MOLECULAR DETECTION OF HUMAN PARASITIC PATHOGENS
MOLECULAR DETECTION OF HUMAN PARASITIC PATHOGENS
EDITED BY DONGYOU LIU
Boca Raton London New York
CRC Press is an imprint of the Taylor & Francis Group, an informa business CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2013 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business
No claim to original U.S. Government works Version Date: 20120608
International Standard Book Number-13: 978-1-4398-1243-3 (eBook - PDF)
This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint.
Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers.
For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged.
Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com This volume is dedicated to a group of international parasitologists whose willingness to share their in-depth knowledge and technical expertise has made an all-inclusive coverage of major human parasitic pathogens possible.
Contents
Preface...... xv Editor ...... xvii Contributors ...... xix
Chapter 1 Introductory Remarks ...... 1 Dongyou Liu
SECTION I Sarcomastigophora
Chapter 2 Acanthamoeba ...... 15 Hélène Yera, Muriel Cornet, and Jean-Louis Bourges
Chapter 3 Balamuthia ...... 25 Govinda S. Visvesvara and Yvonne Qvarnstrom
Chapter 4 Blastocystis ...... 39 Hisao Yoshikawa
Chapter 5 Dientamoeba...... 53 Damien Stark
Chapter 6 Entamoeba ...... 63 Damien Stark and John Ellis
Chapter 7 Giardia ...... 77 Yaoyu Feng and Lihua Xiao
Chapter 8 Leishmania ...... 91 Clotilde Marín, Silvia S. Longoni, and Manuel Sánchez-Moreno
Chapter 9 Naegleria ...... 109 Mineko Shibayama, Jesús Serrano-Luna, Isaac Cervantes-Sandoval, and Víctor Tsutsumi
Chapter 10 Pentatrichomonas ...... 119 Tania Crucitti
Chapter 11 Trichomonas ...... 127 Dongyou Liu, Xiangang Kong, and Changyou Xia
vii viii Contents
Chapter 12 Trypanosoma ...... 135 Emily R. Adams
SECTION II Ciliophora and Apicomplex
Chapter 13 Babesia ...... 151 Dongyou Liu and Xuedong Lu
Chapter 14 Balantidium ...... 161 Dongyou Liu
Chapter 15 Cryptosporidium ...... 167 Una M. Ryan and Rongchang Yang
Chapter 16 Cyclospora ...... 177 Dongyou Liu
Chapter 17 Isospora ...... 187 Chaturong Putaporntip and Somchai Jongwutiwes
Chapter 18 Plasmodium ...... 203 Dongyou Liu and Yi-Wei Tang
Chapter 19 Sarcocystis ...... 215 L. Vangeel, K. Houf, P. Geldhof, J. Vercruysse, R. Ducatelle, and K. Chiers
Chapter 20 Toxoplasma ...... 225 Kjetil Åsbakk and Kristin Wear Prestrud
SECTION III Platyhelminthes: Cestoda
Chapter 21 Diphyllobothrium ...... 237 Barbara Wicht, Raffaele Peduzzi, Hélène Yera, and Jean Dupouy-Camet
Chapter 22 Dipylidium ...... 245 Dongyou Liu
Chapter 23 Echinococcus and Echinococcosis...... 249 Akira Ito, Tetsuya Yanagida, Yasuhito Sako, Minoru Nakao, Kazuhiro Nakaya, Jenny Knapp, and Yuji Ishikawa
Chapter 24 Hymenolepis ...... 265 B.R. Mirdha and Shabeena Rehman Contents ix
Chapter 25 Mesocestoides ...... 277 Kerry A. Padgett, Paul R. Crosbie, and Walter M. Boyce
Chapter 26 Spirometra ...... 287 Dongyou Liu, Jianshun Chen, and Weihuan Fang
Chapter 27 Taenia ...... 297 Munehiro Okamoto and Akira Ito
SECTION IV Platyhelminthes: Trematoda
Chapter 28 Clonorchis ...... 311 Heinz Mehlhorn, Boris Müller, and Jürgen Schmidt
Chapter 29 Dicrocoelium ...... 323 Donato Traversa, Vincenzo Lorusso, and Domenico Otranto
Chapter 30 Echinostomes ...... 333 Rafael Toledo and Bernard Fried
Chapter 31 Fasciola ...... 343 Dongyou Liu and Xing-Quan Zhu
Chapter 32 Fasciolopsis ...... 353 Veena Tandon, B. Roy, and P.K. Prasad
Chapter 33 Gastrodiscoides ...... 365 Dongyou Liu
Chapter 34 Haplorchis ...... 369 Urusa Thaenkham and Jitra Waikagul
Chapter 35 Heterophyidae ...... 379 Ron Dzikowski and Michael G. Levy
Chapter 36 Metagonimus ...... 389 Jae-Ran Yu and Jong-Yil Chai
Chapter 37 Nanophyetus ...... 399 Dongyou Liu
Chapter 38 Opisthorchis ...... 405 Paiboon Sithithaworn, Thewarach Laha, and Ross H. Andrews x Contents
Chapter 39 Paragonimus ...... 423 Hiromu Sugiyama, Takhellambam Shantikumar Singh, and Achariya Rangsiruji
Chapter 40 Plagiorchis ...... 437 Sun Huh
Chapter 41 Schistosoma and Schistosomiasis ...... 441 Antonio Muro and Luís Pérez del Villar
Chapter 42 Trichobilharzia ...... 455 P. Horák, F.M. Schets, L. Kolárˇová, and S.V. Brant
SECTION V Nematoda
Chapter 43 Ancylostoma ...... 469 Jaco J. Verweij
Chapter 44 Angiostrongylus ...... 477 Dongyou Liu
Chapter 45 Anisakis ...... 487 Stefano D’Amelio, Serena Cavallero, Marina Busi, Soa Ingrosso, Elisabetta Giuffra, Sarra Farjallah, and Graça Costa
Chapter 46 Ascaris ...... 501 Dwight D. Bowman and Janice L. Liotta
Chapter 47 Baylisascaris ...... 513 Dongyou Liu
Chapter 48 Brugia ...... 521 Kosum Chansiri and Supatra Areekit
Chapter 49 Capillaria ...... 529 Dongyou Liu
Chapter 50 Dioctophyme ...... 535 Dongyou Liu
Chapter 51 Dirolaria ...... 539 Andrea Gustinelli, Maria Letizia Fioravanti, Monica Caffara, Lorella Garagnani, and Francesco Rivasi
Chapter 52 Dracunculus ...... 549 Dongyou Liu Contents xi
Chapter 53 Enterobius ...... 555 Dongyou Liu
Chapter 54 Gnathostoma ...... 563 Paron Dekumyoy, Tippayarat Yoonuan, and Jitra Waikagul
Chapter 55 Haemonchus ...... 571 Michael B. Hildreth and Aaron Harmon
Chapter 56 Loa ...... 583 Fousseyni S. Touré Ndouo
Chapter 57 Mansonella ...... 595 Ramiro Morales-Hojas
Chapter 58 Marshallagia ...... 603 Jesús M. Pérez and Luca Rossi
Chapter 59 Necator ...... 613 Jan M. Schwenkenbecher
Chapter 60 Oesophagostomum ...... 623 Jaco J. Verweij
Chapter 61 Onchocerca ...... 633 Ramiro Morales-Hojas
Chapter 62 Pseudoterranova and Contracaecum...... 645 Simonetta Mattiucci, Michela Paoletti, Stephen C. Webb, and Giuseppe Nascetti
Chapter 63 Strongyloides ...... 657 Dongyou Liu
Chapter 64 Syphacia ...... 667 Sanford H. Feldman and John H. Porter
Chapter 65 Thelazia ...... 673 Domenico Otranto and Donato Traversa
Chapter 66 Toxocara ...... 681 Anna Borecka
Chapter 67 Trichinella ...... 691 Edoardo Pozio and Giuseppe La Rosa xii Contents
Chapter 68 Trichostrongylus ...... 705 Tai-Soon Yong
Chapter 69 Trichuris ...... 711 C. Cutillas, M. de Rojas, and R. Callejón
Chapter 70 Wuchereria ...... 723 Dongyou Liu and Frank W. Austin
SECTION VI Arthropoda: Arachnida
Chapter 71 Argasidae (Soft Ticks) ...... 733 Dongyou Liu
Chapter 72 Demodex (Hair Follicle Mite) ...... 741 Dongyou Liu
Chapter 73 Dermanyssus, Ornithonyssus, and Trombicula (Red, Fowl, and Harvest Mites) ...... 751 Dongyou Liu
Chapter 74 Ixodidae (Hard Ticks) ...... 761 Dongyou Liu and Chong Yin
Chapter 75 Pyroglyphidae and Glycyphagidae (House Dust and Storage Mites) ...... 777 Dongyou Liu
Chapter 76 Sarcoptes (Itch Mite) ...... 787 Dongyou Liu
SECTION VII Arthropoda: Insecta
Chapter 77 Calliphoridae, Oestridae, and Sarcophagidae (Myiasis-Causing Flies) ...... 799 Dongyou Liu
Chapter 78 Cimex (Bedbug) ...... 811 Dongyou Liu
Chapter 79 Pediculus (Body and Head Lice) ...... 817 Dongyou Liu and Sacha Stelzer-Braid
Chapter 80 Phthirus (Crab Louse) ...... 825 Dongyou Liu Contents xiii
Chapter 81 Pulex, Xenopsylla, and Ctenocephalides (Human, Rat, and Dog/Cat Fleas) ...... 831 Dongyou Liu
Chapter 82 Tunga (Jigger Flea) ...... 837 Dongyou Liu
SECTION VIII Arthropoda: Pentastomida
Chapter 83 Armillifer, Linguatula, and Porocephalus (Tongue Worms) ...... 847 Dongyou Liu
Preface
Parasites are a group of eukaryotic organisms that may be for rapid, sensitive, and specic characterization of parasites. free living or form a symbiotic or parasitic relationship with As a consequence, a large number of original molecular their hosts. Consisting of over 800,000 recognized species, protocols and subsequent modications have been reported. parasites may be unicellular (protozoa) or multicellular (hel- This has created a dilemma to laboratory personnel who are minths and arthropods). In the current six kingdom systems not directly involved in the development of original or modi- for biological organisms (i.e., Bacteria, Protozoa, Fungi, ed protocols to know which are most appropriate to adopt Chromista, Plantae, and Animalia), protozoa constitute an for accurate identication of parasites of interest. independent kingdom, while helminths and arthropods make The purpose of the current book is to address this issue, up parts of the kingdom Animalia. with international scientists in respective parasite research The association of parasites with human populations and diagnosis providing expert summaries on current diag- had occurred long before the emergence of civilization. nostic approaches for major human parasitic pathogens. The identication of hookworm ova and Paragonimus ova Each chapter consists of a brief review on the classication, in human coprolites dated c. 5000 BC and c. 2500 BC in epidemiology, clinical features, and diagnosis of an impor- Brazil and Chile, respectively, reinforces the longstanding tant parasite genus or group; an outline of clinical sample relationship between parasitic pathogens and human races. collection and preparation procedures; a selection of repre- In spite of our unrelenting efforts against parasitic diseases, sentative stepwise molecular protocols; and a discussion on parasites continue to cause signicant human morbidity and further research requirements relating to improved diagno- mortality. For example, Plasmodium, the malaria parasite, sis. This book constitutes a reliable and convenient reference inicts >250 million people worldwide and causes >1 mil- on molecular detection and identication of major human lion deaths per annum; soil-transmitted helminths such as parasitic pathogens; an indispensable tool for upcoming and roundworm (Ascaris), whipworm (Trichuris), and hookworm experienced medical, veterinary, and industrial laboratory (Ancylostoma and Necator) affect >2 billion people glob- scientists engaged in parasite characterization; and an essen- ally, resulting in >200,000 yearly deaths. Other important tial textbook for undergraduate and graduate students major- human parasitic pathogens include Schistosoma, lariae, ing in parasitology. Trypanosoma cruzi, and Leishmania, which together cause A comprehensive book such as this is clearly beyond an miseries to hundreds of millions of people worldwide. individual’s capacity. I am fortunate and honored to have a In view of their diverse range and variety, it is critical large panel of international parasitologists as chapter con- that parasites are correctly identied so that effective treat- tributors. Their detailed knowledge and technical insights ment and control of parasitic diseases can be implemented. have greatly enriched this book. Additionally, the profession- Traditionally, laboratory identication of parasites has relied alism and dedication of executive editor Barbara Norwitz upon various phenotypic procedures that detect their mor- and senior project coordinator Jill Jurgensen at CRC Press phological, biological, and immunological features. Because have enhanced its presentation. Finally, the compilation of these procedures tend to be time consuming and require this all-encompassing volume would have been impossible specialized skills, molecular methods based on nucleic acid without the understanding and support of my family, Liling amplication technologies have been increasingly utilized Ma, Brenda, and Cathy.
xv
Editor
Dongyou Liu, PhD, undertook his veterinary science dermatophyte fungi (Trichophyton, Microsporum, and education at Hunan Agricultural University, China, and Epidermophyton), and listeriae (Listeria species). pursued his postgraduate training in parasitology at the He is the rst author of over 50 original research and review University of Melbourne, Australia, where he worked articles in various international journals and the editor of sev- toward improved immunological diagnosis of human hyda- eral recent biomedical books published by Taylor & Francis tid disease. CRC Press: Handbook of Listeria monocytogenes (2008), Over the last two decades, he has crisscrossed between Handbook of Nucleic Acid Purication (2009), Molecular research and clinical laboratories in Australia and the United Detection of Foodborne Pathogens (2009), Molecular States of America, with focuses on molecular characteriza- Detection of Human Viral Pathogens (2010), Molecular tion and virulence determination of microbial pathogens Detection of Human Bacterial Pathogens (2011), and such as ovine footrot bacterium (Dichelobacter nodosus), Molecular Detection of Human Fungal Pathogens (2011).
xvii
Contributors
Emily R. Adams Marina Busi Koninklijk Instituut voor de Tropen Biomedical Research Department of Public Health Science Amsterdam, the Netherlands Sapienza University of Rome Rome, Italy Ross H. Andrews Department of Parasitology and Liver Fluke and Monica Caffara Cholagiocarcinoma Research Center Department of Veterinary Public Health and Animal Khon Kaen University Pathology Khon Kaen, Thailand University of Bologna Bologna, Italy Supatra Areekit R. Callejón Department of Biochemistry Department of Microbiology and Parasitology Srinakharinwirot University University of Seville Bangkok, Thailand Seville, Spain
Kjetil Åsbakk Serena Cavallero Norwegian School of Veterinary Science Department of Public Health Science Section of Arctic Veterinary Medicine Sapienza University of Rome Tromsø, Norway Rome, Italy
Frank W. Austin Isaac Cervantes-Sandoval Department of Pathobiology and Department of Infectomic and Molecular Population Medicine Pathogenesis Mississippi State University National Polytechnic Institute Mississippi State, Mississippi Mexico City, Mexico
Anna Borecka Jong-Yil Chai Institute of Parasitology Department of Parasitology and Tropical Medicine Polish Academy of Sciences Seoul National University College of Medicine Warsaw, Poland and
Jean-Louis Bourges Institute of Endemic Disease Department of Ophthalmology Seoul National University Medical Research Université Paris Center Paris, France Seoul, Korea Kosum Chansiri Dwight D. Bowman Department of Biochemistry College of Veterinary Medicine Srinakharinwirot University Cornell University Bangkok, Thailand Ithaca, New York Jianshun Chen Walter M. Boyce Zhejiang Fisheries Technical Extension Center and Department of Pathology, Microbiology, and Zhejiang Aquatic Disease Prevention and Immunology Quarantine Center University of California, Davis Zhejiang, China Davis, California K. Chiers S.V. Brant Department of Pathology, Bacteriology, and Avian Department of Biology Diseases University of New Mexico Ghent University Albuquerque, New Mexico Merelbeke, Belgium
xix xx Contributors
Muriel Cornet Ron Dzikowski Department of Microbiology Department of Microbiology and Molecular Genetics Université Paris The Hebrew University–Hadassah Medical School Paris, France Jerusalem, Israel
Graça Costa John Ellis Centre for Macaronesian Studies School of Medical and Molecular Biosciences University of Madeira University of Technology Funchal, Portugal Sydney, New South Wales, Australia
Paul R. Crosbie Weihuan Fang Department of Biology Institute of Preventive Veterinary Medicine California State University Zhejiang University Fresno, California Zhejiang, China
Tania Crucitti Sarra Farjallah STD/HIV Research and Intervention Unit Institute of Applied Sciences and Technology of Gabes Institute of Tropical Medicine Rue Amor Ben El Khatab Antwerp, Belgium Gabes, Tunisia
C. Cutillas Sanford H. Feldman Department of Microbiology and Center for Comparative Medicine Parasitology University of Virginia University of Seville Charlottesville, Virginia Seville, Spain Yaoyu Feng School of Resource and Environmental Engineering Stefano D’Amelio East China University of Science and Technology Department of Public Health Science Shanghai, China Sapienza University of Rome Rome, Italy Maria Letizia Fioravanti Department of Veterinary Public Health and Animal Paron Dekumyoy Pathology Department of Helminthology University of Bologna Mahidol University Bologna, Italy Bangkok, Thailand Bernard Fried Luís Pérez del Villar Department of Biology Tropical Medicine Research Center Lafayette College University of Salamanca Easton, Pennsylvania Salamanca, Spain Lorella Garagnani M. de Rojas Dipartimento ad Attività Integrata di Laboratori Department of Microbiology and Parasitology Anatomia Patologica e Medicina Legale University of Seville Azienda Ospedaliero–Universitaria di Modena Seville, Spain Modena, Italy
R. Ducatelle P. Geldhof Department of Pathology, Bacteriology, and Avian Department of Virology, Parasitology, and Diseases Immunology Ghent University Ghent University Merelbeke, Belgium Merelbeke, Belgium
Jean Dupouy-Camet Elisabetta Giuffra Laboratoire de Parasitologie-Mycologie Parco Tecnologico Padano Université Paris Descartes Polo Universitario Paris, France Lodi, Italy Contributors xxi
Andrea Gustinelli L. Kolár¡ová Department of Veterinary Public Health and Institute of Immunology and Microbiology Animal Pathology Charles University in Prague University of Bologna Prague, Czech Republic Bologna, Italy Xiangang Kong Aaron Harmon Harbin Veterinary Research Institute Department of Biology and Microbiology Chinese Academy Agricultural Sciences South Dakota State University Heilongjiang, China Brookings, South Dakota Thewarach Laha Michael B. Hildreth Department of Parasitology and Liver Fluke and Department of Biology and Cholangiocarcinoma Research Center Microbiology Khon Kaen University South Dakota State University Khon Kaen, Thailand Brookings, South Dakota Giuseppe La Rosa P. Horák Department of Infectious, Parasitic, and Immunomediated Department of Parasitology Diseases Charles University in Prague Istituto Superiore di Sanità Prague, Czech Republic Rome, Italy
K. Houf Michael G. Levy Department of Veterinary Public Health and Department of Population Health and Pathobiology Food Safety North Carolina State University Ghent University Raleigh, North Carolina Merelbeke, Belgium Janice L. Liotta Sun Huh Department of Microbiology and Immunology Department of Parasitology Cornell University Hallym University Ithaca, New York Chuncheon, Korea
Soa Ingrosso Dongyou Liu Department of Public Health Science Royal College of Pathologists of Australasia Quality Sapienza University of Rome Assurance Programs Rome, Italy Surry Hills, New South Wales, Australia
Yuji Ishikawa Silvia S. Longoni Department of Parasitology Department of Parasitology Asahikawa Medical College University of Granada Hokkaido, Japan Granada, Spain
Akira Ito Vincenzo Lorusso Department of Parasitology Department of Veterinary Public Health Asahikawa Medical College University of Bari Hokkaido, Japan Bari, Italy
Somchai Jongwutiwes Xuedong Lu Department of Parasitology Futian Hospital Chulalongkorn University Guandong Medical College Bangkok, Thailand Guangdong, China
Jenny Knapp Clotilde Marín Department of Parasitology Department of Parasitology Asahikawa Medical College University of Granada Hokkaido, Japan Granada, Spain xxii Contributors
Simonetta Mattiucci Domenico Otranto Department of Public Health Sciences Department of Veterinary Public Health Sapienza University of Rome University of Bari Rome, Italy Bari, Italy
Heinz Mehlhorn Kerry A. Padgett Department of Parasitology California Department of Public Health Heinrich Heine University Vector Borne Disease Section North Rhine–Westphalia, Germany Richmond, California
B.R. Mirdha Michela Paoletti Department of Microbiology Department of Public Health Sciences All India Institute of Medical Sciences Sapienza University of Rome New Delhi, India Rome, Italy
Ramiro Morales-Hojas Raffaele Peduzzi Instituto de Biologia Molecular e Celular Istituto cantonale di microbiologia Universidade do Porto Bellinzona, Switzerland Porto, Portugal Jesús M. Pérez Boris Müller Departamento de Biología Animal, Biología Vegetal y Department of Parasitology Ecología Heinrich Heine University Universidad de Jaén North Rhine–Westphalia, Germany Jaén, Spain
Antonio Muro John H. Porter Tropical Medicine Research Center Department of Environmental Sciences University of Salamanca University of Virginia Salamanca, Spain Charlottesville, Virginia
Minoru Nakao Edoardo Pozio Department of Parasitology Department of Infectious, Parasitic, and Immunomediated Asahikawa Medical College Diseases Hokkaido, Japan Instituto Superiore di Sanità Rome, Italy Kazuhiro Nakaya Department of Parasitology P.K. Prasad Asahikawa Medical College Department of Zoology Hokkaido, Japan North-Eastern Hill University Meghalaya, India Giuseppe Nascetti Department of Ecology and Sustainable Economic Kristin Wear Prestrud Development Norwegian School of Veterinary Science Tuscia University Section of Arctic Veterinary Medicine Viterbo, Italy Tromsø, Norway
Fousseyni S. Touré Ndouo Chaturong Putaporntip Medical Parasitology Unit Department of Parasitology International Centre for Medical Research of Franceville Chulalongkorn University Franceville, Gabon Bangkok, Thailand
Munehiro Okamoto Yvonne Qvarnstrom Primate Research Institute Department of Health and Human Services Kyoto University Centers for Disease Control and Prevention Kyoto, Japan Atlanta, Georgia Contributors xxiii
Achariya Rangsiruji Jesús Serrano-Luna Department of Biology Department of Cell Biology Srinakharinwirot University National Polytechnic Institute Bangkok, Thailand Mexico City, Mexico
Shabeena Rehman Takhellambam Shantikumar Singh Department of Biosciences Department of Microbiology Jamia Millia Islamia Sikkim Manipal Institute of Medical Sciences New Delhi, India Sikkim, India
Francesco Rivasi Mineko Shibayama Dipartimento ad Attività Integrata di Laboratori Anatomia Department of Infectomic and Molecular Patologica e Medicina Legale Pathogenesis Azienda Ospedaliero–Universitaria di Modena National Polytechnic Institute Modena, Italy Mexico City, Mexico
Luca Rossi Paiboon Sithithaworn Dipartimento di Produzione Animali, Epidemiologia ed Department of Parasitology and Liver Fluke and Ecologia Cholangiocarcinoma Research Center Università degli Studi di Torino Khon Kaen University Torino, Italy Khon Kaen, Thailand
B. Roy Damien Stark Department of Zoology Division of Microbiology North-Eastern Hill University SydPath St. Vincent’s Hospital Meghalaya, India Darlinghurst, New South Wales, Australia
Una M. Ryan Sacha Stelzer-Braid School of Veterinary and Biomedical Science Virology Research Laboratory Murdoch University Prince of Wales Hospital Murdoch, Western Australia, Australia Randwick, New South Wales, Australia
Yasuhito Sako Hiromu Sugiyama Department of Parasitology Department of Parasitology Asahikawa Medical College National Institute of Infectious Diseases Hokkaido, Japan Tokyo, Japan
Manuel Sánchez-Moreno Veena Tandon Department of Parasitology Department of Zoology University of Granada North-Eastern Hill University Granada, Spain Meghalaya, India
F.M. Schets Yi-Wei Tang National Institute for Public Health and Clinical Microbiology Service Environment Centre for Infectious Disease Control Memorial Sloan-Kettering Cancer Center Netherlands New York, New York Bilthoven, the Netherlands Urusa Thaenkham Jürgen Schmidt Department of Helminthology Department of Parasitology Mahidol University Heinrich Heine University Bangkok, Thailand North Rhine–Westphalia, Germany Rafael Toledo Jan M. Schwenkenbecher Departamento de Biología Celular y Institute of Biological and Environmental Sciences Parasitología University of Aberdeen Universidad de Valencia Aberdeen, Scotland, United Kingdom Burjassot-Valencia, Spain xxiv Contributors
Donato Traversa Lihua Xiao Department of Comparative Biomedical Sciences Division of Foodborne and Waterborne Disease University of Teramo Prevention Teramo, Italy Centers for Disease Control and Prevention Atlanta, Georgia Víctor Tsutsumi Department of Infectomic and Molecular Pathogenesis Tetsuya Yanagida National Polytechnic Institute Department of Parasitology Mexico City, Mexico Asahikawa Medical College Hokkaido, Japan L. Vangeel Department of Pathology, Bacteriology, and Avian Rongchang Yang Diseases School of Veterinary and Biomedical Science Ghent University Murdoch University Merelbeke, Belgium Murdoch, Western Australia, Australia
J. Vercruysse Hélène Yera Department of Virology, Parasitology, and Immunology Laboratory of Parasitology–Mycology Ghent University Université Paris Descartes Merelbeke, Belgium Paris, France
Jaco J. Verweij Chong Yin Department of Parasitology College of Veterinary Medicine Leiden University Medical Center Hunan Agricultural University Leiden, the Netherlands Hunan, China
Govinda S. Visvesvara Tai-Soon Yong Department of Health and Human Services Department of Environmental Medical Biology Centers for Disease Control and Prevention, Public Yonsei University College of Medicine Health Service Seoul, Korea Atlanta, Georgia Tippayarat Yoonuan Jitra Waikagul Department of Helminthology Department of Helminthology Mahidol University Mahidol University Bangkok, Thailand Bangkok, Thailand Hisao Yoshikawa Stephen C. Webb Department of Biological Sciences Cawthron Institute Nara Women’s University Nelson, New Zealand Nara, Japan
Barbara Wicht Jae-Ran Yu Istituto cantonale di microbiologia Konkuk University School of Medicine Bellinzona, Switzerland Seoul, Korea
Changyou Xia Xing-Quan Zhu Harbin Veterinary Research Institute Lanzhou Veterinary Research Institute Chinese Academy of Agricultural Sciences Chinese Academy of Agricultural Sciences Heilongjiang, China Gansu, China 1 Introductory Remarks Dongyou Liu
CONTENTS 1.1 Preamble ...... 1 1.2 Classication, Morphology, and Biology ...... 2 1.2.1 Protozoa ...... 2 1.2.2 Helminths ...... 2 1.2.3 Arthropods ...... 5 1.3 Phenotypic Characterization...... 6 1.3.1 Macroscopic and Microscopic Assessment ...... 6 1.3.2 Biochemical and Immunological Tests ...... 7 1.3.3 In Vitro and In Vivo Cultivation ...... 7 1.4 Genotypic Characterization ...... 7 1.4.1 Target Genes ...... 7 1.4.2 Template Amplication and Product Detection ...... 8 1.4.2.1 Template Amplication...... 8 1.4.2.2 Product Detection ...... 9 1.4.3 Data Analysis ...... 9 1.5 Results Interpretation, Standardization, and Quality Control and Assurance ...... 9 1.5.1 Key Performance Characteristics ...... 9 1.5.2 Results Interpretation ...... 10 1.5.3 Standardization and Validation ...... 10 1.5.4 Quality Control and Assurance ...... 11 1.5.4.1 Quality Control ...... 11 1.5.4.2 Quality Assurance ...... 11 1.6 Conclusion ...... 11 References ...... 11
1.1 PREAMBLE Based on their interactions with their hosts, parasites are often referred to as endoparasites (which live inside the host, Parasites (from Greek parasitos, para—beside, by; sitos— e.g., parasitic worms) and ectoparasites (which live on the wheat; meaning one who eats at the table of another) are surface of the host, e.g., mites). Endoparasites may inhabit eukaryotic organisms that may be free living, or form sym- spaces in the host’s body (intercellular form) or cells in the biotic or parasitic relationship with their hosts. Eukaryotes host’s body (intracellular form). Intracellular parasites often are characterized by having a well-dened chromosome rely on a third organism (generally known as carrier or vec- in a nuclear membrane (which is packed with histones tor) for transmission to the host. In addition, some parasites to form linear chromatin), possessing various independent spend their entire life on the host (e.g., Enterobius), while membrane-bounded cytoplasmic organelles, and undergo- others may undergo freeliving and parasitic stages in their ing endocystosis and exocytosis. This is in clear contrast life cycle (e.g., Strongyloides). with prokaryotes (e.g., bacteria) that lack nuclear mem- The study of parasitic protozoa, helminths, and insects, brane, contain few independent membrane-bounded cyto- their hosts, and the host–parasite relationship is commonly plasmic organelles, and do not undergo endocytosis and known as parasitology. The discipline of parasitology exam- exocytosis. ines the morphology, life cycle, pathogenesis and identi- Consisting of over 800,000 identied species, parasites cation of parasites, and draws on techniques from elds come under the following three categories: protozoa, hel- such as cell biology, ecology, biochemistry, immunology, minths, and arthropods. In the current six kingdom systems molecular biology, bioinformatics, genetics, and evolution. for biological organisms (i.e., Bacteria, Protozoa, Fungi, Although fungi represent another category of eukaryotes Chromista, Plantae, and Animalia), protozoa constitute an with the potential to lead a parasitic life in plant, animal, independent kingdom of their own, while helminths and and human hosts, they are discussed in a distinct branch arthropods constitute parts of the kingdom Animalia [1,2]. of science (mycology). The kingdom Chromista comprises
1 2 Molecular Detection of Human Parasitic Pathogens unicellular plant-like organisms, to which a previous pro- Structurally, many protozoa possess a pellicle that is tozoan taxon Blastocystis belongs. composed of an outer plasma membrane and an inner cyto- skeleton, and that plays a pivotal role in the maintenance of 1.2 CLASSIFICATION, MORPHOLOGY, cell shape. The plasma membrane further separates into a thin outer ectoplasm and an inner endoplasm; the underly- AND BIOLOGY ing cytoskeleton may include additional membranes, micro- Parasites infecting humans commonly belong to three cat- tubules, microlaments, or plates of cellulose or protein. egories: protozoa, helminths, and arthropods. Within the cytoplasm exist nucleus, nuclear membrane, chro- mosomes, endoplasmic reticulum, mitochondria, golgi body, ribosomes, and various specialized structures (e.g., vacuoles 1.2.1 PROTOZOA that may be stomach-like for food digestion, or contractible Protozoa (singular protozoon) are small, unicellular, phago- for eliminating excess water). trophic, nonphotosynthetic, eukaryotic organisms without Protozoa have the ability to reproduce sexually and asex- cell walls. Previously, protozoa (meaning “rst animals,” ually, with each individual protozoan being both male and with Protista and Protoctista, being alternative names) have female. Cell division in protozoa usually involves a closed been classied in the kingdom Animalia, and more recently spindle (constructed inside the intact nuclear envelope), in form an independent, basal eukaryotic kingdom within the contrast to the open spindle of multicellular animals. In par- six kingdom system (Bacteria, Protozoa, Chromista, Plantae, ticular, parasitic protozoa tend to have short generation times Fungi, and Animalia). As protozoa can be of both purely and high rates of reproduction. auto- and heterotrophic, they can t into a plant [auto (photo) Protozoa are both diverse and abundant. With >200,000 trophic], fungal (heterotrophic), or animal (heterotrophic) named species, and present in soil, freshwater, and marine context. Additionally, some protozoa (e.g., dinoagellates) environments, protozoa make one of the largest biomasses demonstrate an intermediate trophic capability (thus called on earth. As heterotrophic organisms, protozoa prey on uni- mixotrophic). Other previous protozoan taxa such as micro- cellular or lamentous algae, bacteria and microfungi, and sporidia and pneumocystis have been transferred to the king- also form an important food source for microinvertebrates, dom Fungi (which covers eukaryotic heterotrophic organisms contributing to the transfer of bacterial and algal production lacking plastids but possessing cell walls containing chitin to successive trophic levels. Moreover, protozoa may be sym- and glycans). Moreover, there is another previous protozoan biotic or parasitic, living attached to or inside other organ- taxon Blastocystis that has been redesignated in the kingdom isms. In fact, of the >200,000 named species, nearly 10,000 Chromista (covering unicellular, photosynthetic, lamen- protozoa are parasitic in invertebrates and in almost every tous, or colonial organisms (“algae”); some with secondary species of vertebrate. loss of plastids). Being the only parasitic member of the king- Protozoa may enter human hosts via the nasal passage dom Chromista, Blastocystis will be covered in Section I, (e.g., Naegleria fowleri), inoculation (e.g., malaria transmit- Sarcomastigophora of the book for convenience in spite of its ted by mosquitoes), ingestion of cysts (e.g., amoebic dysen- changed taxonomical classication. tery), fecal contamination of wounds (e.g., Chagas disease Measuring from 10 to 200 µm in size (with the parasitic transmitted by assassination bugs), intrauterine infection forms tending toward the lower end of this range), protozoa (e.g., malaria), kissing (e.g., Trichomonas tenax), or sexual demonstrate considerable variations in their locomotory intercourse (Trichomonas vaginalis). Examples of signicant structures (pseudopodia, agella, cilia, or no organelle of protozoan diseases in humans include malaria (Plasmodium), locomotion), which have underlined the traditional classi- sleeping sickness (African trypanosomes), Chagas disease cation system for parasitic organisms within this group (i.e., (Trypanasoma cruzi), and leishmaniasis (Leishamania). phyla Sarcomastigophora, Apicomplexa, and Ciliophora) (Table 1.1). 1.2.2 HELMINThS The phylum Sarcomastigophora is further separated into subphyla Mastigophora and Sarcodina. While the subphy- The term helminths (singular helminth) commonly refers to lum Mastigophora (agellates) possesses one or more a- organisms belonging to the phyla Platyhelminthes (platyhel- gella as locomotive organ and reproduces by binary ssion minths or atworms) and Nematoda (nematodes or round- as well as sexual reproduction in some species, the subphy- worms) (Table 1.1). lum Sarcodina (amoebae) moves by pseudopodia (although The phylum Platyhelminthes (platyhelminths or at- agella may be present in the reproductive stages) and worms) is divided into four classes: Cestoda (tapeworms), reproduces by binary ssion. The phylum Apicomplexa (spo- Trematoda (ukes), Monogenea, and Turbellaria. The class rozoans) has no special locomotive organs, reproduces by Cestoda (tapeworms) is further separated into two sub- multiple ssion and gamogony and forms spores at one stage classes, the Cestodaria (which lacks a scolex, is monozoic of their life. The phylum Ciliophora (ciliates) moves by cilia, with a set of reproductive organ, and parasitic in shes and has two nuclei (one macronucleus and one micronucleus), and turtles) and the Eucestoda (the majority of which are seg- reproduces by transverse binary ssion, conjugation, autog- mented and polyzoic, and are parasitic to humans). The class amy, and cytogamy. Trematoda (ukes) comprises two subgroups, the Digenea Introductory Remarks 3
TABLE 1.1 Classification and Characteristics of Major Human Parasitic Pathogens Subphylum/ Notable Human Kingdom Phylum Class Brief Description Pathogens Protozoa Sarcomastigophora Sarcodina Obligate and facultative amoebae; move by pseudopodia; Acantamoeba (amoebae) cytoplasmic streaming assists movement. Asexual reproduction Balamuthia occurs by ssion of the cell Entamoeba Naegleria Mastigophora Intestinal, tissue, and blood-dwelling agellates; move by agella; Dientamoeba (agellates) reproduction in these protozoa generally occurs by ssion, Giardia although sexual reproduction is observed in some species Leishmania Trichomonas Trypanosoma Apicomplexa Sporozoa; unicellular organisms possessing at some stage an apical Babesia (sporozoans) complex composed of polar rings, rhoptries, micronemes, and Cryptosporidium typically a conoid; elaborate life cycles involving a sexual process; Isospora all parasites; no organelle of locomotion; forming spores at one Plasmodium stage in their life cycle Sarcocystis Toxoplasma Ciliophora (ciliates) Unicellular organisms possessing many cilia for locomotion and Balantidium complex oral ciliature for feeding; having somatic, polyploid macronuclei and generative, diploid micronuclei; conjugation may be used for sexual reproduction, and binary ssion also occurs Animalia Nematoda Adenophorea Round worms; appear round in cross section; have body cavities, a Ascaris Secrenentea straight alimentary canal and an anus; have separate sexes Ancylostoma (dioecious reproductive system) Enterobius Necator Strongyloides Trichuris Platyhelminthes Cestoda Tape worms; have a head (scolex) with sucking organs, a segmented Diphyllobothrium body, but no alimentary canal; each body segment is Echinococcus hermaphrodite; adult tapeworms live in the intestine of the Taenia denitive host Trematoda Flukes; a nonsegmented, usually leaf-shaped body, with two Fasciolopsis suckers but no distinct head; have an alimentary canal (but no Opisthorchis anus) and are usually hermaphrodite; however, Schistosomes are Schistosoma thread-like, and have separate sexes Animalia Arthropoda Arachnida Two segments (cephalothorax and abdomen); eight legs; one pair Argas of chelicerae; no attennae Ixodes Consisting of orders Astigmata (mites) and Metastigmata (ticks) Sarcoptes Insecta Three body segments (head, thorax, and abdomen); six legs; one Pediculus pair of attennae Phthirus Consisting of medically relevant orders Anoplura (lice), Diptera Tunga (ies), and Siphunculata (eas) Pentastomida Tongue worms Armillifer Linguatula
and the Aspidogastrea, with only the former occurring in and the gut). Many are hermaphroditic (with both male and humans and other mammals. The class Monogenea cov- female sex organs), undergo self-fertilization, and have an ers mainly ectoparasites of shes; and the class Turbellaria indirect lifecycle. includes a variety of free-living organisms in aquatic and ter- The adult tapeworms in the subclass Eucestoda have an restrial environments. elongate, segmented, at body, which is covered by a syn- The platyhelminths or atworms in the phylum Platyhel cytial outer layer formed by the tegumental cells (known minthes have a bilaterally symmetrical, dorsoventrally at- as tegument). The anterior end of the body contains a tened body with three well-dened germ layers (ectoderm, holdfast organ called a scolex, which may have a rostel- endoderm, and mesoderm), a denite head end, but without lum, suckers, bothria, bothridia, tentacles, hooks, and/or a coelom (a uid-lled cavity between the outer body wall spines for attachment to the gut of the host. The scolex is 4 Molecular Detection of Human Parasitic Pathogens followed by a short unsegmented portion (called the neck) Most digeneans have a forked digestive system (consisting and the remainder of the body (or strobila) is composed of of oral sucker, the pharynx, esophagus, and blind-end- a number of segments (or proglottids). Each segment (pro- ing caeca) that opens at the mouth as an anus is absent; glottid) harbors one or more sets of the male and female and a single set of reproductive organs. The tegument of reproductive organs. There is a lack of body cavity and digeneans is syncitial (which consists of a mass of proto- alimentary canal. plasm containing many nuclei but not divided into cells) Of the two orders within the subclass Eucestoda, the order and is involved in the direct nutrient uptake. The digenetic Pseudophyllidea is characterized by its lack of suckers and trematodes are normally hermaphroditic (with both male hooks in the scolex, and its use of a pair of bothria (dorsal and female reproductive systems), occasionally partly or and ventral longitudinal grooves on the scolex) for attach- entirely dioecious (e.g., schistosomes have separate sexes). ment; and the order Cyclophyllidea is characterized by pres- The female reproductive system comprises an ovary, ovi- ence of a ring of four suckers and sometimes one or more duct, sometimes a seminal receptacle (for storing sperm rings of apical hooks on the scolex. The presence or absence from the copulatory partner), ootype (where the egg is of scolex, the shape of the segments (proglottids) and the formed) surrounded by Mehlis’ gland, yolk gland (vitellar- arrangement and form of the reproductive system(s) within ium) which opens into the ootype or near it, and the uterus the segments (e.g., the position of the genital pore, the size (in which eggs mature), which sometimes forms a muscu- of the cirrus sac, the shape of the ovary, uterus, and vitel- lar terminal called metraterm, and which opens through a larium or yolk gland) are useful features for differentiation common gonopore together with the male system. A nar- of cestodes. The medically important members in the order row duct, Laurer’s canal, links the female system near the Pseudophyllidea are found in the family Diphyllobothriidae, ootype to the dorsal surface. The male reproductive system while those in the order Cyclophyllidea are found in the comprises of one testis or several testes, and sperm ducts families Anoplocephalidae, Davaineidae, Dipylidiidae, which unite and widen terminally to form a seminal vesicle Hymenolepididae, Mesocestoididae, and Taeniidae. in a cirrus pouch or sac. During their lifecycle, cestodes generally involve a nal The lifecycle of digenetic trematodes includes an adult host, and one or two intermediate hosts; the only cestode spe- stage in the nal host, and ve larval stages (miracidium, cies capable of autoinfection without the use of intermediate sporocyst, redia, cercaria, and metacercaria) in one, two, or hosts is Hymenolepis. Adult cestodes live in the intestine of more intermediate hosts. The eggs of the digenea are oval vertebrates, and the eggs or gravid segments containing eggs and operculate and are normally passed in the faces of the are discharged with the feces. The egg of the cyclophyllid- nal host. Under suitable conditions of moisture, warmth, eans is eaten by a terrestrial invertebrate (e.g., an arthropod) light, and salinity, the egg hatches to release a ciliated larva, or a vertebrate, in which it hatches and releases a hexacanth miracidium. The miracidium enters a rst intermediate host (six-hooked) larva (called onchosphere). The egg of the (a snail), and forms a sporocyst after losing ciliated coat. pseudophyllideans hatches in water and releases a ciliated, Within the sporocyst the germinal cells multiply and pro- motile hexacanth larva (called coracidium), which is eaten by duce either daughter sporocysts (which lacks a pharynx) or an aquatic arthropod (e.g., a copepod). The hexacanth pen- rediae (which has a pharynx or a sac-like gut), and the spo- etrates the gut wall of intermediate host and develops into rocyst or the redia then produces the cercaria. After leaving a procercoid in the body cavity. The procercoid may further the snail host through an opening, or the tegument or being develop, either in the same host or in a second intermediate expelled, the cercaria encysts on or in a second intermedi- host if the rst host is eaten, into a resting, encysted, stage ate host (typically an invertebrate) or on vegetation, and the that is referred to as cysticercus, cysticercoid, plerocercoid, encysted cercaria matures to produce the infective stage, tetrathyridium, or hydatid. The nal host acquires the worm the metacercaria. The metacercaria contained in the second by feeding on the intermediate host harboring the encysted intermediate host is ingested by a denitive host and devel- stage. In rare instances, asexual multiplication of larval ops into the adult worm in the gut, liver, bile ducts, lungs, heads (protoscoleces) can take place in vertebrates which act or blood system. However, the metacercaria is absent in the as an intermediate host. family Schistosomatidae and its cercaria actively penetrates The presence of adult tapeworm in humans may result in the skin of the denitive host. diarrhea, loss of appetite, and illthrift. On the other hand, the The medically important trematodes are found in the orders presence of encysted larval tapeworms in the tissues (e.g., the Strigeida (family Schistosomatidae), Echinostomida (fami- brain, liver, and lungs) may have serious consequences. For lies Echinostomatidae and Fasciolidae), and Plagiorchiida example, about 50,000 people die annually of cysticercosis, (families Dicrocoeliidea, Heterophyidae, Opsithorchiidae, an infection with Taenia solium larvae. Paragonimidae, and Plagiorchiidae). Features useful for dis- The adult worms of the digenetic trematodes in the class tinguishing trematode groups include the presence or absence Trematoda (ukes) are usually dorsoventrally attened, of spines on the tegument, the position and number of suck- with some having long and narrow, leaf-shaped, or thick ers, the form and arrangement of the reproductive organs, eshy bodies. Adult trematodes typically have an anterior and the shape of the gut and excretory vesicle. For species oral sucker surrounding the mouth, and a ventral sucker level discrimination, details such as sucker ratio and egg size (sometimes termed the acetabulum) on the ventral surface. are informative. Introductory Remarks 5
Besides taking in nutrients through their body surface, host, nematode larvae migrate through the body and develop digeneans also feed by browsing. The feeding process and in adult worms after the third and fourth molts (ecdyses) in the presence of spines on the trematode body can cause dam- their normal habitat. age and irritation. The toxins produced by digeneans can Heavily armed with teeth or other sclerotized mouthparts, elicit inammation, tissue reactions, brosis, obstruction, nematodes such as hookworms browse on the gut wall, caus- and so on. ing considerable damage. Migrating adults in the tissues and The phylum Nematoda is divided into two classes, larvae (termed as larva migrants) can also cause serious prob- Adenophorea and Secernentea, which reect the presence lems in sensitive regions such as the brain, liver, or eyes. The and absence of small sensory structures (phasmids) on the medically important families in the class Adenophorea include tail and the nature of the excretory system. In addition, in Trichinellidae and Trichuridae; those in the class Secern the Adenophora the rst-stage larva is infective to the deni- entea include Ancylostomatidae, Anisakidae, Ascarididae, tive (nal) host, whereas in the Secernentea it is the third- Dracunculidae, Onchocercidae, Gnathostomatidae, Angio stage larva. A majority of animal parasites are found in the strongylidae, Oxyuridae, Strongyloididae, Thelaziidae, and Secernentea. Trichostrongylidae. The phylum Nematoda is characterized by (i) a bilateral Apart from phyla Platyhelmithes and Nematoda, there symmetrical and triploblastic (or triblastic) body; (ii) a body is another phylum of parasites Acanthocephalans (“thorny- wall consisting of an outer layer of cuticle and an inner layer headed worms”) that commonly occur in animals, and rarely of longitudinal muscles with scattered nuclei but without cell in humans (causing intestinal perforation and peritonitis). As boundaries; (iii) a pseudocoel body cavity between the body dioecious worms with a body cavity, Acanthocephalans have wall and the digestive tract, that is not lined by the mesoder- lost their digestive system and reduced their muscular, ner- mal (or coelomic) epithelium; (iv) a complete digestive sys- vous, and excretory systems. The body of Acanthocephalans tem with a mouth at the anterior end, a muscular esophagus consists of a large trunk and a retractable anterior proboscis and an intestine leading to anus; (v) a lack of the circulatory armed with a regular array of hooks used for attachment to and respiratory system; (vi) a circumventric nerve ring with the gut wall. The number of rows of hooks and the number of longitudinal nerves in the nervous system; (vii) a pair of lon- hooks per row in the proboscis are of diagnostic importance. gitudinal excretory canals with excretory pores in the excre- The body wall is a syncytium with large, fragmented nuclei, tory system; (viii) separate sexes, with males and females along with a complex outer tegument and an inner layer of differing in size and certain morphological features (sexual muscles, as well a lacunar system of interconnecting canals dimorphism); (ix) eggs with plug at either end. The nature of that may serve as a circulatory system. The males harbor the anterior regions of the alimentary canal (e.g., the esopha- two testes and a series of accessory glands (called cement gus), the form of the head (presence, number of lips, teeth, glands). The females have ovarian balls, and the body cavity etc.), the form of the male tail such as the arrangement of cau- of mature females is full of eggs. dal papillae (sensory structures used during copulation), and During the lifecycle of Acanthocephalans, female adults the length and shape of the spicule(s) (sclerotized copulatory in the intestine produce eggs that are voided with the feces. aids) are important features for nematode identicaiton. In After ingestion by a suitable arthropod, the egg releases a the majority of cases, males carry more taxonomically use- larva armed with hooks (called an acanthor), which then ful information than females, with the females being often enters the hemocoel and develops into an acanthella with the unidentiable at the specic level in the absence of the males. internal organs of the adult beginning to emerge. When the Nematodes undergo ve developmental stages (four larval proboscis develops, it invaginates into the body and the acan- and one adult), which are separated by a molt of the cuticle, thella becomes encysted and is known as a cystacanth. The with the rst one or two molts occurring within the egg. The nal host acquires the infection either by feeding directly on life cycle of nematodes may be direct or indirect. Direct life an arthropod harboring the cystacanth larva or a vertebrate cycle involves the ingestion of eggs or larvae with food or paratenic host harboring a juvenile form (a reencysted cysta- direct penetration of larvae through the skin. Indirect life canth). Inside the intestine of a suitable nal host, the parasite cycle utilizes invertebrate or vertebrate (or other invertebrate) attaches to the gut wall and matures into adult. as intermediate or paratenic hosts. The larvae normally reside The proboscis with its armature of hooks penetrates the in the tissues of intermediate hosts, while the adults occur in gut wall, causing some local inammation and abdominal the alimentary canal of vertebrates, and occasionally in other pain. Some juvenile parasites may perforate the intestine, parts of the body following larval migration. causing peritonitis. Due to their rare occurrences in humans, With around 12,000 described species, nematodes (round- Acanthocephalans are not discussed in details in the book. worms) are ubiquitous in freshwater, marine, and terrestrial environments, and occur in both invertebrate and vertebrate 1.2.3 ARThROpODS as parasites. The denitive host may acquire the infection by (i) ingestion or penetration of an active, nonparasitic third- Arthropods are invertebrate animals that are characterized stage larva; (ii) ingestion of a passive infective egg contain- by their bilaterally symmetrical, segmented bodies (with ing an infective larva; (iii) ingestion of an intermediate host each segment containing a pair of appendages), jointed limbs containing the infective larva. Once inside the denitive or appendages (specialized for feeding, locomotion, sensing), 6 Molecular Detection of Human Parasitic Pathogens and exoskeleton or cuticle (containing a noncellular material either one or two pairs of wings. The head contains mouth- made of α-chitin, a derivative of glucose, which is secreted parts (which are used for biting, sucking, stinging, licking), by the epidermis). antennae (which are used as tactile organs or as olfactory The cuticle is composed of three main layers: the epicuti- organs), and eyes (which may be compound eyes or simple cle (a thin outer waxy coat that moisture-proofs the other lay- eyes called “ocelli” or “little eyes”). The thorax is further ers and provides protection), the exocuticle (which consists of divided into three segments (pro-thorax, meso-thorax, and chitin and chemically hardened proteins), and the endocuticle meta-thorax), with each segment containing one pair of legs (which consists of chitin and unhardened proteins). Together, (with each leg being made up of coxa, trochanter, femur, the exocuticle and endocuticle are known as the procuticle. tibia, and tarsus). The abdomen has the gonopore (genital While each body segment and limb section is encased in opening) at the posterior end. hardened cuticle, the joints between body segments and As the exoskeleton is unstretchable and restrictive to between limb sections are covered by exible cuticle. The growth, arthropods undergo molting to shed (replace) the old cuticle can have setae (bristles) which grow from specialized exoskeleton after growing a new one. On the basis of their cells in the epidermis, and which function as appendages. differences in the type of metamorphosis (showing variable Other notable features of arthropods include a coelom (or external changes after each molt), class Insecta can be dis- hemocoel), which is a membrane-lined cavity between the tinguished into apterygota, exopterygota, and endopterygota. gut and the body wall that runs most of the length of the body The apterygota have no metamorphosis (i.e., except for the and through which blood ows, and that accommodates the size, all larval stages resemble the adults, which are wing- internal organs (e.g., the reproductive, excretory, and circu- less), the exopterygota undergo a simple metamorphosis latory systems, as well as ladder-like nervous system). The (i.e., there is a gradual change in the external appearance head is formed by fusion of varying numbers of segments, from egg, the nymphal stages to an adult), and the endoptery- and the brain is formed by fusion of the ganglia of these seg- gota has a complete metamorphosis (i.e., there are signicant ments that encircle the esophagus. The heart of arthropods morphological changes from egg, larvae, pupa to adult). is a muscular tube that situates under the back and runs for Of the 20 or so orders within the class Insecta, the follow- most of the length of the hemocoel. By contracting in ripples ing are most notable: Collembola (springtails), Ephemeroptera that run from rear to front, the heart pushes blood forward. (mayies), Odonata (dragonies), Orthoptera (grasshoppers Along the heart are a series of paired ostia, nonreturn valves and crickets), Blattaria (cockroaches), Mantodea (mantids), that allow blood to enter the heart but prevent it from leaving Phasmida (walkingsticks), Isoptera (termites), Anoplura before it reaches the front. Arthropods rely on various com- (sucking lice), Heteroptera (cicadas, hoppers, and aphids), binations of compound eyes and pigment-pit ocelli for vision. Coleoptera (beetles), Lepidoptera (butteries and moths), Arthropods reproduce by internal fertilization via either an Diptera (ies), Siphonaptera (eas), and Hymenoptera (ants, appendage or the ground (rather than by direct injection). wasps, and bees). To date, around 1,000,000 insect species Taxonomically, arthropods are classied in the phylum have been described. Arthropoda (from Greek árthron, “joint”; podós “foot”; together “jointed feet”). The phylum Arthropoda is further 1.3 PHENOTYPIC CHARACTERIZATION divided into ve classes: Arachnida (or Chelicerata, e.g., ticks, mites, spiders, and scorpions), Chilopoda (e.g., centi- Phenotypic techniques for the identication of parasitic pro- pedes), Diplopoda (e.g., millipedes), Crustacea (e.g., crabs, tozoa, helminths, and arthropods include macroscopic and shrimp, lobsters, shrimp, water eas, copepods, barnacles, microscopic assessments (morphometrics) of parasitic mate- sowbugs, and woodlice), and Insecta (or Hexapoda, e.g., ies, rials in clinical and histological specimens, biochemical and eas, wasps, and bees). A majority of medically important immunological characterization, and in vitro and in vivo arthropods are found in the classes Arachnida and Insecta. cultivation, and so on. Class Arachnida have two distinct body regions or tag- mata (cephlothorax and abdomen), two pairs of mouthparts 1.3.1 MACROSCOpIC AND MICROSCOpIC ASSESSMENT (the rst pair being chelicerae, which are three-segmented and pincher-like; the second pair being six-segmented pedi- Macroscopic examination is useful for detection of proglottids palps), and four pairs of legs (most with seven segments; of tapeworms and sometimes adult ascarid worms in stools. compared to insects, an extra segment located between the Direct saline preparations may be employed for identication third segment (the femur) and the fourth segment (the tibia) of trophozoite stages of protozoa. Use of the Kato–Katz method is called the patella), but no antennae. Class Arachnida can is helpful for quantication of the number of eggs present in be further separated into the orders Araneae (spiders, con- stools [3]. Use of modied acid-fast preparations or UV illu- sisting of >2500 described species), Acari or Acarina (mites mination for the examination of auramine-stained preparations and ticks, consisting of >30,000 described species), and facilitates easy identication of Cryptosporidium and Isospora. Scorpiones (scorpions, with >40 species in the United States). To improve the sensitivity of diagnosing protozoal cysts Class Insecta have three distinct body regions or tagmata and helminth eggs, stool samples may be concentrated (head, thorax, and abdomen), three pairs of legs (most with using the formol–ether method and its variants, which also six segments), and one pair of antennae. Many insects have helps remove fecal debris [4,5]. Flotation, sedimentation, Introductory Remarks 7
McMaster method, and Baermann method are other com- However, as a general rule, changes detected by the bio- mon techniques for coprological examination of intestinal chemical tests are non-specic, and further conrmation is parasites such as Entamoeba and Giardia, with a minimum invariably required to achieve a correct diagnosis. of three samples of stool being necessary to make a diagno- Immunological tests [e.g., complement xation test (CFT), sis [6,7]. Similarly, concentration techniques are essential for immunodiffusion (ID), indirect hemagglutination (IHA), identication of parasites in blood. For example, the thick indirect immunouorescent antibody test (IFA), enzyme- lm technique utilizes the hypotonic Field’s stain to lyse linked immunosorbent assay (ELISA), radioimmunoassay red cells, allowing a large number of red cells to be scanned (RIA), and Western blot] demonstrate a higher specicity quickly for the presence of the characteristic ring-shaped than biochemical tests for diagnosis of parasitic infections. merozoites of malaria parasites. Buffy-coat concentration For instance, application of uorescent antibodies permits permits the diagnosis of Leishmania donovani and African rapid and specic diagnosis of P. falciparum and P. vivax. trypanosomes, while centrifugation of blood through a In some cases, immunological tests facilitate differentiation DEAE column also works well for African trypanosomes. In between recent and latent infections, identication of carrier addition, ltration of a measured volume of blood through a status, and verication of parasite elimination after therapies. micropore membrane is useful to concentrate microlariae. Nonetheless, given the frequent antigen sharing among vari- Subsequent staining and microscopic examination enables ous closely related parasites, the results of immunological species identication of microlariae [8]. tests need to be interpreted with accompanied clinical mani- Rectal biopsy samples obtained at sigmoidoscopy or proc- festations and parasitological ndings. toscopy is valuable in the diagnosis of schistosomiasis, espe- cially in travelers who have few eggs present in stools. For 1.3.3 IN ViTRO AND IN VivO CULTIVATION some parasites, biopsy represents the only practical means of making the diagnosis (e.g., cutaneous leishmaniasis or Although culture methods are infrequently used in parasi- onchocerciasis). Additionally, string test may be used to tology, for diagnosis of certain parasites (e.g., Strongyloides diagnose giardiasis, strongyloidiasis, and cryptosporidiosis. and hookworms), an improved yield is obtainable by cultur- In this method, a length of nylon “string” with a weighted end ing stools, especially when the numbers of larvae are low [9]. is swallowed and travels to the upper part of the jejunum (bile In addition, in vitro cultivation of schistosome eggs yields staining of the string indicates that the string has reached the the miracidia for easy identication. Typically, fresh stool or jejunum). After a period of time, the string is pulled up and urine specimens are placed in 10 volumes of unchlorinated the uid expressed is examined. water (e.g., spring water) in an Erlenmeyer ask, which is As morphometric analysis relies on having material in covered. The sediment is then examined after 24 h for the good condition, specimens may be xed by using 10% for- presence of miracidia. Similarly, blood and buffy coat may malin, 70–80% alcohol, Berland’s uid (19 parts glacial be cultured for Leishmania on Schneider’s insect medium for acetic acid to 1 part pure formalin), or boiling water (espe- two weeks to visualize promastigotes. cially in the case of cestodes). Apart from large tapeworms For sensitive diagnosis of Trypanosoma cruzi in blood, and acanthocephlans, specimens should not be xed under uninfected triatomid bugs are permitted to feed on a sus- the pressure (e.g., coverslip or glass slide). After xation for pected patient for ~30 min. The bugs are kept and sacriced about a minute, the specimens are stored in 80% alcohol. monthly for 3 months. Microscopic examination of the dis- Nematodes and acanthocephalans are often examined sected guts helps reveal the presence of trypanosomes. as temporary mounts between a glass slide and a coverslip in a clearing agent (e.g., beechwood creosote, lactophenol, 1.4 GENOTYPIC CHARACTERIZATION or glycerine). Glycerine is only useful for smaller worms (of <15 mm in length). Trematodes and cestodes are mounted Since phenotypic characterization of parasitic protozoa, hel- on slides permanently, followed by staining in a carmine- minths, and arthropods is not only labor intensive, but also based stain (e.g., Mayer’s paracarmine for 1–20 min, depend- requires specialized skills, molecular techniques have been ing on size), destaining in acid alcohol (until the worm is a increasingly utilized for rapid and reliable identication of pale pink color), dehydrating, clearing, and mounting (in parasites. Canada balsam or other mountant).
1.4.1 TARGET GENES 1.3.2 BIOChEMICAL AND IMMUNOLOGICAL TESTS In general, while introns and noncoding regions (e.g., the Biochemical tests (e.g., liver function tests) may be employed internal transcribed spacer or ITS of ribosomal RNA (rRNA) for preliminary diagnosis of blood-dwelling parasites caus- gene) exhibit a high mutational rate (with a high level of inter- ing malaria, theileriosis, babesiosis, trypanosomiasis, and specic sequence variation and a low level of intraspecic lariasis, and so on. Many patients suffering from malaria difference), the coding genes with specic translational prod- and lariasis often show elevated levels of bilirubin, ala- ucts (e.g., the mitochondrial DNA or mtDNA) demonstrate a nine aminotransferase (ALAT), aspartate aminotransfer- relatively low mutational rate (with a low level of interspe- ase (ASAT), and alkaline phosphatase in serum samples. cic sequence variation). For differentiation of protozoa, 8 Molecular Detection of Human Parasitic Pathogens trematoda, cestoda, nematodes, and arthropods at species species, ≥99%; genus, 93–99%; and inconclusive, ≤93%. The level, a target gene or region with a low intraspecic vari- rRNA IGS regions also experience more genetic drift and are ability is preferred to a gene with a high degree of variation. not as highly conserved. This makes the rRNA IGS regions On the other hand, for comparison of two larval populations another potential target for identication. of the same species (e.g., Hypoderma lineatum) originating mtDNA consists of two rRNA genes, 13 protein coding from two geographic areas, a target gene with a high level of genes (including cytochrome c oxidase subunit 1 (cox1), intraspecic variation is preferred (e.g., tandemly repeated cytochrome b (cob), and NADH dehydrogenase subunit 1 sequences or microsatellites of ITS, which are units of repeti- (nad1)) and 22 tRNA genes, which are highly conserved tion between one and ve nucleotides) [10]. within vertebrates and insects, with the exception of tRNAs. Ribosome is an essential cellular organelle that is Because of its conserved sequences and its multiple copies, involved in protein synthesis in all living organisms. As mtDNA is more readily detected than nuclear DNA mark- the key component of the ribosome, rRNA molecules com- ers and thus is widely used for taxonomic, population, and prises two complex folded subunits of differing sizes (small evolutionary investigations in eukaryotes including mam- and large), whose main functions are to provide a mecha- mals, parasites, and arthropods [12]. However, the existence nism for decoding messenger RNA (mRNA) into amino of mitochondrial pseudogenes integrated into a nuclear acids (at the center of the small ribosomal subunit) and to genome may sometimes impact negatively on the reliability interact with transfer RNA (tRNA) during translation by of polymerase chain reaction (PCR)-based mitochondrial providing peptidyltransferase activity (large subunit). The studies [13]. two rRNA subunits in eukaryotes have sedimentation coef- Being the terminal catalyst in the respiratory mitochon- cient values of 40S (Svedberg units) and 60S. The small drial chain, the cytochrome c oxidase subunit I (cox1) of the rRNA subunit (40S) in eukaryotes contains a single RNA mitochondrial genome is particularly suitable as a molecular species (i.e., 18S rRNA) whereas the large rRNA subunit marker for the taxonomic differentiation and evolutionary (60S) in eukaryotes comprises three RNA species (5S, 5.8S, studies of insects because of its large size and the presence of and 25–28S rRNA) [11]. highly conserved and variable regions. The COI amino acid The eukaryotic rRNA genes exist as a family of multiple- sequence comprises 12 transmembrane helices (M1–M12), copy genes that are arranged in a head-to-toe manner, six external loops (E1–E6), ve internal loops (I1–I5), one with each copy consisting of 18S RNA, ITS 1 (internal carboxyl (COOH), and one amino (NH2) terminal. The nico- transcribed spacer 1), 5.8S RNA, ITS 2, 28S RNA, IGS tinamide dehydrogenase subunit 4 (nad4) gene is also useful 1 (intergenic spacer), 5S, and IGS 2. Having been homog- for studying genetic variation both within and among popula- enized through concerted evolution, the tandemly repeated tions of key parasitic helminthes. copies of rRNA genes contain highly similar nucleotide sequence, and thus are often treated as a single-locus gene. 1.4.2 TEMpLATE AMpLIFICATION AND PRODUCT DETECTION Along with ITS, the 18S, 5.8S, and 25–28S rRNAs are transcribed as a 35–40S precursor, with all spacers being 1.4.2.1 Template Amplification later spliced out of the transcript. A nontranscribed or The early molecular procedures are nonamplication based IGS region exists between the copies of the 18S, 5.8S, and and suffer from poor sensitivity and time-consumingness. 25–28S rRNA repeats, serving to separate the repeats from With the development of PCR and other nucleic acid ampli- one another on the chromosome. A 5S RNA gene takes a cation technologies (such as ligase chain reaction (LCR), position within the IGS region and is transcribed in the nucleic acid sequence-based amplication (NASBA), opposite direction. transcription-mediated amplication (TMA), strand dis- In general the 25–28S RNA gene is conserved across placement amplication (SDA), rolling circle amplica- widely divergent taxa, and only the rst 600–900 bases con- tion (RCA), cycling probe technology (CPT), branched tain three divergent domains (D1, D2, D3) that are useful for DNA (bDNA), and loop-mediated isothermal amplication phylogenetic study of eukaryotic organisms such as para- (LAMP)), it has become possible to rapidly and specically sitic worms and arthropods. The 18S rRNA (small subunit detect a single copy of nucleic acid in a few hours. (SSU)) gene also includes alternating regions of sequence Given their efciency, simplicity, robustness, and versatil- conservation and heterogeneity. The conserved regions are ity, PCR and its derivatives have been widely adopted in both often targeted for phylogenetic analysis of higher taxonomic research and clinical laboratories for identication and deter- orders (e.g., phylum, family, and genus), while the regions mination of parasitic protozoa, helminths, and arthropods. of sequence diversity are valuable for characterization of The amplication formats have also evolved from single- isolates to the genus or species level (with isolates showing round PCR to nested PCR, multiplex PCR, reverse transcrip- sequence identity >97% in the 18S rRNA gene being consid- tion-PCR (RT-PCR), real-time PCR, and quantitative PCR ered as the identical species). Compared to the rRNA genes, (Q-PCR), and so on, leading to enhanced assay sensitivity the rRNA ITS 1 and ITS 2 regions tend to be more variable, (nested PCR) and versatility (multiplex PCR detecting mul- offering valuable targets for species-specic identication. tiple genes and/or organisms; and RT-PCR detecting RNA The identity of the testing organism is dened by its ITS instead of DNA) as well as accurate quantitation (Q-PCR) sequence similarity (%) to the type strain or control isolate: and instant result availability (real-time PCR) [14]. Introductory Remarks 9
In particular, PCR-restriction fragment-length polymor- amplicon are then compared with those stored at reference phism (PCR-RFLP) has been widely applied for the identi- databases such as GenBank, and the phylogenetic relation- cation of closely related taxa of insects (e.g., Calliphoridae ships of related parasitic protozoa, helminths, and arthropods and Sarcophagidae families). Single-strand conformation are displayed in the form of trees, constructed with distance polymorphism analysis (SSCP) is a PCR-based mutation matrix methods (resulting in phenograms), and maximum- scanning technique that is based on the different electropho- parsimony methods (resulting in cladograms). retic mobility of ssDNA in a nondenaturing gel. The SSCP analysis is useful for assessing genetic distance and identify- 1.4.3 DATA ANALYSIS ing haplotypes existing in insect populations from different geographic areas belonging to the same species (e.g., ies or Pairwise nucleotide comparison of test sample with publicly mosquitoes) or to identify morphologically indistinguishable available gene databases (such as GenBank) by BLASTn species (e.g., mosquitoes). Random amplied polymorphic analysis provides a straightforward means to identify the DNA (RAPD) analysis relies on the use of a single arbi- presence or absence of a particular sequence motif or even trary primer under low annealing temperature conditions to a single nucleotide and to ascertain the relatedness of organ- amplify DNA sequences and screen genetic variation across isms of interest. For evolutionary studies, multiple sequence a genome. alignment and the true homology between variable sites and portions of sequences must be addressed. One or more 1.4.2.2 Product Detection alignment approaches may be employed: (i) on the basis of In its conventional (standard) form, PCR products are sepa- secondary structure and functional domains; (ii) using one rated by agarose gel electrophoresis with or without modica- of a range of specialist alignment programs with various tion (e.g., enzymatic digestion), stained with a DNA-binding weighting options and gap penalties (e.g., ClustalX); (iii) by dye (e.g., ethidium bromide or gel star), and visualized under eye, often in relation to previously aligned sequences. Three UV light. For PCR products <100 bp or for distinction of main categories of phylogenetic analysis with molecular data products with minor size differences, polyacrylamide gel include distance methods (e.g., phenetic methods), cladistics/ electrophoresis, and its derivatives (e.g., single-strand con- parsimony, and maximum-likelihood analysis. Bootstrap formational polymorphism analysis (SSCP), denaturing gra- analysis is the most commonly employed statistical method dient gel electrophoresis (DGGE), and temperature-gradient to provide support for the evolutionary relationships pre- gel electrophoresis (TGGE)) may be employed. sented [15]. To further improve the sensitivity of PCR product detec- tion, enzymatic signal amplication (e.g., ELISA and ow 1.5 RESULTS INTERPRETATION, cytometry), DNA microarray (also known as DNA chip, gene or genome chip, or gene array), and line probe assay STANDARDIZATION, AND QUALITY (LiPA) (which uses a nitrocellulose strip instead of a glass, CONTROL AND ASSURANCE silicon, plastic chip as in the case of DNA array) have been 1.5.1 KEY PERFORMANCE ChARACTERISTICS developed. Recent advances in instrument automation and uorescent dye chemistry facilitate instant monitoring of The performance of a diagnostic assay is often evaluated by PCR amplicons (real-time PCR) without additional manual using several key parameters, including detection limit, sen- handling. Real-time PCR may be based on a double-stranded sitivity, specicity, accuracy, intra-assay precision, interassay DNA intercalating dye (e.g., SYBR Green), or specically precision, and linearity (as in the case of a quantitative assay). designed probes that target a region of amplicon and incor- Detection limit (or limit of detection) is the lowest concentra- porate a uorescent dye. Examples of these probes include tion or quantity of bacteria that can be detected by a given hydrolysis dual-labeled probes (TaqMan®), hybridization assay. Sensitivity is the percentage of samples containing probes (LightCycler), molecular beacons, peptide nucleic bacteria of interest that are identied by the assay as posi- acid (PNA) probes, TaqMan minor groove binding (MGB™) tive for the bacteria. Specicity is the percentage of samples probes, locked nucleic acid (LNA®) primers and probes, without bacteria of interest that are identied by the assay as and scorpions™. DNA sequencing analysis provides a most negative for the bacteria. Accuracy (or trueness) is the degree accurate way to ascertain the identity of PCR amplicons. The of conformity of an assay’s measurements to the actual (true) classical chain termination sequencing method (or Sanger value. It is often estimated by analyses of reference materials method) utilizes primers or dideoxynucleotides that are or comparisons of results with those obtained by a reference labeled with radioactive isotope or uorescent tag, and the method. The closer an assay’s measurements to the accepted sequencing products are detected by exposure to x-rays or value, the more accurate the assay is. Precision is the degree UV-light. More recently, the 454 Genome Sequencer (Roche), of mutual agreement among a series of assay’s individual the Genome Analyzer II (Illumina), and the SOLiD3 System measurements, values, or results. Usually characterized in (Applied Biosystems) (collectively referred to as “next- terms of the standard deviation of the measurements, pre- generation sequencing” sequencing or NGS technologies) cision can be stratied into (i) repeatability—the variation have been adopted for high-throughput sequencing analy- arising using the same instrument and operator in a single sis of PCR products. The nucleotide sequences of the PCR run (i.e., intra-assay precision) or repeating during a short 10 Molecular Detection of Human Parasitic Pathogens time period; and (ii) reproducibility—the variation arising may be due to the impurity of the processed sample. Enzymes using the same measurement process among different instru- (e.g., DNA polymerase, reverse transcriptase) used in PCR ments and operators from one run to another (i.e., inter-assay and RT-PCR are impeded by components in blood and feces precision) or over longer time periods. Linearity refers to the (e.g., heme, hemoglobulin, lactoferrin, immunoglobulin G, tendency of measurements by a quantitative assay to form leukocyte DNA, polysaccharides, and urea), in foods (e.g., a straight line when plotted on a graph. Data from linearity phenolics, glycogen, calcium ions, fat, and other organic experiments may be subjected to linear regression analysis substances), in environmental specimens (e.g., phenolics, with an ideal regression coefcient of 1. In the case of a non- humic acids, and heavy metals); and in added anticoagulants linear curve, other objective, statistically valid methods may (e.g., EDTA and heparin) as well as nucleic acid purication be utilized. reagents (e.g., detergents, lysozyme, NaOH, alcohol, EDTA, EGTA, phenol, and high salt concentrations) [16]. Any impu- rities and contaminations present in the samples after nucleic 1.5.2 RESULTS INTERpRETATION acid isolation may contribute to false-negative results. A use- A positive result by a molecular assay for a given pathogen ful way to determine the effective of nucleic acid purica- normally conrms the etiologic relationship if the clini- tion procedure for removing inhibitory substances is to spike cal syndrome is compatible with the pathogen identied. samples with well-dened DNA or RNA prior to and after Considering the sensitive nature of the amplied methods sample preparation (as process and amplication internal such as PCR, it is important to rule out the possibility of a controls). In light of the high sensitivity of PCR, the occur- false-positive result. Occasionally, false-positive results may rence of false-negative results is probably a truly underesti- originate from the low diagnostic specicity of the assay, in mated problem. which primers bind to irrelevant sequences and occasion- ally a homologous sequence that is shared among related or 1.5.3 STANDARDIZATION AND VALIDATION unknown bacteria. More often, false-positive results in the molecular testing come from contamination, which may As molecular tests such as PCR and sequencing offer arise during manual handling of the samples in the testing improved sensitivity, specicity, accuracy, precision, and laboratory either at the pre or postextraction (while setting result availability for identication and diagnosis of para- up the PCR) stages. This risk is heightened when a high sites, they have been increasingly adopted and applied in copy-number polynucleotide (or plasmid) is used as a quanti- routine diagnostic laboratories. Considering the possibility cation standard and distributed around the laboratory, con- of false-positive and false-negative results that may occur in taminating reaction source. Additionally, contamination may these highly sensitive tests, it is essential to properly stan- be attributable to samples that are referred from other labora- dardize and validate them prior to their adoption, and to put tories, which do not utilize manipulation techniques that are in place appropriate quality control measures to ensure their mandatory for the molecular testing. These may include the consistent performance. use of unplugged pipette tips, infrequent changing of gloves Standardization of molecular tests addresses the need and using pipette for long periods without decontamination. for standardized reagents and common units, contamina- Another cause of contamination is by amplication products tion control mechanisms, inhibition control mechanisms, from previous tests. Contamination may also occur by leak- clinically relevant dynamic ranges and internal controls, and age from tubes or microtiter plates with lids not tightly closed so on. Validation helps to verify the sensitivity, specicity, or by breakage of glass capillaries leading to spillage of the accuracy, repeatability (intra-assay precision), reproducibil- amplication mixture. Besides the adoption of stringent labo- ity (inter-assay precision), detection limit, and linearity (if ratory practice, the risk of contamination with PCR products quantitative) of molecular tests. may be reduced by replacing nucleotide dTTP with dUTP Before validating a method, it is important to have all in PCR, and implementing a digestion step with Uracil- instruments calibrated and maintained throughout the test- DNA-glycosylase (UNG) to remove previous PCR prod- ing process. The validation process may involve a series of ucts containing dUTP prior to each amplication reaction. steps including: (i) testing of dilution series of positive sam- Furthermore, inclusion of multiple negative controls, such as ples (or plasmid construct) to determine the limits of detec- no-template controls (NTC) and no-amplication controls tion of the assay and their linearity over concentrations to (NAC) may help identify the likely source of contamination be measured in quantitative test (using minimal number of and prevent false-positive results. Moreover, microbial DNA reference calibrators such as previously tested patient sam- may come with PCR reagents. ples or pooled sera); (ii) evaluating the sensitivity and speci- Similarly, a negative result by a molecular assay for a city of the assay, along with the extent of cross-reactivity given pathogen normally indicates the absence of the patho- with other genomic material; (iii) establishing the day-to-day gen. However, it is equally important to rule out the possibil- variation of the assay’s performance; (iv) assuring the qual- ity of false-negative results. One possible cause is due to the ity of assembled assays using quality control procedures that low sensitivity of the assay employed. Alternatively, insuf- monitor the performance of reagent batches; and (v) align- cient amount of bacteria may be present in the sample (due ing the in-house primer and probe sequences with a genome to sample degradation or prior antibiotic treatment). Another sequence databank to avoid extended specicity testing. Introductory Remarks 11
1.5.4 QUALITY CONTROL AND ASSURANCE sample to Site A to check for reproducibility. (v) A detailed record of distributions is kept to provide an audit trail. 1.5.4.1 Quality Control Quality control strategies for nucleic acid-based tests include (i) designation of a “clean” area for reaction setup 1.6 CONCLUSION (e.g., room under negative air pressure; positive-displace- ment pipettes; aerosol-block pipette tips; UV-equipped Parasites are eukaryotic organisms that are separated into PCR cabinet); (ii) use of personal protective equipment three categories: protozoa, helminths, and arthropods. (PPE) (e.g., disposable gloves and lab coats to prevent intro- Although a majority of the 800,000 or so parasite species are duction of contaminating DNA or nucleases); (iii) use of free-living or form symbiotic relationship with their hosts, uracil-N-glycosylase (UNG) in real-time PCR (to eliminate causing no obvious adverse effects, a small proportion have cross-over amplicon contamination); (iv) use of a “hot-start” the capacity to induce clinical symptoms of various sever- method (to minimize false priming events by withholding a ity in affected individuals. Due to the fact that many parasite crucial reaction component until appropriate temperature is species share considerable morphological and biological simi- reached); (v) Use of external positive and negative controls larities, and that clinical features resulting from the infection (to monitor reaction performance and contamination) and with parasitic protozoa, helminths, and arthropods are often homologous or heterologous internal controls (to monitor nonspecic, it is important that laboratory techniques are presence of inhibitors). applied for their correct identication in order to facilitate the A variety of test controls may be considered for diagnostic implementation of effective control and prevention measures. PCR. These include (i) internal amplication control (IAC) While phenotypic procedures such as macroscopic and (negative sample spiked with sufcient pathogen and pro- microscopic examination, biochemical and immunological cessed throughout the entire protocol); (ii) processing posi- assays, and in vitro/in vivo culture provide a useful means for tive control (PPC) (negative sample spiked with sufcient diagnosis of parasitic infections, they suffer from the draw- closely related, but nontarget, strain processed throughout backs of being slow, variable at times, costly, and technically the entire protocol); (iii) reagent control (blank) (containing demanding. The recent development of molecular methodol- all reagents, but no nucleic acid apart from the primers); (iv) ogies offers an opportunity to improve the sensitivity, speci- premises control (tube containing the master mixture left city, and speed of parasite detection and identication. In open in the PCR setup room to detect possible contaminating particular, PCR amplication of ribosomal RNA and other DNA in the environment (carried out at regular intervals as gene regions followed by sequencing analysis represents a part of the quality assurance program); (v) standard (three rapid and accurate approach for identication and differen- to four samples containing 10-fold dilution series of known tiation of parasitic pathogens. number of target DNA copies in a range). There is no doubt that with an increasing adoption and utilization of molecular methods in and clinical laboratories, reliable identication and detection of parasitic protozoa, 1.5.4.2 Quality Assurance helminths, and arthropods directly from patient materials is One way to assess preparedness of the diagnostic laborato- no longer an impossible dream. Furthermore, since molecu- ries is through the conduct of an external quality assurance lar techniques enable precise epidemiological tracking and (EQA) program providing characterized specimens contain- phylogenetic analysis of parasitic pathogens involved in dis- ing pathogens of interest. The design of a quality assurance ease epidemics, they will play an even more prominent role program has the following components: (i) internal quality in the future control and prevention campaigns against para- control (IQC) materials are distributed every month and sitic infections in human populations. comprising three pools of clinical samples of known patho- gen status (typically one negative, one positive containing 1 log10 over the lower limit of detection of the assay, and one REFERENCES low positive containing up to 1 log10 of the lower limit of detection of the assay). These are incorporated in test runs 1. http://www.uniprot.org/; last accessed on 20 May 2011. 2. http://www.data.gbif.org/; Global Biodiversity Information on a weekly basis. The purpose of IQC is to provide samples Facility data portal, published by Field Museum of Natural of known status for repeated testing in parallel with clini- History, Museum of Vertebrate Zoology, University of cal samples to ensure reproducibility of the test system in Washington Burke Museum, and University of Turku; last an individual laboratory. (ii) EQA distributions of panels accessed on 20 May 2011. of ve unknown samples distributed quarterly. Results are 3. Katz, N., Chaves, A., and Pellegrino, J. A simple device for returned to the QA laboratory for assessment. EQA compares quantitative stool thick-smear technique in Schistosomiasis the performance of different testing sites using specimens of mansoni. Rev. Inst. Med. Trop. Sao Paulo 14, 397, 1972. known, but undisclosed, content. (iii) Aliquots of all samples 4. Allen, A.V.H., and Ridley, D.S. Further observations on the formol-ether concentration technique for fecal parasites. sent from the reference laboratory are posted back to Site A J. Clin. Pathol. 23, 545, 1970. for repeat testing to check for integrity of the pools and for 5. Ramsay, A. et al. A eld evaluation of the formol detergent transport problems. (iv) A nal element of the pilot program method for concentrating fecal parasites. J. Trop. Med. Hyg. involves Sites B, C, and D sending an aliquot of every 50th 94, 210, 1991. 12 Molecular Detection of Human Parasitic Pathogens
6. Bell, D.R. Diagnosis of parasitic diseases by ltration. Ann. 12. Meusemann, K. et al. A phylogenomic approach to Soc. Belg. Med. Trop. 55, 489, 1975. resolve the arthropod tree of life. Mol. Biol. Evol. 27, 2541, 7. Peters, P.A. et al. Rapid, accurate quantication of schisto- 2010. some eggs via nuclepore lters. J. Parasitol. 62, 154, 1976. 13. Jex, A.R., Littlewood, D.T., and Gasser, R.B. Toward next- 8. Scheiber, P., Braun-Munzinger, R.A., and Southgate, B.A. generation sequencing of mitochondrial genomes—Focus on A new technique for the determination of microlariael densi- parasitic worms of animals and biotechnological implications. ties in onchocerciasis. Bull. World Health Organ. 53, 130, 1976. Biotechnol. Adv. 28, 151, 2010. 9. Arakaki, T.J. et al. Efcacy of agar-plate culture in detection of 14. Bretagne, S., and Costa, J.-M. Towards a nucleic acid-based Strongyloides stercoralis infection. Parasitology 76, 425, 1990. diagnosis in clinical parasitology and mycology. Clin. Chim. 10. Barker, G.C. Microsatellite DNA: A tool for population genetic Acta 363, 221, 2006. analysis. Trans. R. Soc. Trop. Med. Hyg. 96, S21, 2002. 15. Liu, L. et al. Estimating species trees using multiple-allele 11. Hillis, D.M., and Dixon, M.T. Ribosomal DNA: Molecular DNA sequence data. Evolution 62, 2080, 2008. evolution and phylogenetic inference. Quart. Rev. Biol. 66, 16. Wilson, I.G. Inhibition and facilitation of nucleic acid ampli- 411, 1991. cation. Appl. Environ. Microbiol. 63, 3741, 1997. Section I
Sarcomastigophora
References
1 Chapter 1 - Introductory Remarks
6. Bell, D.R. Diagnosis of parasitic diseases by �ltration. Ann. Soc. Belg. Med. Trop. 55, 489, 1975.
7. Peters, P.A. et al. Rapid, accurate quanti�cation of schistosome eggs via nuclepore �lters. J. Parasitol. 62, 154, 1976.
8. Scheiber, P., Braun-Munzinger, R.A., and Southgate, B.A. A new technique for the determination of micro�lariael densities in onchocerciasis. Bull. World Health Organ. 53, 130, 1976.
9. Arakaki, T.J. et al. Ef�cacy of agar-plate culture in detection of Strongyloides stercoralis infection. Parasitology 76, 425, 1990.
10. Barker, G.C. Microsatellite DNA: A tool for population genetic analysis. Trans. R. Soc. Trop. Med. Hyg. 96, S21, 2002.
11. Hillis, D.M., and Dixon, M.T. Ribosomal DNA: Molecular evolution and phylogenetic inference. Quart. Rev. Biol. 66, 411, 1991. 12. Meusemann, K. et al. A phylogenomic approach to resolve the arthropod tree of life. Mol. Biol. Evol. 27, 2541, 2010. 13. Jex, A.R., Littlewood, D.T., and Gasser, R.B. Toward nextgeneration sequencing of mitochondrial genomes—Focus on parasitic worms of animals and biotechnological implications. Biotechnol. Adv. 28, 151, 2010. 14. Bretagne, S., and Costa, J.-M. Towards a nucleic acid-based diagnosis in clinical parasitology and mycology. Clin. Chim. Acta 363, 221, 2006. 15. Liu, L. et al. Estimating species trees using multiple-allele DNA sequence data. Evolution 62, 2080, 2008. 16. Wilson, I.G. Inhibition and facilitation of nucleic acid ampli�cation. Appl. Environ. Microbiol. 63, 3741, 1997.
Section I
Sarcomastigophora 2 Chapter 2 - Acanthamoeba
1. Visvesvara, G.S., Moura, H., and Schuster, F.L. Pathogenic and opportunistic free-living amoebae: Acanthamoeba spp., Balamuthia mandrillaris, Naegleria fowleri, and Sappinia diploidea. FEMS Immunol. Med. Microbiol. 50, 1, 2007.
2. Adl, S.M. et al. The new higher level classi�cation of eukaryotes with emphasis on the taxonomy of protists. J. Eukaryot. Microbiol. 52, 399, 2005.
3. Marciano-Cabral, F., and Cabral, G. Acanthamoeba spp. as agents of disease in humans. Clin. Microbiol. Rev. 16, 273, 2003.
4. Khan, N.A. Acanthamoeba: Biology and increasing importance in human health. FEMS Microbiol. Rev. 30, 564, 2006.
5. Page, F.C. Re-de�nition of the genus Acanthamoeba with descriptions of three species. J. Protozool. 14, 709, 1967.
6. Illingworth, C.D., and Cook, S.D. Acanthamoeba keratitis. Surv. Ophthalmol. 42, 493, 1998.
7. Alsam, S. et al. Acanthamoeba interactions with human brain microvascular endothelial cells. Microb. Pathog. 35, 235, 2003.
8. Clarke, D.W., and Niederkorn, J.Y. The pathophysiology of Acanthamoeba keratitis. Trends Parasitol. 22, 175, 2006.
9. Kim, W.T. et al. Comparison of speci�c activity and cytopathic effects of puri�ed 33 kDa serine proteinase from Acanthamoeba strains with different degree of virulence. Korean J. Parasitol. 44, 321, 2006. 10. Radford, C.F. et al. Risk factors for Acanthamoeba keratitis in contact lens users: A case–control study. Br. Med. J. 310, 1567, 1995. 11. Sharma, S., Garg, P., and Rao, G.N. Patient characteristics, diagnosis, and treatment of non-contact lens related Acanthamoeba keratitis. Br. J. Ophthalmol. 84, 1103, 2000. 12. Pasricha, G. et al. Use of 18S rRNA gene-based PCR assay for diagnosis of Acanthamoeba keratitis in non-contact lens wearers in India. J. Clin. Microbiol. 41, 3206, 2003. 13. Syam, P.P., Narendran, R., and van der Hoek, J. Persistent Acanthamoeba keratitis in a non-contact lens wearer following exposure to bird seed dust. Br. J. Ophthalmol. 89, 388, 2005. 14. Hurt, M. et al. Pathogenic Acanthamoeba spp. secrete a mannose-induced cytolytic protein that correlates with the ability to cause disease. Infect. Immun. 71, 6243, 2003. 15. Dart, J.K., Saw, V.P., and Kilvington, S. Acanthamoeba keratitis: Diagnosis and treatment update, 2009. Am. J. Ophthalmol. 148, 487, 2009. 16. Cardine, S. et al. Clinical management and prognosis in Acanthamoeba keratitis: A retrospective study of 25 cases. J. Fr. Ophtalmol. 25, 1007, 2002. 17. Moshari, A. et al. Chorioretinitis after keratitis caused by Acanthamoeba: Case report and review of the literature. Ophthalmology 108, 2232, 2001. 18. Hef�er, K.F. et al. Acanthamoeba endophthalmitis in acquired immunode�ciency syndrome. Am. J. Ophthalmol. 122, 584, 1996. 19. Martinez, A.J., and Visvesvara, G.S. Free-living, amphizoic and opportunistic amebas. Brain Pathol. 7, 583, 1997. 20. Yagi, S., Schuster, F.L., and Visvesvara, G.S. Demonstration of Balamuthia and Acanthamoeba mitochondrial DNA in sectioned archival brain and other tissues by polymerase chain reaction. Parasitol. Res. 102, 491, 2008. 21. Alsam, S. et al. Extracellular proteases of Acanthamoeba castellanii (encephalitis isolate belonging to T1 genotype) contribute to increased permeability in an in vitro model of the human blood–brain barrier. J. Infect. 51, 150, 2005. 22. Kong, H.H., Kim, T.H., and Chung, D.I. Puri�cation and characterization of a secretory serine proteinase of Acanthamoeba healyi isolated from GAE. J. Parasitol. 86, 12, 2000. 23. Lehmann, O.J. et al. Polymerase chain reaction analysis of corneal epithelial and tear samples in the diagnosis of Acanthamoeba keratitis. Invest. Ophthalmol. Vis. Sci. 39, 1261, 1998. 24. Mathers, W.D. et al. Con�rmation of confocal microscopy diagnosis of Acanthamoeba keratitis using polymerase chain reaction analysis. Arch. Ophthalmol. 118, 178, 2000. 25. Stothard, D.R. et al. Fluorescent oligonucleotide probes for clinical and environmental detection of Acanthamoeba and the T4 18S rRNA gene sequence type. J. Clin. Microbiol. 37, 2687, 1999. 26. Epstein, R.J. et al. Rapid diagnosis of Acanthamoeba keratitis from corneal scrapings using indirect �uorescent antibody staining. Arch. Ophthalmol. 104, 1318, 1986. 27. Schuster, F.L. et al. Detection of antibodies against free-living amoebae Balamuthia mandrillaris and Acanthamoeba species in a population of patients with encephalitis. Clin. Infect. Dis. 42, 1260, 1986. 28. Chappell, C.L. et al. Standardized method of measuring Acanthamoeba antibodies in sera from healthy human subjects. Clin. Diagn. Lab. Immunol. 8, 724, 2001.
29. Grunnet, M.L., Cannon, G.H., and Kushner, J.P. Fulminant amebic meningoencephalitis due to Acanthamoeba. Neurology 31, 174, 1981.
30. Bloch, K.C. and Schuster, F.L. Inability to make a premortem diagnosis of Acanthamoeba species infection in a patient with fatal granulomatous amebic encephalitis. J. Clin. Microbiol. 43, 3003, 2005.
31. McKellar, M.S. et al. Fatal granulomatous Acanthamoeba encephalitis mimicking a stroke, diagnosed by correlation of results of sequential magnetic resonance imaging, biopsy, in vitro culture, immuno�uorescence analysis, and molecular analysis. J. Clin. Microbiol. 44, 4265, 2006.
32. Barete, S. et al. Fatal disseminated Acanthamoeba lenticulata infection in a heart transplant patient. Emerg. Infect. Dis. 13, 736, 2007.
33. Petry, F. et al. Early diagnosis of Acanthamoeba infection during routine cytological examination of cerebrospinal �uid. J. Clin. Microbiol. 44, 1903, 2006.
34. Zam�r, O. et al. Diagnostic par PCR des kératites à Acanthamoeba spp. J. Fr. Ophthalmol. 29, 1034, 2006.
35. Yera, H. et al. Comparison of PCR, microscopic examination and culture in the early diagnosis and characterization of Acanthamoeba isolates from ocular infections. Eur. J. Clin. Microbiol. Inf. Dis. 26, 221, 2007.
36. Schroeder, J.M. et al. Use of subgenic 18S ribosomal DNA PCR and sequencing for genus and genotype identi�cation of Acanthamoeba from humans with keratitis and from sewage sludge. J. Clin. Microbiol. 39, 1903, 2001.
37. Sharma, S. et al. Acanthamoeba keratitis in non-contact lens wearers in India: DNA typing-based validation and a simple detection assay. Arch. Ophthalmol. 122, 1430, 2004.
38. Dhivya, S. et al. Comparison of a novel semi-nested polymerase chain reaction (PCR) with a uniplex PCR for the detection of Acanthamoeba genome in corneal scrapings. Parasitol. Res. 100, 1303, 2007.
39. Shirwadkar, C.G. et al. Acanthamoeba encephalitis in patient with systemic lupus, India. Emerg. Infect. Dis. 12, 84, 2006.
40. MacLean, R.C. et al. Identi�cation of Acanthamoeba sp. in paraf�n-embedded CNS tissue from an HIV+ individual by PCR. Diagn. Microbiol. Infect. Dis., 57, 289, 2007.
41. Meersseman, W. et al. Rapidly fatal Acanthamoeba encephalitis and treatment of cryoglobulinemia. Emerg. Infect. Dis. 13, 469, 2007.
42. Yagi, S., Schuster, F.L., and Bloch, K. Demonstration of presence of Acanthamoeba mitochondrial DNA in brain tissue and cerebrospinal �uid by PCR in samples from a patient who died of granulomatous amebic encephalitis. J. Clin. Microbiol. 45, 2090, 2007.
43. Walochnik, J. et al. Granulomatous amoebic encephalitis caused by Acanthamoeba amoebae of genotype T2 in a human immunode�ciency virus-negative patient. J. Clin. Microbiol. 46, 338, 2008.
44. Rivière, D. et al. Development of a real-time PCR assay for quanti�cation of Acanthamoeba trophozoites and cysts. J. Microbiol. Methods. 64, 78, 2006.
45. Qvarnstrom, Y. et al. Multiplex real-time PCR assay for the simultaneous detection of Acanthamoeba spp., Balamuthia mandrillaris, and Naegleria fowleri. J. Clin. Microbiol. 44, 3589, 2006.
46. Thompson, P.P. et al. Validation of real-time PCR for laboratory diagnosis of Acanthamoeba keratitis. J. Clin. Microbiol. 46, 3232, 2008.
47. Goldschmidt, P. et al. New tool for the simultaneous detection of 10 different genotypes of Acanthamoeba available from the American Type Culture Collection. Br. J. Ophthalmol. 93, 1086, 2009. 48. Pussard, M., and Pons. R. Morphologie de la paroi kystique et taxonomie du genre Acanthamoeba (Protozoa, Amoebida). Protistologica 13, 557, 1977. 49. Visvesvara, G.S. Classi�cation of Acanthamoeba. Rev. Infect. Dis. 13(Suppl 5), S369, 1991. 50. Gast, R.J. et al. Subgenus systematics of Acanthamoeba: Four nuclear 18S rDNA sequence types. J. Eukaryot. Microbiol. 43, 498, 1996. 51. Stothard, D.R. et al. The evolutionary history of the genus Acanthamoeba and the identi�cation of eight new 18S rRNA gene sequence types. J. Eukaryot. Microbiol. 45, 45, 1998. 52. Hewett, M.K. et al. Identi�cation of a new Acanthamoeba jacobsi Sawyer, Nerad and Visvesvara (1992) (Lobosea: Acanthamoebidae). Acta Protozool. 42, 325, 2003. 53. Horn, M. et al. Novel bacterial endosymbionts of Acanthamoeba spp. related to the Paramecium caudatum symbiont Caedibacter caryophilus. Environ. Microbiol. 1, 357, 1999. 54. Gast, R.J., Development of an Acanthamoeba-speci�c reverse dot-blot and the discovery of a New ribotype. J. Eukaryot. Microbiol. 48, 609, 2001. 55. Booton, G.C. et al. 18S ribosomal DNA typing and tracking of Acanthamoeba species isolates from corneal scrape specimens, contact lenses, lens cases, and home water supplies of Acanthamoeba keratitis patients in Hong Kong. J. Clin. Microbiol. 40, 1621, 2002. 56. Fuerst, P.A. et al. Genotypic identi�cation of no-keratitis infections caused by the opportunistically pathogenic ameba genus Acanthamoeba. J. Eukaryot. Microbiol. 50 (Suppl), 512, 2003. 57. Zhang, Y. et al. Identi�cation of 18S ribosomal DNA genotype of Acanthamoeba from patients with keratitis in North China. Invest. Ophthalmol. Vis. Sci. 45, 1904, 2004. 58. Booton, G.C. et al. Identi�cation and distribution of Acanthamoeba species genotypes associated with nonkeratitis infections. J. Clin. Microbiol. 43, 1689, 2005. 59. Yera, H. et al. The genotypic characterization of Acanthamoeba isolates from human ocular samples. Br. J. Ophthalmol. 92, 1139, 2008. 60. Spanakos, G. et al. Genotyping of pathogenic Acanthamoebae isolated from clinical samples in Greece—Report of a clinical isolate presenting T5 genotype. Parasitol. Int. 55, 147, 2006. 61. Di Cave, D. et al. Acanthamoeba T4 and T15 genotypes associated with keratitis infections in Italy. Eur. J. Clin. Microbiol. Inf. Dis. 28, 607, 2009. 62. Lackner, P. et al. Acute granulomatous Acanthamoeba encephalitis in an immunocompetent patient. Neurocrit. Care 12, 91, 2010. 63. Ledee, D.R. et al. Advantages of using mitochondrial 16S rDNA sequences to classify clinical isolates of Acanthamoeba, Invest. Ophthalmol. Vis. Sci. 44, 1142, 2003. 64. Köhsler, M. et al. ITS1 sequence variabilities correlate with 18S rDNA sequence types in the genus Acanthamoeba (Protozoa: Amoebozoa). Parasitol. Res. 98, 86, 2006. 65. Goldschmidt, P. et al. Effects of topical anaesthetics and �uorescein on the real-time PCR used for the diagnosis of Herpesviruses and Acanthamoeba keratitis. Br. J. Ophthalmol. 90, 1354, 2006. 66. Goldschmidt, P. et al. Resistance of Acanthamoeba to classic DNA extraction methods used for the diagnosis of corneal infections. Br. J. Ophthalmol. 92, 112, 2008. 3 Chapter 3 - Balamuthia
TABLE 3.1
PCR Primers and Probes for Specific Detection of Balamuthia mandrillaris
PCR
Assay Oligonucleotide Name Sequence (5’–3’) Amplicon Size (Base Pairs) Detection Mode Reference
1 5′Balspec16S CGCATGTATGAAGAAGACCA 1075 Gel electrophoresis 18 3′Balspec16S TTACCTATATAATTGTCGATACCA
2 5′Balspec16S CGCATGTATGAAGAAGACCA 230 Gel electrophoresis 95 Bal16Sr610 CCCCTTTTTAACTCTAGTCATATAGT
3 BalaF1451 TAA CCT GCT AAA TAG TCA TGC CAA T 171 Fluorescence (real-time PCR) 96 BalaR1621 CAA ACT TCC CTC GGC TAA TCA BalaP1582 (R)-AGT ACT TCT ACC AAT CCA ACC GCC A-(Q)
4 RNaseP/FW GGC AGG TTC CGA GGA GAC A 82 Fluorescence (real-time PCR) 97 RNaseP/RV GTG GCC TTG TGT ATT GAA CTT AAC ATT RNaseP/probe (R)-TG GAA CCA TAC CTT GGG TGA CAC GAT G-(Q)
10. Matin, A. et al. Increasing importance of Balamuthia mandrillaris. Clin. Microbiol. Rev. 21, 435, 2008.
11. Gelman, B.B. et al. Amoebic encephalitis due to Sappinia diploidea. JAMA 285, 2450, 2001.
12. Gelman, B.B. et al. Neuropathological and ultrastructural features of amebic encephalitis caused by Sappinia diploidea. J. Neuropathol. Exp. Neurol. 62, 990, 2003.
13. Qvarnstrom, Y. et al. Molecular con�rmation of Sappinia pedata as causative agent of amebic encephalitis. J. Infect. Dis. 199, 1139, 2009.
14. Amaral-Zettler, L.A. et al. A molecular reassessment of the leptomyxid amoebae. Protist 151, 275, 2000.
15. Stothard, D.R. et al. The evolutionary history of the genus Acanthamoeba and the identi�cation of eight new 18S rRNA gene sequence types. J. Eukaryot. Microbiol. 45, 45, 1998.
16. Booton, G.C. et al. Genotyping of Balamuthia mandrillaris based on nuclear 18S and mitochondrial 16S rRNA genes. Am. J. Trop. Med. Hyg. 68, 65, 2003.
17. Booton, G.C. et al. Balamuthia mandrillaris: Identi�cation of clinical and environmental isolates using genus-speci�c PCR. J. Eukaryot. Microbiol. 50 (Suppl), 508–509, 2003.
18. Schuster, F.L. et al. Environmental isolation of Balamuthia mandrillaris associated with a case of amebic encephalitis. J. Clin. Microbiol. 41, 3175, 2003.
19. Dunnebacke, T.H. et al. Balamuthia mandrillaris from soil samples. Microbiology 150, 2837, 2004.
20. Page, F.C. A New Key to Freshwater and Soil Gymnamoebae. Freshwater Biological Association, Ambleside, UK, 1988.
21. Adl, S.M. et al. The new higher level classi�cation of eukaryotes with emphasis on the taxonomy of protists. J. Eukaryot. Microbiol. 52, 399, 2005.
22. Siddiqui, R., Ortega-Rivas, A., and Khan, N.A. Balamuthia mandrillaris resistance to hostile conditions. J. Med. Microbiol. 57, 428, 2008.
23. Siddiqui, R., Khan, N.A., and Jarroll, E.L. The cyst wall carbohydrate composition of Balamuthia mandrillaris. Parasitol. Res. 104, 1439, 2009.
24. Jayasekera, S. et al. Post-mortem culture of Balamuthia mandrillaris from the brain and cerebrospinal �uid of a case of granulomatous amoebic meningoencephalitis, using human brain microvascular endothelial cells. J. Med. Microbiol. 53, 1007, 2004.
25. Kidederlen, A.F. et al. Cytopathogenicity of Balamuthia mandrillaris, an opportunistic causative agent of granulomatous amoebic encephalitis. J. Eukrayot. Microbiol. 53, 456, 2006.
26. Dunnebacke, T.H. The amoeba Balamuthia mandrillaris feeds by entering into mammalian cells in culture. J. Eukaryot. Microbiol. 54, 451, 2007.
27. Visvesvara, G.S. et al. Paravahlkamp�a francinae n. sp. masquearding as an agent of primary amebic meningoencephalitis. J. Eukaryot. Microbiol. 56, 357, 2009.
28. Rocha-Azevedo, B. et al. The interaction between the amoeba Balamuthia mandrillaris and extracellular glycoproteins in vitro. Parasitology 134, 51, 2007.
29. Schuster, F.L., and Visvesvara, G.S. Axenic growth and drug sensitivity studies of Balamuthia mandrillaris, an agent of amebic meningoencephalitis in humans and other animals. J. Clin. Microbiol. 34, 385, 1996.
30. Schuster, F.L. Cultivation of pathogenic and opportunistic free-living amebas. Clin. Microbiol. Rev. 15, 342, 2002. 31. Matin, A. et al. Evaluation of prokaryotic and eukaryotic cells as food source for Balamuthia mandrillaris. Arch. Microbiol. 186, 261, 2006. 32. Niyyati, M. et al. Isolation of Balamuthia mandrillaris from urban dust, free of known infectious involvement. Parasitol. Res. 106, 279, 2009. 33. Visvesvara, G.S. In vitro cultivation of microsporidia of clinical importance. Clin Microbiol. Rev. 15, 401, 2002. 34. Greub, G., and Raoult, D. Microorganisms resistant to freeliving amoebae. Clin. Microbiol. Rev. 17, 413, 2004. 35. Shadrach, W.S. et al. Balamuthia mandrillaris, a free-living ameba and opportunistic agent of encephalitis, is a potential host for Legionella pneumophila bacteria. Appl. Environ. Microbiol. 71, 2244, 2005. 36. Michel, R. et al. Free-living amoebae serve as a host for the Chlamydia-like bacterium Simkania negevensis. Acta Protozool. 44, 113–121, 2005. 37. Hoffmann, E.O. et al. A case of primary amebic meningoencephalitis. Light and electron microscopy and immunohistologic studies. Am. J. Trop. Med. Hyg. 27, 29, 1978. 38. Cox, E.C. Amebic meningoencephalitis caused by Acanthamoeba species in a four month old child. J. S. C. Med. Assoc. 76, 459, 1980. 39. Matson, D.O. et al. Acanthamoeba meningoencephalitis masquerading as neurocysticercosis. Pediatr. Infect. Dis. 7, 121, 1988. 40. Kidney D.D., and Kim, S.H. CNS infections with free-living amebas: Neuroimaging �ndings. Am. J. Radiol. 171, 808, 1988. 41. Anzil, A.P. et al. Amebic meningoencephalitis in a patient with AIDS caused by a newly recognized opportunistic pathogen. Arch. Pathol. 115, 21, 1991. 42. Gordon, S.M. et al. Culture isolation of Acanthamoeba spp. and leptomyxid amebas in AIDS and non-AIDS patients with amebic meningoencephalitis. Clin. Infect. Dis. 15, 1024, 1992. 43. Popek, E.J., and Nea�e, R.C. Cae 4. Granulomatous meningoencephalitis due to leptomyxid ameba. Pediatr. Pathol. 12, 871, 1992. 44. Griesemer, D.A. et al. Amebic meningoencephalitis caused by Balamuthia mandrillaris. Pediatr. Neurol. 10, 249, 1994. 45. Rowen, J.L. et al. Balamuthia mandrillaris: A newly recognized agent for amebic meningoencephalitis. Pediatr. J. Infect. Dis. 14, 705, 1995. 46. Lowichik, A. et al. Leptomyxid amebic meningoencephalitis mimicking brain stem glioma. Am. J. Neuroradiol. 16 (Suppl), 926, 1995. 47. Denney, C.F. et al. Amebic meningoencephalitis caused by Balamuthia mandrillaris: Case report and review. Clin. Infect. Dis. 25, 1354, 1997. 48. Duke, B.J. et al. Balamuthia mandrillaris meningoencephalitis presenting with acute hydrocephalus. Pediatr. Neurol. 26, 107, 1997. 49. Deol, I. et al. Encephalitis due to a free-living amoeba (Balamuthia mandrillaris): Case report with literature review. Surg. Neurol. 53, 611, 2000. 50. Katz, J.D. et al. A case of Balamuthia mandrillaris meningoencephalitis. Arch. Neurol. 57, 1210, 2000. 51. Bakardjiev, A. et al. Amebic encephalitis caused by Balamuthia mandrillaris: Report of four cases. Pediatr. Infect. Dis. 22, 447, 2003. 52. Deetz, T.R. et al. Successful treatment of Balamuthia amoebic encephalitis: Presentation of two cases. Clin. Infect. Dis. 37, 1304, 2003.
53. Jung, S. et al. Balamuthia mandrillaris meningoencephalitis in an immunocompetent patient. Arch. Pathol. Lab. Med. 128, 466, 2004.
54. Rahimian, J., Kleinman, G., and Parta, M. Amebic encephalitis caused by Balamuthia. Infect. Med. 22, 382, 2005.
55. Zagardo, M.T., Castellani, R.J., Zoarski, G.H., and Bauserman, S.C. Granulomatous amebic encephalitis caused by leptomyxid amebae in an HIV-infected patient. Am. J. Neuroradiol. 18, 903, 1997.
56. Pritzker, A.S. et al. Fatal granulomatous amebic encephalitis caused by Balamuthia mandrillaris presenting as a skin lesion. J. Am. Acad. Dermatol. 50, S38–S41, 2004.
57. CDC, Balamuthia amebic encephalitis—California. 1999–20 Mort. Morb. Wkly Rep. 57, 768, 2008.
58. Kansagra, A.P. et al. Balamuthia mandrillaris meningoencephalitis in an immunocompromised patient. J. Neurosurg. 111, 301, 2009.
59. Glaser, C.A. et al. Beyond viruses: Clinical pro�les and etiologies associated with encephalitis. Clin. Infect. Dis. 43, 1565, 2006.
60. Schuster, F.L. et al. Under the radar: Balamuthia amebic encephalitis. Clin. Infect. Dis. 48, 879, 2009.
61. Taratuto, A.L. Leptomyxid amoeba encephalitis: Report of the �rst case in Argentina. Trans. R. Soc. Trop. Med. Hyg. 85, 77, 1991.
62. Galarza, M. et al. Pediatric granulomatous cerebral amebiasis: A delayed diagnosis. Pediatr. Neurol. 26, 153, 2002.
63. Chemelli, L. et al. Granulomatous amoebic encephalitis due to leptomyxid amoebae: Report of the �rst Brazilian case. Trans. Roy. Soc. Trop. Med. Hyg. 86, 635, 1992.
64. Silva-Vergara, M.L. et al. Disseminated Balamuthia mandrillaris amoeba infection in an AIDS patient from Brazil. Am. J. Trop. Med. Hyg. 77, 1096, 2007.
65. Oddó, D., Ciani, S., and Vial, P. Encefalitis amebiana granulomatosa por Balamuthia mandrillaris. Primer caso diagnostico en Chile. Rev. Chil. Infect. 23, 232, 2006.
66. Cuevas, M. et al. Meningoencefalitis granulomatosa por Balamuthia mandrillaris: Reporte de un caso y revisión de la literature. Rev. Chil. Infect. 23, 237, 2006.
67. Riestra-Castaneda, J.M. et al. Granulomatous amebic encephalitis due to Balamuthia mandrillaris (Leptomyxiidae): Report of four cases from Mexico. Am. J. Trop. Med. Hyg. 56, 603, 1997.
68. Uribe-Uribe, N.O. et al. Granulomatous amebic encephalitis by Balamuthia mandrillaris. Patologia 39, 141, 2001.
69. Candanedo-Gonzalez, F., and Gamboa-Domínguez, A. Acute amebic thyroiditis by Balamuthia mandrillaris. Patologia 38, 31–34, 2000.
70. Recavarren-Arce, S. et al. Amoeba angeitic lesions of the central nervous system in Balamuthia mandrillaris amoebiasis. Human Pathol. 30, 269, 1999.
71. Ballona, R., and Aquije, M. Compromiso cutáneo en encefalitis branulomatosa amebiana fatal causada por Balamuthia mandrillaris. Folia Dermatol. 14, 28, 2003.
72. Seas, C.R., and Bravo, F.P. Encefalitis amebiana granulomatosa por Balamuthia mandrillaris: un enfermedad fatal reconocida cada vez más frecuentemente en América Latina. (Editorial) Rev. Chil. Infect. 23, 197, 2006.
73. Valverde, J., Arrese, J.E., and Pierard, G.E. Granulomatous cutaneous centrofacial and meningocerebral amebisis. Am. J. Dermatol. 7, 267, 2006.
74. Bravo, F.G. et al. Cutaneous manifestations of infection by free-living amebas. In: Tyring, S.K., Lupi, O., and Hengge, U.R. (eds), Tropical Dermatology. Elsevier Inc., Philadelphia, pp. 49–55, 2006. 75. Martinez, J. et al. Granulomatous amebic encephalitis: A review and report of a spontaneous case from Venezuela. Acta Neuropathol. 87, 430, 1994. 76. Rodríguez, O. et al. Infección del sistema nervioso central por amebas de vida libre: Comunicación de tres nuevos casos venezolanos. Rev. Neurol. 26, 1005, 1998. 77. Bhgwandeen, S.B. et al. A case of hartmannellid amebic meningoencephalitis in Zambia. Am. J. Clin. Pathol. 63, 483, 1975. 78. Rutherfoord, G.S. Amoebic meningo-encephalitis due to a free-living amoeba. A case report. S. Afr. Med. J. 69, 52, 1986. 79. Carter, R.F. et al. A fatal case of meningoencephalitis due to a free-living amoeba of uncertain identity—Probably Acanthamoeba sp. Pathology 13, 51, 1981. 80. Reed, R.P. et al. Fatal granulomatous amoebic encephalitis caused by Balamuthia mandrillaris. Med. J. Aust. 167, 82, 1997. 81. Kodet, R. et al. Amebic encephalitis caused by Balamuthia mandrillaris in a Czech child: Description of the �rst case from Europe. Pathol. Res. Pract. 194, 423, 1998. 82. White, J.M.L. et al. Granulomatous amoebic encephalitis. Lancet 364, 20, 2004. 83. Bodi, I. et al. Fatal granulomatous amoebic meningoencephalitis due to Balamuthia mandrillaris. Pathol. Res. Practice. 204, 925, 2008. 84. Tavares, M. et al. Diagnosis of �rst case of Balamuthia amebic encephalitis in Portugal by immuno�uorescence and PCR. J. Clin. Microbiol. 44, 2660, 2006. 85. Shirabe, T., Monobe, Y., and Visvesvara, G.S. An autopsy case of amebic meningoencephalitis. The �rst Japanese case caused by Balamuthia mandrillaris. Neuropathology 22, 213, 2002. 86. Prasad, K. et al. Fatal subacute necrotizing brainstem encephalitis in a young man due to a rare parasitic (Balamuthia) infection. Pract. Neurol. 8, 122, 2008. 87. Sangruchi, T. et al. Spontaneous granulomatous amebic encephalitis. Report of 4 cases from Thailand. SouthEast Asian J. Trop. Med. Public Health 25, 309, 1994. 88. Intalpaporn, P. et al. Balamuthia mandrillaris meningoencephalitis: The �rst case in Southeast Asia. Am. J. Trop. Med. Hyg. 70, 666, 2004. 89. Schuster, F.L. et al. Balamuthia amebic encephalitis risk, Hispanic Americans. Emerg. Infect. Dis. 10, 1510, 2004. 90. Chappell, C.I. et al. Standardized method of measuring Acanthamoeba antibodies in sera from healthy subjects. Clin. Diagn. Lab. Immunol. 8, 724, 2001. 91. Foreman, O. et al. Disseminated infection with Balamuthia mandrillaris in a dog. Vet. Pathol. 41, 506, 2004. 92. Healy, J.F. Balamuthia amebic encephalitis: Radiographic and pathologic �ndings. AJNR 23, 486, 2002. 93. Schumacher, D.J., Tien, R.D., and Lane, K. Neuroimaging �ndings in rare amebic infections of the central nervous system. Am. J. Neuroradiol. 16, 930, 1995. 94. Booton, G.C. et al. Identi�cation of Balamuthia mandrillaris by PCR assay using the mitochondrial 16S rRNA gene as a target. J. Clin. Microbiol. 41, 453, 2003. 95. Yagi, S. et al. Detection of Balamuthia 16S rRNA gene DNA in clinical specimens by PCR. J. Clin. Microbiol. 43, 3192, 2005. 96. Qvarnstrom, Y. et al. Multiplex real-time pCR assay for simultaneous detection of Acanthamoeba spp., Balamuthia mandrillaris and Naegleria fowleri. J. Clin. Microbiol. 44, 3589, 2006. 97. Kiderlen, A.F., Radam, E., and Lewin, A. Detection of Balamuthia mandrillaris DNA by real-time PCR targeting the RNase P gene. BMC Microbiol. 8, 210, 2008.
98. Guarner, J. et al. Histopathologic spectrum and immunohistochemical diagnosis of amebic meningoencephalitis. Modern Pathol., 20, 1230, 2007.
99. Matin, A. et al. Balamuthia mandrillaris exhibits metalloprotease activities. FEMS Immunol. Med. Microbiol. 47, 83, 2006.
100. Matin, A., and Khan, N.A. Demonstration and partial characterization of ecto-ATPase in Balamuthia mandrillaris and its possible role in the host–cell interactions. Lett. Appl. Microbiol. 47, 348, 2008.
101. Jayasekera, S. et al. Balamuthia mandrillaris stimulates interleukin-6 release in primary brain microvascular endothelial cells via a phosphatidyliositol 3-kinase-dependent pathway. Microbes Infect. 7, 1345, 2005.
102. Matin, A. et al. Balamuthia mandrillaris interactions with human brain microvascular endothelial cells in vitro. J. Med. Microbiol. 56, 1110, 2007.
103. Matin, A. et al. Effects of human serum on Balamuthia mandrillaris interaction with human brain microvascular endothelial cells. J. Med. Microbiol. 56, 30, 2007.
104. Siddiqui, R. et al. Effect of antimicrobial compounds on Balamuthia mandrillaris encystment and human brain microvascular endothelial cell cytopathogenicity. Antimicrob. Agents Chemother. 51, 4471, 2007.
105. Shadrach, W.S. et al. The pathogenic amoeba Balamuthia mandrillaris possesses cell-associated phospholipase A, lysophospholipase A, and lipase activities. J. Med. Microbiol. 293 (Suppl. 38), 100, 2004.
106. Visvesvara, G.S., Moura, H., and Schuster, F.L. Balamuthia mandrillaris: In vitro cultivation and isoenzyme electrophoresis reveal differences in certain strains. In: Billot-Bonef, S., Cabanes, P.A., Marciano–Cabral, F., Pernin, P., and Pringuez, E. (eds), Proceedings of the IXth International Meeting on the Biology and Pathogenicity of Free-Living Amoebae. John Libbey Eurotext, Montrouge, France, pp. 166–171, 2001.
107. Anderson, M.P. et al. Pneumonia and meningoencephalitis due to amoeba in a lowland gorilla. J. Zool. Anim. Med. 17, 87, 1986.
108. Rideout, B.A. et al. Fatal infections with Balamuthia mandrillaris (a free-living amoeba) in gorillas and other old world primates. Vet. Pathol. 34, 15, 1997.
109. Can�eld, P.J. et al. Amoebic meningoencephalitis in an orang utan (Pongo pygmaeus) caused by Balamuthia mandrillaris. Austral. Vet. Pathol. 75, 97, 1997.
110. Kinde, H. et al. Amebic meningoencephalitis caused by Balamuthia mandrillaris (leptomyxid ameba) in a horse. J. Vet. Diagn. Invest. 10, 378, 1998.
111. Kinde, H. et al. Infections caused by pathogenic free-living amebas (Balamuthia mandrillaris and Acanthamoeba sp.) in horses. J. Vet. Diagn. Invest. 19, 317, 2007.
112. Fuentealba, I.C. et al. Amebic meningoencephalitis in a sheep. JAVMA, 200, 363–365, 1992. 113. Finnin, P.J. et al. Multifocal Balamuthia mandrillaris infection in a dog in Australia. Parasitol. Res. 100, 423, 2006. 114. Janitschke, K. et al. Animal model for Balamuthia mandrillaris CNS infection: Contrast and comparison in immunode�cient and immunocompetent mice: A murine model of “granulomatous’’ amoebic encephalitis. J. Neuropathol. Exp. Neurol. 55, 815, 1996. 115. Kidederlen, A.F., and Laube, U. Balamuthia mandrillaris, an opportunistic agent of encephalitis, infects the brain via the olfactory nerve pathway. Parasitol. Res. 94, 49, 2004. 116. Kiderlen, A.F. et al. Oral infection of immunocompetent and immunode�cient mice with Balamuthia mandrillaris amoebae. Parasitol. Res. 100, 775, 2007. 117. Huang, Z.H., Ferrante, A., and Carter, R.F. Serum antibodies to Balamuthia mandrillaris, a free-living amoeba recently demonstrated to cause granulomatous amoebic encephalitis. J. Infect. Dis. 179, 1305, 1999. 118. Schuster, F.L. et al. Detection of antibodies against free-living amoebae Balamuthia mandrillaris and Acanthamoeba species in a population of patients with encephalitis. Clin. Infect. Dis. 42, 1260, 2006. 119. Schuster, F.L. et al. Balamuthia mandrillaris, agent of amebic encephalitis: Detection of serum antibodies and antigenic similarity of isolates by enzyme immunoassay. J. Eukaryot. Microbiol. 55, 313, 2008. 120. Schuster, F.L. et al. Survey of sera from encephalitis patients for Balamuthia mandrillaris antibody. J. Eukaryot. Microbiol. IWOP: 10S, 2001. 121. Schuster, F.L., and Visvesvara, G.S. Ef�cacy of novel antimicrobials against clinical isolates of opportunistic amebas. J. Eukaryot. Microbiol. 45, 612, 1998. 122. Schuster, F.L., and Visvesvara, G.S. Opportunistic amoebae: Challenges in prophylaxis and treatment. Drug Resist. Updates 7, 41, 2004. 123. Schuster, F.L., Guglielmo, B.J., and Visvesvara, G.S. In-vitro activity of miltefosine and voriconazole on clinical isolates of free-living amoebas: Balamuthia mandrillaris, Acanthamoeba spp., and Naegleria fowleri. J. Eukaryot. Microbiol. 53, 121, 2006. 124. Yagi, S., Schuster, F.L., and Visvesvara, G.S. Demonstration of Balamuthia and Acanthamoeba mitochondrial DNA in sectioned archival brain and other tissues by the polymerase chain reaction. Parasitol. Res. 102, 211, 2008. 125. Greer, C.E., Lund, J.K., and Manos, M.M. PCR ampli�cation from paraf�n-embedded tissues; recommendation on �xatives for long-term storage and prospective studies. PCR Methods Appl. 1, 46, 1991. 126. Kiderlen, A.F., Radam, E., and Tata, P.S. Assessment of Balamuthia mandrillaris-speci�c serum antibody concentrations by �ow cytometry. Parasitol. Res. 104, 663, 2009. 4 Chapter 4 - Blastocystis
1. Brumpt, E. Blastocystis hominis n sp. et formes voisines. Bull. Soc. Pathol. Exot. 5, 725, 1912.
2. Zierdt, C.H. Blastocystis hominis—past and future. Clin. Microbiol. Rev. 4, 61, 1991.
3. Boreham, P.F.L., and Stenzel, D.J. Blastocystis in humans and animals: Morphology, biology, and epizootiology, Adv. Parasitol. 32, 1, 1993.
4. Stenzel, D.J., and Boreham, P.F.L. Blastocystis hominis revisited. Clin. Microbiol. Rev. 9, 563, 1996.
5. Tan, K.S.W., Singh, M., and Yap, E.H. Recent advances in Blastocystis hominis research: Hot spots in terra incognita. Int. J. Parasitol. 32, 789, 2002.
6. Tan, K.S.W. Blastocystis in humans and animals: New insights using modern methodologies. Vet. Parasitol. 126, 121, 2004.
7. Tan, K.S.W. New insights on classi�cation, identi�cation, and clinical relevance of Blastocystis spp. Clin. Microbiol. Rev. 21, 639, 2008.
8. Amin, O.M. Seasonal prevalence of intestinal parasites in the United States during 2000. Am. J. Trop. Med. Hyg. 66, 799, 2002.
9. Cirioni, O. et al. Prevalence and clinical relevance of Blastocystis hominis in diverse patient cohorts. Eur. J. Epidemiol. 15, 389, 1999.
10. Noor Azian, M.Y. et al. Prevalence of intestinal protozoa in an aborigin community in Pahang, Malaysia. Trop. Biomed. 24, 55, 2007.
11. Windsor, J.J. et al. Incidence of Blastocystis hominis in faecal samples submitted for routine microbiological analysis. Br. J. Biomed. Sci. 59, 154, 2002.
12. Abe, N. et al. A survey of Blastocystis sp. in livestock, pets, and zoo animals in Japan. Vet. Parasitol. 106, 203, 2002.
13. Chen, X.Q. et al. A survey of Blastocystis sp. in rodents. Lab. Anim. Sci. 47, 91, 1997. 14. Cordón, G.P. et al. Intestinal parasitism in the animals of the zoological garden “Peña Escrita” (Almuñecar, Spain). Vet. Parasitol. 156, 302, 2008.
15. Duda, A., Stenzel, D.J., and Boreham, P.F. Detection of Blastocystis sp. in domestic dogs and cats. Vet. Parasitol. 76, 9, 1998.
16. Jones-Engel, L. et al. Prevalence of enteric parasites in pet macaques in Sulawesi, Indonesia. Am. J. Primatol. 62, 71, 2004.
17. Lee, M.G., and Stenzel, D.J. A survey of Blastocystis in domestic chickens. Parasitol. Res. 85, 109, 1999.
18. Legesse, M., and Erko, B. Zoonotic intestinal parasites in Papio anubis (baboon) and Cercopithecus aethiops (vervet) from four localities in Ethiopia. Acta Trop. 90, 231, 2004.
19. Pakandl, M. Occurrence of Blastocystis sp. in pigs. Folia Parasitol. 38, 297, 1991.
20. Ponce Gordo, F. et al. Parasites from farmed ostriches (Struthio camelus) and rheas (Rhea americana) in Europe. Vet. Parasitol. 107, 137, 2002. 21. Teow, W.L. et al. A survey of Blastocystis in reptiles. Parasitol. Res. 78, 453, 1992. 22. Yamada, M. et al. Light microscopical study of Blastocystis spp. in monkeys and fowls. Parasitol. Res. 73, 527, 1987. 23. Yoshikawa, H. et al. A survey of Blastocystis infection in anuran and urodele amphibians. Vet. Parasitol. 122, 91, 2004. 24. Yoshikawa, H. et al. Ultrastructural and phylogenetic studies on Blastocystis isolates from cockroaches. J. Eukaryot. Microbiol. 54, 33, 2007. 25. Zaman, V. et al. Isolation of Blastocystis from the cockroach (Dictyoptera: Blattidae). Parasitol. Res. 79, 73, 1993. 26. Abe, N., Wu, Z., and Yoshikawa, H. Molecular characterization of Blastocystis isolates from birds by PCR with diagnostic primers and restriction fragment length polymorphism analysis of the small subunit ribosomal RNA gene. Parasitol. Res. 89, 393, 2003. 27. Abe, N., Wu, Z., and Yoshikawa, H. Zoonotic genotypes of Blastocystis hominis detected in cattle and pigs by PCR with diagnostic primers and restriction fragment length polymorphism analysis of the small subunit ribosomal RNA gene. Parasitol. Res. 90, 124, 2003. 28. Arisue, N., Hashimoto, T., and Yoshikawa, H. Sequence heterogeneity of the small subunit ribosomal RNA genes among Blastocystis isolates. Parasitology 126, 1, 2003. 29. Clark, C.G. Extensive genetic diversity in Blastocystis hominis. Mol. Biochem. Parasitol. 87, 79, 1997. 30. Hoevers, J. et al. Restriction-fragment-length polymorphism analysis of small-subunit rRNA genes of Blastocystis hominis isolates from geographically diverse human hosts. Parasitol. Res. 86, 57, 2000. 31. Snowden, K. et al. Restriction-fragment-length polymorphism analysis of small-subunit rRNA genes of Blastocystis isolates from animal hosts. Parasitol. Res. 86, 62, 2000. 32. Yoshikawa, H. et al. DNA polymorphism revealed by arbitrary primers polymerase chain reaction among Blastocystis strains isolated from humans, a chicken, and a reptile. J. Eukaryot. Microbiol. 43, 127, 1996. 33. Abe, N. Molecular and phylogenetic analysis of Blastocystis isolates from various hosts. Vet. Parasitol. 120, 235, 2004. 34. Abe, N., Wu, Z., and Yoshikawa, H. Molecular characterization of Blastocystis isolates from primates. Vet. Parasitol. 113, 321, 2003. 35. Noël, C. et al. Phylogenetic analysis of Blastocystis isolates from different hosts based on the comparison of small-subunit rRNA gene sequences. Mol. Biochem. Parasitol. 126, 119, 2003. 36. Noël, C. et al. Molecular phylogenies of Blastocystis isolates from different hosts: Implications for genetic diversity, identi�cation of species, and zoonosis. J. Clin. Microbiol. 43, 348, 2005. 37. Yoshikawa, H. et al. Problems in speciation in the genus Blastocystis. Trends Parasitol. 20, 251, 2004. 38. Stensvold, C.R. et al. Terminology for Blastocystis subtypes— a consensus. Trends Parasitol. 23, 93, 2007. 39. Stensvold, C.R. et al. Symptomatic infection with Blastocystis sp. subtype 8 successfully treated with trimethoprim-sulfamethoxazole. Ann. Trop. Med. Parasitol. 102, 271, 2008. 40. Yan, Y. et al. Blastocystis sp. subtype 5: A possibly zoonotic genotype. Parasitol. Res. 101, 1527, 2007. 41. Yoshikawa, H. et al. Polymerase chain reaction-based genotype classi�cation among human Blastocystis hominis populations isolated from different countries. Parasitol. Res. 92, 22, 2004. 42. Yoshikawa, H., and Oishi, K. Ultrastructural localization of basic proteins of Blastocystis hominis. Protoplasma 200, 31, 1997.
43. Yoshikawa, H., Kuwayama, N., and Enose, Y. Histochemical detection of carbohydrates of Blastocystis hominis. J. Eukaryot. Microbiol. 42, 70, 1995.
44. Yoshikawa, H., Satoh, J., and Enose, Y. Light and electron microscopic localization of lipids in Blastocystis hominis. J. Electron Microsc. 44, 100, 1996.
45. Stenzel, D.J., Dunn, L.A., and Boreham, P.F.L. Endocytosis in cultures of Blastocystis hominis. Int. J. Parasitol. 19, 787, 1989.
46. Stenzel, D.J., Boreham, P.F.L., and McDougall, R. Ultrastructure of Blastocystis hominis in human stool samples. Int. J. Parasitol. 21, 807, 1991.
47. Chen, X.Q. et al. Description of a Blastocystis species from Rattus norvegicus. Parasitol. Res. 83, 313, 1997.
48. Dunn, L.A., Boreham, P.F.L., and Stenzel, D.J. Ultrastructural variation of Blastocystis hominis stocks in culture. Int. J. Parasitol. 19, 43, 1989.
49. Tan, T.C., and Suresh, K.G. Predominance of amoeboid forms of Blastocystis hominis in isolates from symptomatic patients. Parasitol. Res. 98, 189, 2006.
50. Tan, T.C., and Suresh, K.G. Amoeboid form of Blastocystis hominis—a detailed ultrastructural insight. Parasitol. Res. 99, 737, 2006.
51. Moe, K.T. et al. Observations on the ultrastructure and viability of the cystic stage of Blastocystis hominis from human feces. Parasitol. Res. 82, 439, 1996.
52. Stenzel, D.J., and Boreham, P.F. A cyst-like stage of Blastocystis hominis. Int. J. Parasitol. 21, 613, 1991.
53. Yoshikawa, H. et al. Freeze-fracture and cytochemical studies on the in vitro cyst form of reptilian Blastocystis pythoni. J. Eukaryot. Microbiol. 50, 70, 2003.
54. Zaman, V., Howe, J., and Ng, M. Ultrastructure of Blastocystis hominis cysts. Parasitol. Res. 81, 465, 1995.
55. Zaman, V. The diagnosis of Blastocystis hominis cysts in human faeces. J. Infect. 33, 15, 1996.
56. Zaman, V., and Khan, K.Z. A concentration technique for obtaining viable cysts of Blastocystis hominis from faeces. J. Pak. Med. Assoc. 44, 220, 1994.
57. Pakandl, M. Blastocystis sp. from pigs: Ultrastructural changes occurring during polyxenic cultivation in Iscove’s modi�ed Dulbecco’s medium. Parasitol. Res. 85, 743, 1999.
58. Yoshikawa, H., and Hayakawa, A. Morphological changes in the central vacuole of Blastocystis hominis during in vitro culture. Protoplasma 194, 63, 1996.
59. Yoshikawa, H., Yamada, M., and Yoshida, Y. Freeze-fracture study of Blastocystis hominis. J. Protozool. 35, 522, 1988.
60. Zierdt, C.H. Studies of Blastocystis hominis. J. Protozool. 20, 114, 1973.
61. Burghelea, B., and Radulescu, S. Ultrastructural evidence for a possible differentiation way in the life-cycle of Blastocystis hominis, Roum. Arch. Microbiol. Immunol. 50, 231, 1991.
62. Govind, S.K., Khairul, A.A., and smith, H.V. Multiple reproductive processes in Blastocystis. Trends Parasitol. 18, 528, 2002.
63. Singh, M. et al. Elucidation of the life cycle of the intestinal protozoan Blastocystis hominis. Parasitol. Res. 81, 446, 1995.
64. Tan, K.S.W., and Stenzel, D.J. Multiple reproductive processes in Blastocystis: Proceed with caution. Trends Parasitol. 19, 290, 2003.
65. Tan, T.C., and Suresh, K.G. Evidence of plasmotomy in Blastocystis hominis. Parasitol. Res. 101, 1521, 2007.
66. Zhang, X. et al. Morphology and reproductive mode of Blastocystis hominis in diarrhea and in vitro. Parasitol. Res. 101, 43, 2007. 67. Suresh, K. et al. Sac-like pouches in Blastocystis from the house lizard Cosymbotus platyurus. Parasitol. Res. 83, 523, 1997. 68. Silberman, J.D. et al. Human parasite �nds taxonomic home. Nature 380, 398, 1996. 69. Arisue, N. et al. Phylogenetic position of Blastocystis hominis and of stramenopiles inferred from multiple molecular sequence data. J. Eukaryot. Microbiol. 49, 42, 2002. 70. Patterson, D.J. Stramenopiles: Chromophytes from a protistan perspective. In: Green, J.C., Leadbeater, B.S.C., and Diver, W.L. (ed.), The Chromophyte Algae: Problems and Perspectives. Oxford, Clarendon Press, pp. 357–379, 1989. 71. Leipe, D.D. et al. 16S-like rDNA sequences from Developayella elegans, Labyrinthuloides haliotidis, and Proteromonas lacertae con�rm that the stramenopiles are a primarily heterotrophic group. Europ. J. Protistol. 32, 449, 1996. 72. Basualdo, J.A. et al. Intestinal parasitoses and environmental factors in a rural population of Argentina 2002–2003. Rev. Inst. Med. Trop. S. Paulo 49, 251, 2007. 73. Minvielle, M.C. et al. Epidemiological survey of Giardia spp. and Blastocystis hominis in an Argentinian rural community. Korean J. Parasitol. 42, 121, 2004. 74. Rayan, H.Z., Ismail, O.A., and El Gayar, E.K. Prevalence and clinical feature of Dientamoeba fragilis infections in patients suspected to have intestinal parasitic infection. J. Egypt. Soc. Parasitol. 37, 599, 2007. 75. Aguiar, J.I.A. et al. Intestinal protozoa and helminths among Terena Indians in the State of Mato Grosso do Sul: High prevalence of Blastocystis hominis. Rev. Soc. Bras. Med. Trop. 40, 631, 2007. 76. Graczyk, T.K. et al. The association of Blastocystis hominis and Endolimax nana with diarrheal stools in Zambian schoolage children. Parasitol. Res. 98, 38, 2005. 77. Arani, A.S. et al. Prevalence of intestinal parasites in a population in south of Teheran, Iran. Rev. Inst. Med. Trop. S. Paulo 50, 145, 2008. 78. Wong, K.H. et al. Predominance of subtype 3 among Blastocystis isolates from a major hospital in Singapore. Parasitol. Res. 102, 663, 2008. 79. Navarro, C. et al. High prevalence of Blastocystis sp. in pigs reared under intensive growing systems: frequency of ribotypes and associated risk factors. Vet. Parasitol. 153, 347, 2008. 80. Burden, D.J., Anger, H.S., and Hammet, N.C. Blastocystis sp. infection in pigs. Vet. Microbiol. 3, 227, 1978/1979. 81. Quílez, J. et al. Occurrence of Blastocystis sp. in cattle in Aragón, northeastern Spain. Parasitol. Res. 81, 703, 1995. 82. Lim, Y.A.L. et al. Intestinal parasites in various animals at a zoo in Malaysia. Vet. Parasitol. 157, 154, 2008. 83. Scicluna, S.M., Tawari, B., and Clark, C.G. DNA barcoding of Blastocystis Protist 157, 77, 2006. 84. Stensvold, C.R. et al. Blastocystis: unravelling potential risk factors and clinical signi�cance of a common but neglected parasite. Epidemiol. Infect. 137, 1655, 2009. 85. Thathaisong, U. et al. Blastocystis isolates from a pig and a horse are closely related to Blastocystis hominis. J. Clin. Microbiol. 41, 967, 2003. 86. Parkar, U. et al. Direct characterization of Blastocystis from faeces by PCR and evidence of zoonotic potential. Parasitology 134, 359, 2007. 87. Rivera, W.L. Phylogenetic analysis of Blastocystis isolates from animal and human hosts in the Philippines. Vet. Parasitol. 156, 178, 2008.
88. Yoshikawa, H. et al. Molecular comparative studies among Blastocystis isolates obtained from humans and animals. J. Parasitol. 89, 585, 2003.
89. Yoshikawa, H., Abe, N., and Wu, Z. PCR-based identi�cation of zoonotic isolates of Blastocystis from mammals and birds. Microbiology 150, 1147, 2004. 90. Dogruman-Al, F. et al. A possible link between subtype 2 and asymptomatic infections of Blastocystis hominis. Parasitol. Res. 103, 685, 2008.
91. Dogruman-Al, F. et al. Blastocystis subtypes in irritable bowel syndrome and in�ammatory bowel diseases in Ankara, Turkey. Mem. Inst. Oswald Cruz 104, 724, 2009.
92. Li, L.H. et al. Molecular epidemiology of human Blastocystis in a village in Yunnan province, China. Parasitol. Int. 56, 281, 2007.
93. Li, L.H. et al. Cross-sectional surveys and subtype classi�cation of human Blastocystis isolates from four epidemiological settings in China. Parasitol. Res. 102, 83, 2007.
94. Özyurt, M. et al. Molecular epidemiology of Blastocystis infections in Turkey. Parasitol. Int. 57, 300, 2008.
95. Stensvold, C.R. et al. Blastocystis: Subtyping isolates using pyrosequencing™ technology. Exp. Parasitol. 116, 111, 2007.
96. Stensvold, C.R. et al. Detecting Blastocystis using parasitologic and DNA-based methods: A comparative study. Diagn. Microbiol. Infect. Dis. 59, 303, 2007.
97. Souppart, L. et al. Molecular epidemiology of human Blastocystis isolates in France. Parasitol. Res. 105, 413, 2009.
98. Leelayoova, S. et al. Drinking water: A possible source of Blastocystis spp. subtype 1 infection in schoolchildren of a rural community in central Thailand. Am. J. Trop. Med. Hyg. 79, 401, 2008.
99. Stensvold, C.R. et al. Subtype distribution of Blastocystis isolates from synanthropic and zoo animals and identi�cation of a new subtype. Int. J. Parasitol. 39, 473, 2009.
100. Khan, Z.A., and Alkhalife, I.S. Prevalence of Blastocystis hominis among “healthy” food handlers in Dammam, Soudi Arabia. J. Egypt. Soc. Parasitol. 35, 395, 2005.
101. Sadek, Y. et al. Intestinal parasites among food-handlers in Qualyobia Governorate, with reference to the pathogenic parasite Blastocystis hominis. J. Egypt. Soc. Parasitol. 27, 471, 1997.
102. Salim, H.R. et al. Blastocystis in animal handlers. Parasitol. Res. 85, 1032, 1999.
103. Yoshikawa, H. et al. Molecular characterization of Blastocystis isolates from children and rhesus monkeys in Kathmandu, Nepal. Vet. Parasitol. 160, 295, 2009.
104. Iguchi, A. et al. Infectivity of different genotypes of human Blastocystis hominis isolates in chickens and rats. Parasitol. Int. 56, 107, 2007.
105. Yoshikawa, H. et al. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J. Clin. Microbiol. 38, 1324, 2000.
106. Stark, D. et al. Prevalence of enteric protozoa in human immunode�ciency virus (HIV)-positive and HIV-negative men who have sex with men from Sydney, Australia. Am. J. Trop. Med. Hyg. 76, 549, 2007.
107. Rene, B.A. et al. Subtype analysis of Blastocystis isolates from Blastocystis cyst excreting patients. Am. J. Trop. Med. Hyg. 80, 588, 2009.
108. Yoshikawa, H. et al. Fecal–oral transmission of the cyst form of Blastocystis hominis in rats. Parasitol. Res. 94, 391, 2004.
109. Tanizaki, A. et al. Infectivity of Blastocystis isolates from chickens, quails and geese in chickens. Parasitol. Res. 96, 57, 2005. 110. Suresh, K., Smith, H.V., and Tan, T.C. Viable Blastocystis cysts in Scottish and Malaysian sewage samples. Appl. Environ. Microbiol. 71, 5619, 2005. 111. Leelayoova, S. et al. Evidence of waterborne transmission of Blastocystis hominis. Am. J. Trop. Med. Hyg. 70, 658, 2004. 112. Nimri, L., and Batchoun, R. Intestinal colonization of symptomatic and asymptomatic schoolchildren with Blastocystis hominis. J. Clin. Microbiol. 32, 2865, 1994. 113. Taamasri, P. et al. Transmission of intestinal blastocystosis related to the quality of drinking water. Southeast Asian J. Trop. Med. Public Health 31, 112, 2000. 114. Stensvold, C.R. et al. Pursuing the clinical signi�cance of Blastocystis—diagnostic limitations. Trends Parasitol. 25, 23, 2009. 115. Carrascosa, M., Martínez, J., and Pérez-Castrillon, J.L. Hemorrhagic proctosigmoiditis and Blastocystis hominis infection. Ann. Intern. Med. 124, 278, 1996. 116. Chen, T.L. et al. Clinical characteristics and endoscopic �ndings associated with Blastocystis hominis in healthy adults. Am. J. Trop. Med. Hyg. 69, 213, 2003. 117. Kaya, S. et al. Pathogenicity of Blastocystis hominis, a clinical reevaluation. Turkiye Parazitol. Derg. 31, 184, 2007. 118. Kuo, H-Y. et al. Clinical signi�cance of Blastocystis hominis: Experience from a medical center in northern Taiwan. J. Microbiol. Immunol. Infect. 41, 222, 2008. 119. Leder, K. et al. No correlation between clinical symptoms and Blastocystis hominis in immunocompetent individuals. J. Gastroenterol. Hepatol. 20, 1390, 2005. 120. Sun, T. et al. Questionable clinical signi�cance of Blastocystis hominis infection. Am. J. Gastroenterol. 84, 1543, 1989. 121. Zaki, M. et al. Clinical report of Blastocystis hominis infection in children. J. Trop. Med. Hyg. 94, 118, 1991. 122. Qadri, S.M.H., Al-Okaili, G.A., and Al-Dayel, F. Clinical signi�cance of Blastocystis hominis. J. Clin. Microbiol., 27, 2407, 1989. 123. Doyle, P.W. et al. Epidemiology and pathogenicity of Blastocystis hominis. J. Clin. Microbiol. 28, 116, 1990. 124. Markell, E.K., and Udkow, M.P. Association of Blastocystis hominis with human disease? J. Clin. Microbiol. 28, 1085, 1990. 125. Rosenblatt, J.E. Blastocystis hominis. J. Clin. Microbiol. 28, 2379, 1990. 126. Zierdt, C.H. Pathogenicity of Blastocystis hominis. J. Clin. Microbiol. 29, 662, 1991. 127. Lee, M.J. Pathogenicity of Blastocystis hominis. J. Clin. Microbiol. 29, 2089, 1991. 128. Boorom, K.F. et al. Oh my aching gut: Irritable bowel syndrome, Blastocystis, and asymptomatic infection. Parasite Vector 1, 40, 2008. 129. Giacometti, A. et al. Irritable bowel syndrome in patients with Blastocystis hominis infection. Eur. J. Clin. Microbiol. Infect. Dis. 18, 436, 1999. 130. Stark, D. et al. Irritable bowel syndrome: A review on the role of intestinal protozoa and the importance of their detection and diagnosis. Int. J. Parasitol. 37, 11, 2007. 131. Yakoob, J. et al. Irritable bowel syndrome in search of an etiology: Role of Blastocystis hominis. Am. J. Trop. Med. Hyg. 70, 383, 2004. 132. Tungtrongchitr, A. et al. Blastocystis hominis infection in irritable bowel syndrome patients. Southeast Asian J. Trop. Med. Pub. Health 35, 705, 2004. 133. Eroglu, F. et al. Identi�cation of Blastocystis hominis isolates from asymptomatic and symptomatic patients by PCR. Parasitol. Res. 105, 1589, 2009.
134. Hussein, E.M. et al. Pathophysiological variability of different genotypes of human Blastocystis hominis Egyptian isolates in experimentally infected rats. Parasitol. Res. 102, 853, 2008.
135. Yan, Y. et al. Genetic variability of Blastocystis hominis isolates in China. Parasitol. Res. 99, 597, 2006.
136. Tan, T.C., Suresh, K.G., and Smith, H.V. Phenotypic and genotypic characterization of Blastocystis hominis isolates implicates subtype 3 as a subtype with pathogenic potential. Parasitol. Res. 104, 85, 2008.
137. Kaneda, Y. et al. Ribodemes of Blastocystis hominis isolates in Japan. Am. J. Trop. Med Hyg. 65, 393, 2001.
138. Böhm-Gloning, B., Knobloch, J., and Walderich, B. Five subgroups of Blastocystis hominis from symptomatic and asymptomatic patients revealed by restriction site analysis of PCR-ampli�ed 16S-like rDNA. Trop. Med. Int. Health 2, 771, 1997.
139. Dogruman-Al, F. et al. PCR-based subtyping of Blastocystis isolates from symtomatic and asymptomatic individuals in a major hospital in Ankara, Turkey. Parasitol. Res. 106, 263, 2009.
140. Motazedian, H., Ghasemi, H., and Sadjjadi, S.M. Genomic diversity of Blastocystis hominis from patients in southern Iran. Ann. Trop. Med. Parasitol. 102, 85, 2008.
141. Leelayoova, S. et al. In-vitro cultivation: A sensitive method for detecting Blastocystis hominis. Ann. Trop. Med. Parasitol. 96, 803, 2002.
142. Suresh, K., and Smith, H. Comparison of methods for detecting Blastocystis hominis. Eur. J. Clin. Microbiol. Infect. Dis. 23, 509, 2004.
143. Termmathurapoj, S. et al. The usefulness of short-term in vitro cultivation for the detection and molecular study of Blastocystis hominis in stool specimens. Parasitol. Res. 93, 445, 2004.
144. Hussain, R. et al. Signi�cantly increased IgG2 subclass antibody levels to Blastocystis hominis in patients with irritable bowel syndrome. Am. J. Trop. Med. Hyg. 56, 301, 1997.
145. Mahmoud, M.S.E., and Saleh, W.A. Secretory and humoral antibody responses to Blastocystis hominis in symptomatic and asymptomatic human infections. J. Egypt. Soc. Parasitol. 33, 13, 2003. 146. Zierdt, C.H., Zierdt, W.S., and Nagy, B. Enzyme-linked immunosorbent assay for detection of serum antibody to Blastocystis hominis in symptomatic infections. J. Parasitol. 81, 127, 1995.
147. Kaneda, Y. et al. Serologic response to Blastocystis hominis infection in asymptomatic individuals. Tokai J. Exp. Clin. Med. 25, 51, 2000.
148. Garavelli, P.L. et al. Serum antibody detected by �uorescent antibody test in patients with symptomatic Blastocystis hominis infection. Recenti Prog. Med. 86, 398, 1995. 149. Chen, J. et al. Lack of serum immune response to Blastocystis hominis Lancet 330, 1021, 1987. 150. Kukoschke, K.G. and Müller, H.E., SDS–PAGE and immunological analysis of different axenic Blastocystis hominis strains. J. Med. Microbiol. 35, 35, 1991. 151. Lanuza, M.D. et al. Soluble-protein and antigenic heterogeneity in axenic Blastocystis hominis isolates: Pathogenic implications. Parasitol. Res. 85, 93, 1999. 152. Müller, H.E. Four serologically different groups within the species Blastocystis hominis. Zbl. Bakt. 280, 403, 1994. 153. Yoshikawa, H. et al. Antigenic differences between Blastocystis hominis and Blastocystis sp. revealed by polyclonal and monoclonal antibodies. J. Protozool. Res. 5, 118, 1995. 154. Grabensteiner, E., and Hess, M. PCR for the identi�cation and differentiation of Histomonas meleagridis, Tetratrichomonas gallinarum and Blastocystis spp. Vet. Parasitol. 142, 223, 2006. 155. Stensvold, R. et al. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J. Parasitol. 92, 1081, 2006. 156. Utzinger, J. et al. Microscopic diagnosis of sodium acetate– acetic acid–formalin-�xed stool samples for helminths and intestinal protozoa: A comparison among European reference laboratories. Clin. Microbiol. Infect. 16, 267, 2010. 157. Abe, N., and Yoshikawa, H. Stool examination and diagnosis of recently attracted intestinal parasitic protozoa Blastocystis hominis. Rinsho Kensa 41, 938 (1997) (in Japanese). 158. Boeck, W.C., and Drbohlav, J. The cultivation of Entamoeba histolytica. Am. J. Hyg. 5, 371, 1925. 159. Clark, C.G., and Diamond, L.S, Methods for cultivation of luminal parasitic protists of clinical importance. Clin. Microbiol. Rev. 15, 329, 2002. 160. Rivera, W.L., and Tan, M.A. Molecular characterization of Blastocystis isolates in the Philippines by riboprinting. Parasitol. Res. 96, 253, 2005. 161. Yoshikawa, H. et al. Genomic polymorphism among Blastocystis hominis strains and development of subtype-speci�c diagnostic primers. Mol. Cell. Probes 12, 153, 1998. 162. Menounos, P.G. et al. Direct detection of Blastocystis sp. in human faecal samples and subtype assignment using single strand conformational polymorphism and sequencing. Mol. Cell. Probes 22, 24, 2008. 163. Jones II, M.S. et al. Detection of Blastocystis from stool samples using real-time PCR. Parasitol. Res. 103, 551, 2008. 164. Yoshikawa et al. Evaluation of DNA extraction kits for molecular diagnosis of human Blastocystis subtypes from fecal samples. Rarasitol. Res. 109, 1045, 2011. 5 Chapter 5 - Dientamoeba
1. Jepps, M.W., and Dobell, C. Dientamoeba fragilis n.g., n. sp., new intestinal amoeba from man. Parasitology 10, 352–367, 1918.
2. Delgado-Viscogliosi, P. et al. Molecular phylogeny of parabasalids based on small subunit rRNA sequences, with emphasis on the Trichomonadinae subfamily. J. Eukaryot. Microbiol. 47, 70–75, 2000. 3. Dobell, C. Researches on the intestinal protozoa of monkeys and man. X. The life history of Dientamoeba fragilis: Observations, experiments, and speculations. Parasitology 32, 417–461, 1940. 4. Dwyer, D.M. Analysis of the antigenic relationships among Trichomonas, Histomonas, Dientamoeba, and Entamoeba. II. Gel diffusion methods. J. Protozool 19, 326–332, 1972. 5. Dwyer, D.M. Analysis of the antigenic relationships among Trichomonas, Histomonas, Dientamoeba, and Entamoeba. I. Quantitative �uorescent antibody methods. J. Protozool 19, 316–325, 1972. 6. Dwyer, D.M. Analysis of the antigenic relationships among Trichomonas, Histomonas, Dientamoeba and Entamoeba. 3. Immunoelectrophoresis technics. J. Protozool 21, 139–145, 1974. 7. Camp, R.R., Mattern, C.F. and Honigberg, B.M. Study of Dientamoeba fragilis Jepps & Dobell. I. Electron microscopic observations of the binucleate stages. II. Taxonomic position and revision of the genus. J. Protozool 21, 69–82, 1974. 8. Bird, R.G., Sargeaunt, P., and Upton, C.P. Uni- and binucleate trophozoites of Dientamoeba fragilis. Trans. R. Soc. Trop. Med. Hyg. 64, 18, 1970. 9. Silberman, J.D., Clark, C.G., and Sogin, M.L. Dientamoeba fragilis shares a recent common evolutionary history with the trichomonads. Mol. Biochem. Parasitol. 76, 311–314, 1996. 10. Gerbod, D. et al. Phylogenetic position of the trichomonad parasite of turkeys, Histomonas meleagridis (Smith) Tyzzer, inferred from small subunit rRNA sequence. J. Eukaryot. Microbiol. 48, 498–504, 2001. 11. Stark, D.J., Beebe, N., Marriott, D., Ellis, J.T., and Harkness, J. Dientamoebiasis: Clinical importance and recent advances. Trends Parasitol. 22, 92–96, 2006. 12. Windsor, J.J., and Johnson, E.H. Dientamoeba fragilis: The un�agellated human �agellate. Br. J. Biomed. Sci. 56, 293–306, 1999. 13. Wenrich, D.H. Studies on Dientamoeba fragilis (protozoa). IV. Further observations, with an outline of present-day knowledge of this species. J. Parasitol. 30, 322–337, 1944. 14. Johnson, E.H., Windsor, J.J., and Clark, C.G. Emerging from obscurity: Biological, clinical, and diagnostic aspects of Dientamoeba fragilis. Clin. Microbiol. Rev. 17, 553–570, table of contents, 2004. 15. Burrows, R.B., Swerdlow, M.A., Frost, J.K., and Leeper, C.K. Pathology of Dientamoeba fragilis infections of the appendix. Am. J. Trop. Med. Hyg. 3, 1033–1039, 1954. 16. Swerdlow, M.A., and Burrows, R.B. Dientamoeba fragilis, an intestinal pathogen. J. Am. Med. Assoc. 158, 176–178, 1955. 17. Burrows, R.B., and Swerdlow, M.A. Enterobius vermicularis as a probable vector of Dientamoeba fragilis. Am. J. Trop. Med. Hyg. 5, 258–265, 1956. 18. Girginkardesler, N., Kurt, O., Kilimcioglu, A.A., and Ok, U.Z. Transmission of Dientamoeba fragilis: Evaluation of the role of Enterobius vermicularis. Parasitol. Int. 57, 72–75, 2008. 19. Yang, J., and Scholten, T. Dientamoeba fragilis: A review with notes on its epidemiology, pathogenicity, mode of transmission, and diagnosis. Am. J. Trop. Med. Hyg. 26, 16–22, 1977. 20. Kean, B.H., and Malloch, C.L. The neglected ameba: Dientamoeba fragilis. A report of 100 “pure” infections. Am. J. Dig. Dis. 11, 735–746, 1966. 21. Cuffari, C., Oligny, L., and Seidman, E.G. Dientamoeba fragilis masquerading as allergic colitis. J. Pediatr. Gastroenterol. Nutr. 26, 16–20, 1998.
22. Stark, D., Beebe, N., Marriott, D., Ellis, J., and Harkness, J. Prospective study of the prevalence, genotyping, and clinical relevance of Dientamoeba fragilis infections in an Australian population. J. Clin. Microbiol. 43, 2718–2723, 2005.
23. Windsor, J.J., Rafay, A.M., Shenoy, A.K., and Johnson, E.H. Incidence of Dientamoeba fragilis in faecal samples submitted for routine microbiological analysis. Br. J. Biomed. Sci. 55, 172–175, 1998.
24. Ayadi, A., and Bahri, I. Dientamoeba fragilis: Pathogenic �agellate?. Bull. Soc. Pathol. Exot. 92, 299–301, 1999.
25. Johnson, J.A., and Clark, C.G. Cryptic genetic diversity in Dientamoeba fragilis. J. Clin. Microbiol. 38, 4653–4654, 2000.
26. Peek, R., Reedeker, F.R., and van Gool, T. Direct ampli�cation and genotyping of Dientamoeba fragilis from human stool specimens. J. Clin. Microbiol. 42, 631–635, 2004.
27. Windsor, J.J., Clark, C.G., and Macfarlane, L. Molecular typing of Dientamoeba fragilis. Br. J. Biomed. Sci. 61, 153–155, 2004.
28. Vandenberg, O. et al. Clinical and microbiological features of dientamoebiasis in patients suspected of suffering from a parasitic gastrointestinal illness: A comparison of Dientamoeba fragilis and Giardia lamblia infections. Int. J. Infect. Dis. 10, 255–261, 2006.
29. Hussein, E.M., Al-Mohammed, H.I., and Hussein, A.M. Genetic diversity of Dientamoeba fragilis isolates of irritable bowel syndrome patients by high-resolution melting-curve (HRM) analysis. Parasitol. Res. 105, 1053–1060, 2009.
30. Crotti, D., and D’Annibale, M.L. Role of Dientamoeba fragilis in human bowel infections. Infez. Med. 15, 30–39, 2007.
31. Spencer, M.J., Garcia, L.S., and Chapin, M.R. Dientamoeba fragilis. An intestinal pathogen in children? Am. J. Dis. Child. 133, 390–393, 1979.
32. Stensvold, C.R., Arendrup, M.C., Molbak, K., and Nielsen, H.V. The prevalence of Dientamoeba fragilis in patients with suspected enteroparasitic disease in a metropolitan area in Denmark. Clin. Microbiol. Infect. 13, 839–842, 2007.
33. Schuster, H., and Jackson, R.S. Prevalence of Dientamoeba fragilis among patients consulting complementary medicine practitioners in the British Isles. J. Clin. Pathol. 62, 182–184, 2009.
34. Rayan, H.Z., Ismail, O.A., and El Gayar, E.K. Prevalence and clinical features of Dientamoeba fragilis infections in patients suspected to have intestinal parasitic infection. J. Egypt. Soc. Parasitol. 37, 599–608, 2007.
35. Crotti, D., and D’Annibale, M.L. Intestinal infections caused by Dientamoeba fragilis and Giardia duodenalis in our experience. Recenti Prog. Med. 98, 361–366, 2007.
36. Girginkardesler, N., Coskun, S., Cuneyt Balcioglu, I., Ertan, P., and Ok, U.Z. Dientamoeba fragilis, a neglected cause of diarrhea, successfully treated with secnidazole. Clin. Microbiol. Infect. 9, 110–113, 2003.
37. Crotti, D., and D’Annibale, M.L. Human intestinal parasitosis: Role of Dientamoeba fragilis in human infections. Ann. Ig. 19, 27–34, 2007. 38. Crotti, D. et al. Dientamoeba fragilis is more prevalent than Giardia duodenalis in children and adults attending a day care centre in Central Italy. Parasite 12, 165–170, 2005. 39. Butler, W.P. Dientamoeba fragilis. An unusual intestinal pathogen. Dig. Dis. Sci. 41, 1811–1813, 1996. 40. Shein, R., and Gelb, A. Colitis due to Dientamoeba fragilis. Am. J. Gastroenterol. 78, 634–636, 1983. 41. Preiss, U., Ockert, G., Bromme, S., and Otto, A. Dientamoeba fragilis infection, a cause of gastrointestinal symptoms in childhood. Klin. Padiatr. 202, 120–123, 1990. 42. Norberg, A., Nord, C.E., and Evengard, B. Dientamoeba fragilis—A protozoal infection which may cause severe bowel distress. Clin. Microbiol. Infect. 9, 65–68, 2003. 43. Dardick, K.R. Tetracycline treatment of Dientamoeba fragilis. Conn. Med. 47, 69–70, 1983. 44. Vandenberg, O., Souayah, H., Mouchet, F., Dediste, A., and van Gool, T. Treatment of Dientamoeba fragilis infection with paromomycin. Pediatr. Infect. Dis. J. 26, 88–90, 2007. 45. Kurt, O., Girginkardesler, N., Balcioglu, I.C., Ozbilgin, A., and Ok, U.Z. A comparison of metronidazole and singledose ornidazole for the treatment of dientamoebiasis. Clin. Microbiol. Infect. 14, 601–604, 2008. 46. Oxner, R.B., Paltridge, G.P., Chapman, B.A., Cook, H.B., and Sheppard, P.F. Dientamoeba fragilis: A bowel pathogen? N. Z. Med. J. 100, 64–65, 1987. 47. van Gool, T., Weijts, R., Lommerse, E., and Mank, T.G. Triple Faeces test: An effective tool for detection of intestinal parasites in routine clinical practice. Eur. J. Clin. Microbiol. Infect. Dis. 22, 284–290, 2003. 48. Hiatt, R.A., Markell, E.K., and Ng, E. How many stool examinations are necessary to detect pathogenic intestinal protozoa? Am. J. Trop. Med. Hyg. 53, 36–39, 1995. 49. Sawangjaroen, N., Luke, R., and Prociv, P. Diagnosis by faecal culture of Dientamoeba fragilis infections in Australian patients with diarrhoea. Trans. R. Soc. Trop. Med. Hyg. 87, 163–165, 1993. 50. Windsor, J.J., Macfarlane, L., Hughes-Thapa, G., Jones, S.K., and Whiteside, T.M. Detection of Dientamoeba fragilis by culture. Br. J. Biomed. Sci. 60, 79–83, 2003. 51. Clark, C.G., and Diamond, L.S. Methods for cultivation of luminal parasitic protists of clinical importance. Clin. Microbiol. Rev. 15, 329–341, 2002. 52. Stark, D., Beebe, N., Marriott, D., Ellis, J., and Harkness, J. Detection of Dientamoeba fragilis in fresh stool specimens using PCR. Int. J. Parasitol. 35, 57–62, 2005. 53. Stark, D., Beebe, N., Marriott, D., Ellis, J., and Harkness, J. Evaluation of three diagnostic methods, including real-time PCR, for detection of Dientamoeba fragilis in stool specimens. J. Clin. Microbiol. 44, 232–235, 2006. 54. Verweij, J.J. et al. Real-time PCR for the detection of Dientamoeba fragilis in fecal samples. Mol. Cell. Probes 21, 400–404, 2007.
6 Chapter 6 - Entamoeba
20. World Health Organization. World Health Organization/Pan American Health Organization/UNESCO report of a consultation of experts on amoebiasis. Wkly. Epidemiol. Rec. 72, 97, 1997.
21. Hutson, C.D., and Petri, W.A. Jr. Amoebiasis: Clinical implications of the recognition of Entamoeba dispar. Curr. Infect. Dis. Rep. 1, 441, 1999.
22. Walsh, J.A. Problems in recognition and diagnosis of amoebiasis: Estimation of the global magnitude of morbidity and mortality. Rev. Infect. Dis. 8, 228, 1986.
23. Petri, W.A. Jr. Recent advances in amoebiasis. Crit. Rev. Clin. Lab. Sci. 33, 1, 1996.
24. Leroy, A.G. et al. Contact-dependent transfer of the galactose-speci�c lectin of Entamoeba histolytica to the lateral surface of enterocytes in culture. Infect. Immun. 63, 4253, 1995.
25. Stanley, S.L. et al. Role of the Entamoeba histolytica cysteine proteinase in amoebic liver abscess formation in severe combined immunode�cient mice. Infect. Immun. 63, 1587, 1995.
26. Vazquez, J. et al. Characterization of adhesion plates induced by the interaction of Entamoeba histolytica trophozoites with �bronectin. Cell Motil. Cytoskeleton 32, 37, 1995.
27. Weiss, S.J. Tissue destruction by neutrophils. N. Engl. J. Med. 320, 365, 1988.
28. Mann, B.J. Structure and function of the Entamoeba histolytica Gal/GalNAc lectin. Int. Rev. Cytol 216, 59, 2002.
29. Chadee, K. et al. Rat and human colonic mucins bind to and inhibit adherence lectin of Entamoeba histolytica. J. Clin. Invest. 80, 1245, 1991.
30. Ravdin, J.I. et al. Characterization of cell surface carbohydrate receptors for Entamoeba histolytica adherence lectin. Infect. Immun. 57, 2179, 1989.
31. Leippe, M. Amoebapores. Parsitol. Today 13, 178, 1997. 32. Que, X., and Reed, S.L. Cysteine proteinases and the pathogenesis of amoebiasis. Clin. Microbiol. Rev. 13, 196, 2000.
33. Stenson, W.F. et al. Amoebic infection in the human colon induces cyclooxgenase-2. Infect. Immun. 69, 3382, 2001.
34. Stanley, S.L. Jr. Amoebiasis. Lancet 361, 1025, 2003.
35. Gathiram, V., and Jackson, T.F.H.G. A longitudinal study of asymptomatic carriers of pathogenic zymodemes of Entamoeba histolytica. S. Afr. Med. J. 72, 669, 1987.
36. Haque, R. et al. Amoebiasis and mucosal IgA antibody against the Entamoeba histolytica adherence lectin in Bangladeshi children. J. Infect. Dis. 183, 1787, 2001.
37. Lewis, E.A., and Antia, A.U. Amoebic colitis: Review of 295 cases. Trans. R. Soc. Trop. Med. Hyg. 63, 633, 1969.
38. Brandt, H., and Perez-Tamayo, R. Pathology of human amoebiais. Hum. Pathol. 1, 351, 1970.
39. Kapoor, O.P., Nathwani, B.N., and Joshi, V.R. Amoebic peritonitis: A study of 73 cases. J. Trop. Med. Hyg. 75, 11, 1972.
40. Takahashi, T. et al. Fulminant amoebic colitis: Analysis of 55 cases. Dis. Colon Rectum 40, 1362, 1997.
41. Adams, E.B., and MacLeod, I.N. Invasive amoebiasis. I. Amoebic dysentery and its complications. Medicine 56, 315, 1977.
42. Mhlanga, B.R. et al. Amoebiasis complicating carcinomas— A diagnostic dilemma. Am. J. Trop. Med. Hyg. 46, 759, 1992.
43. Lysy, J. et al. Crohn’s colitis complicated by superimposed invasive amoebic colitis. Am. J. Gastroenterol. 86, 1063, 1991.
44. Rustgi, A.K., and Richter, J.M. Pyogenic and amoebic liver abscess. Med. Clin. N. Am. 73, 487, 1989.
45. Orbison, J.A. et al. Amoebic brain abscess: Review of the literature and report of �ve additional cases. Medicine 30, 247, 1951. 46. Kirsc, D., and Diaz-Rivera, R.S. Perinephric abscess: A previously unrecognized complication of amoebiasis. Am. J. Med. Sci. 206, 373, 1943. 47. Heinz, K.P. Amoebic infection of the female genital tract: A report of three cases. S. Afr. Med. J. 47, 1795, 1973. 48. O’Leary, R.K., and Posen, J. Amoebiasis of the penis. S. Afr. Med. J. 65, 113, 1984. 49. Ramdial, P.K. et al. Amoebiasis cutis revisited. J. Cutan. Pathol. 34, 620, 2007. 50. Pehrson, P.O., and Bengtsson, E. A long-term follow up study of amoebiasis treated with metronidazole. Scand. J. Infect. Dis. 16, 195, 1984. 51. Lasserre, R. et al. Single-day drug treatment of amoebic liver abscess. Am. J. Trop. Med. Hyg. 32, 723, 1983. 52. Akgun, Y., Tacyildiz, I.H., and Celik, Y. Amoebic liver abscess: Changing trends over 20 years. World J. Surg. 23, 102, 1999. 53. Pillai, D.R., and Kain, K.C. Immunochromatographic stripbased detection of Entamoeba histolytica-E. dispar and Giardia lamblia coproantigen. J. Clin. Microbiol. 37, 3017, 1999. 54. Proctor, E.M. Laboratory diagnosis of amoebiasis. Clin. Lab. Med. 11, 829, 1991. 55. Clark, C.G., and Diamond, L.S. Methods for cultivation of luminal parasitic protists of clinical importance. Clin. Microbiol. Rev. 15, 329, 2002. 56. Balamuth, W. Improved egg yolk medium for the cultivation of Entamoeba histolytica and other intestinal protozoa. Am. J. Clin. Pathol. 16, 380, 1946. 57. Jones, W.R. The experimental infection of rats with Entamoeba histolytica; with a method for evaluating the anti-amoebic properties of new compounds. Ann. Trop. Med. Parasitol. 40, 130, 1946. 58. Diamond, L.S. A new liquid medium for xenic cultivation of Entamoeba histolytica and other lumen dwelling protozoa. J. Parasitol. 68, 958, 1982. 59. Robinson, G.L. The laboratory diagnosis of human parasitic amoebae. Trans. R. Soc. Trop. Med. Hyg. 62, 285, 1968. 60. Diamond, L.S. Axenic cultivation of Entamoeba histolytica. Science 134, 336, 1961. 61. Sargeaunt, P.G. et al. The reliability of Entamoeba histolytica zymodemes in clinical laboratory diagnosis. Arch. Invest. Med. 18, 69, 1987. 62. Zengzhu, G. et al. Analysis by enzyme-linked immunosorbent assay and PCR of human liver abscess aspirates from patients in China for Entamoeba histolytica. J. Clin. Microbiol. 37, 3034, 1999. 63. Haque, R. et al. Rapid diagnosis of Entamoeba infection by using Entamoeba and Entamoeba histolytica stool antigen detection kits. J. Clin. Microbiol. 33, 2558, 1995. 64. Garcia, L.S., Shimizu, R.Y., and Bernard C.N. Detection of Giardia lamblia, Entamoeba histolytical Entamoeba dispar, and Cryptosporidium parvum antigens in human fecal specimens using the Triage parasite panel enzyme immunoassay. J. Clin. Microbiol. 38, 3337, 2000. 65. Pillai, D.R. et al. Entamoeba histolytica and Entamoeba dispar: Epidemiology and comparison of diagnostic methods in a setting of nonendemicity. Clin. Infect. Dis. 29, 1315, 1999. 66. Levia, B. et al. Overdiagnosis of Entamoeba histolytica and Entamoeba dispar in Nicaragua: A microscopic, triage parasite panel and PCR study. Arch. Med. Res. 37, 529, 2006. 67. Haque, R. et al. Entamoeba histolytica and Entamoeba dispar infection in children in Bangladesh. J. Infect. Dis. 175, 734, 1997. 68. Solaymani-Mohammadi, S. et al. Comparison of a stool antigen detection kit and PCR for diagnosis of Entamoeba histolytica and Entamoeba dispar infections in asymptomatic cyst passers in Iran. J. Clin. Microbiol. 44, 2258, 2006.
69. Gatti, S. et al. Amoebic infections due to the Entamoeba histolytica-Entamoeba dispar complex: A study of the incidence in a remote rural area of Ecuador. Am. J. Trop. Med. Hyg. 67, 123, 2002.
70. Stark, D. et al. Comparison of stool antigen detection kits with PCR for the diagnosis of amoebiasis. J. Clin. Microbiol. 46, 1678, 2008.
71. Gonin, P., and Trudel, L. Detection and differentiation of Entamoeba histolytica and Entamoeba dispar isolates in clinical samples by PCR and enzyme-linked immunosorbent assay. J. Clin. Microbiol. 41, 237, 2003.
72. Visser, L.G. et al. Diagnostic methods for differentiation of Entamoeba histolytica and Entamoeba dispar in carriers: Performance and clinical implications in a non-endemic setting. Int. J. Med. Microbiol. 296, 397, 2006.
73. Jelinek, T. et al. Evaluation of an antigen-capture enzyme immunoassay for detection of Entamoeba histolytica in stool samples. Eur. J. Clin. Microbiol. Infect. Dis. 15, 752, 1996.
74. Ong, S.J. et al. Use of the ProSpecT microplate enzyme immunoassay for the detection of pathogenic and non-pathogenic Entamoeba histolytica in faecal specimens. Trans. R. Soc. Trop. Med. Hyg. 90, 248, 1996.
75. Mirelman, D., Nuchamowitz, Y., and Stolarsky, T. Comparison of use of enzyme-linked immunosorbent assay-based kits and PCR ampli�cation of rRNA genes for simultaneous detection of Entamoeba histolytica and E. dispar. J. Clin. Microbiol. 35, 2405, 1997.
76. Edman, U. et al. Characterisation of an immuno-dominant variable surface antigen from pathogenic and non-pathogenic Entamoeba histolytica. J. Exp. Med. 172, 879, 1990.
77. Tannich, E., and Burchard, G.D. Differentiation of pathogenic from non-pathogenic Entamoeba histolytica by restriction fragment analysis of a single gene ampli�ed in vitro. J. Clin. Microbiol. 29, 250, 1991.
78. Gar�nkel, L. et al. DNA probes speci�c for Entamoeba histolytica possessing pathogenic and nonpathogenic zymodemes. Infect. Immun. 57, 926, 1989.
79. Romero, J.L. et al. Use of polymerase chain reaction and nonradioactive DNA probes to diagnose Entamoeba histolytica infection. Arch. Med. Res. 23, 277, 1992.
80. Bhattacharya, S. et al. Circular DNA of Entamoeba histolytica encodes ribosomal RNA. J. Protozool. 36, 455, 1989.
81. Troll, H., Marti, H., and Weiss, N. Simple differential detection of Entamoeba histolytica and Entamoeba dispar in fresh stool specimens by sodium acetate-acetic acid-formalin concentration and PCR. J. Clin. Microbiol. 35, 1701, 1997.
82. Ramos, F. et al. Prevalence of Entamoeba histolytica and Entamoeba dispar in a highly endemic rural population. Arch. Med. Res. 31, 34, 2000.
83. Verweij, J.J. et al. Differentiation of Entamoeba histolytica and Entamoeba dispar cysts using polymerase chain reaction on DNA isolated from faeces with spin columns. Eur. J. Clin. Microbiol. Infect. Dis. 19, 358, 2000.
84. Heckendorn, F. et al. Species-speci�c �eld testing of Entamoeba spp. in an area of high endemicity. Trans. Roy. Soc. Trop. Med. Hyg. 96, 521, 2002.
85. Ramos, F. et al. Entamoeba histolytica and Entamoeba dispar: Prevalence infection in a rural Mexican community, Exp. Parasitol. 110, 327, 2005.
86. Hamzah, Z. et al. Differential detection of Entamoeba histolytica, Entamoeba dispar, and Entamoeba moshkovskii by a single-round PCR assay. J. Clin. Microbiol. 44, 3196, 2006. 87. Acuna-Soto, R. et al. Application of the polymerase chain reaction to the epidemiology of pathogenic and nonpathogenic Entamoeba histolytica. Am. J. Trop. Med. Hyg. 48, 58, 1993. 88. Britten, D. et al. An improved colorimetric PCR-based method for the detection and differentiation of Entamoeba histolytica and Entamoeba dispar. J. Clin. Microbiol. 35, 1108, 1997. 89. Tachibana, H.. et al. Distinguishing pathogenic isolates of Entamoeba histolytica by polymerase chain reaction. J. Infect. Dis. 164, 825, 1991. 90. Tachibana, H. et al. Asymptomatic cyst passers of Entamoeba histolytica but not Entamoeba dispar in institutions for the mentally retarded in Japan. Parasitol. Int. 49, 31, 2000. 91. Rivera, W.L. et al. Differentiation of Entamoeba histolytica and E. dispar DNA from cysts present in stool specimens by polymerase chain reaction: Its �eld application in the Philippines. Parasitol. Res. 82, 585, 1996. 92. Rivera, W.L., Santos S.R., and Kanbara, H. Prevalence and genetic diversity of Entamoeba histolytica in an institution for the mentally retarded in the Philippines. Parasitol. Res. 98, 106, 2006. 93. Ayeh-Kumi, P.F. et al. Entamoeba histolytica: Genetic diversity of clinical isolates from Bangladesh as demonstrated by polymorphisms in the serine-rich gene. Exp. Parasitol. 99, 80, 2001. 94. Freitas, M.A. et al. A single step duplex PCR to distinguish Entamoeba histolytica from Entamoeba dispar. Parasitology 128, 625, 2004. 95. Zindrou, S. Speci�c detection of Entamoeba histolytica DNA by hemolysin gene targeted PCR. Acta Trop. 78, 117, 2001. 96. Nunez, Y.O. et al. Multiplex polymerase chain reaction ampli�cation and differentiation of Entamoeba histolytica and Entamoeba dispar DNA from stool samples. Am. J. Trop. Med. Hyg. 64, 293, 2001. 97. Khairnar, K., and Parija, S.C. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 7, 47, 2007. 98. Zaman, S. et al. Direct ampli�cation of Entamoeba histolytica DNA from amoebic liver abscess pus using polymerase chain reaction. Parasitol. Res. 86, 724, 2000. 99. Tachibana, H. et al. Detection of pathogenic Entamoeba histolytica DNA in liver abscess �uid by polymerase chain reaction. Int. J. Parasitol. 22, 1193, 1992. 100. Khan, U. et al. Detection of Entamoeba histolytica using polymerase chain reaction in pus samples from amoebic liver abscess. Indian J. Gastroenterol. 25, 55, 2006. 101. Qvarnstrom, Y. et al. Comparison of real-time PCR protocols for differential laboratory diagnosis of amoebiasis. J. Clin. Microbiol. 43, 5491, 2005. 102. Blessman, J. et al. Real-time PCR for detection and differentiation of Entamoeba histolytica and Entamoeba dispar in fecal samples. J. Clin. Microbiol. 40, 4413, 2002. 103. Kebede, A. et al. The use of real-time PCR to identify Entamoeba histolytica and E. dispar infections in prisoners and primary-school children in Ethiopia. Ann. Trop. Med. Parasitol. 98, 43, 2004. 104. Verweij, J.J. et al. Simultaneous detection of Entamoeba histolytica, Giardia lamblia, and Cryptosporidium parvum in faecal samples by using multiplex real-time PCR. J. Clin. Microbiol. 42, 1220, 2004. 105. Verweij, J.J. et al. Short communication: Prevalence of Entamoeba histolytica and Entamoeba dispar in northern Ghana. Trop. Med. Int. Health. 8, 1153, 2003.
106. Roy, S. et al. Real-time PCR assay for the diagnosis of Entamoeba histolytica infection. J. Clin. Microbiol. 43, 2168, 2005.
107. Furrows, S.J., Moody, A.H., and Chiodini, P.L. Comparison of PCR and antigen detection methods for diagnosis of Entamoeba histolytica infection. J. Clin. Pathol. 57, 1264, 2004.
108. Lebbad, M., and Svard, S.G. PCR differentiation of Entamoeba histolytica and Entamoeba dispar from patients with amoeba infection initially diagnosed by microscopy. Scand. J. Infect. Dis. 37, 680, 2005.
109. Katzwinkel-Wladarsch, S., Loscher, T., and Rinder, H. Direct ampli�cation and differentiation of pathogenic and nonpathogenic Entamoeba histolytica DNA from stool specimens. Am. J. Trop. Med. Hyg. 51, 115, 1994.
110. Ramos, F. et al. The effect of formalin �xation on the polymerase chain reaction characterisation of Entamoeba histolytica. Trans. R. Soc. Trop. Med. Hyg. 93, 335, 1999.
111. Evangelopoulos, A. et al. A nested, multiplex, PCR assay for the simultaneous detection and differentiation of Entamoeba histolytica and Entamoeba dispar in faeces. Ann. Trop. Med. Parasitol. 94, 233, 2000.
112. Myjak, P., Kur, J., and Pietkiewicz, H. Usefulness of new DNA extraction procedure for PCR technique in species identi�cation of Entamoeba isolates. Wiad. Parazytol. 43, 163, 1997. 113. Verweij, J.J., Polderman, A.M., and Clark, C.G. Genetic variation among human isolates of uninucleated cyst-producing Entamoeba species. J. Clin. Microbiol. 39, 1644, 2001. 114. van Hal, S.J. et al. Amoebiasis: Current status in Australia. Med. J. Aust. 186, 412, 2007. 115. Paul, J., Srivastava, S., and Bhattacharya, S. Molecular methods for diagnosis of Entamoeba histolytica in a clinical setting: An overview. Exp. Parasitol. 116, 35, 2007. 116. Wang, Z., Vora, G.J., and Stenger, D.A. Detection and genotyping of Entamoeba histolytica, Entamoeba dispar, Giardia lamblia, and Cryptosporidium parvum by oligonucleotide microarray. J. Clin. Microbiol. 42, 3262, 2004. 117. Palacios, G. et al. Panmicrobial oligonucleotide array for diagnosis of infectious diseases. Emerg. Infect. Dis. 13, 73, 2007. 118. Cruz, H.J., Rosa, C.C., and Oliva, A.G. Immunosensors for diagnostic applications. Parasitol. Res. 88, S4, 2002. 119. Ricci, F. et al. A review on novel developments and applications of immunosensors in food analysis. Anal. Chim. Acta 605, 111, 2007. 120. Johnson-White, B., Lin, B., and Ligler, F.S. Combination of immunosensor detection with viability testing and con�rmation using the polymerase chain reaction and culture. Anal. Chem. 79, 140, 2007. 121. Haque, R. et al. Comparison of PCR, isoenzyme analysis, and antigen detection for diagnosis of Entamoeba histolytica infection. J. Clin. Microbiol. 36, 449, 1995. 7 Chapter 7 - Giardia
1. Yason, J.A., and Rivera, W.L. Genotyping of Giardia duodenalis isolates among residents of slum area in Manila, Philippines. Parasitol. Res. 101, 681, 2007.
2. Thompson, R.C. Giardiasis as a re-emerging infectious disease and its zoonotic potential. Int. J. Parasitol. 30, 1259, 2000.
3. Hellard, M.E. et al. Prevalence of enteric pathogens among community based asymptomatic individuals. J. Gastroenterol. Hepatol. 15, 290, 2000.
4. Thompson, R.C. The zoonotic signi�cance and molecular epidemiology of Giardia and giardiasis. Vet. Parasitol. 126, 15, 2004.
5. Thompson, R.C., and Monis, P.T. Variation in Giardia: Implications for taxonomy and epidemiology. Adv. Parasitol. 58, 69, 2004.
6. Appelbee, A.J., Thompson, R.C., and Olson, M.E. Giardia and Cryptosporidium in mammalian wildlife–current status and future needs. Trends Parasitol. 21, 370, 2005.
7. Caccio, S.M. et al. Unravelling Cryptosporidium and Giardia epidemiology. Trends Parasitol. 21, 430, 2005.
8. Monis, P.T. et al. Molecular systematics of the parasitic protozoan Giardia intestinalis. Mol. Biol. Evol. 16, 1135, 1999.
9. Monis, P.T. et al. Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infect. Genet. Evol. 3, 29, 2003.
10. Read, C.M., Monis, P.T., and Thompson, R.C. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect. Genet. Evol. 4, 125, 2004. 11. Sulaiman, I.M. et al. Triosephosphate isomerase gene characterization and potential zoonotic transmission of Giardia duodenalis. Emerg. Infect. Dis. 9, 1444, 2003. 12. Wielinga, C.M., and Thompson, R.C. Comparative evaluation of Giardia duodenalis sequence data. Parasitology 134, 1795, 2007. 13. Hopkins, R.M. et al. Ribosomal RNA sequencing reveals differences between the genotypes of Giardia isolates recovered from humans and dogs living in the same locality. J. Parasitol. 83, 44, 1997. 14. Monis, P.T., and Andrews, R.H. Molecular epidemiology: Assumptions and limitations of commonly applied methods. Int. J. Parasitol. 28, 981, 1998. 15. Leonhard, S. et al. The molecular characterisation of Giardia from dogs in southern Germany. Vet. Parasitol. 150, 33, 2007. 16. Ey, P.L. et al. Genetic analysis of Giardia from hoofed farm animals reveals artiodactyl-speci�c and potentially zoonotic genotypes. J. Eukaryot. Microbiol. 44, 626, 1997. 17. Huang, D.B., and White, A.C. An updated review on Cryptosporidium and Giardia. Gastroenterol. Clin. North Am. 35, 291, 2006. 18. Karanis, P., Kourenti, C., and Smith, H. Waterborne transmission of protozoan parasites: A worldwide review of outbreaks and lessons learnt. J. Water Health 5, 1, 2007. 19. Savioli, L., Smith, H., and Thompson, A. Giardia and Cryptosporidium join the “Neglected Diseases Initiative.” Trends Parasitol. 22, 203, 2006. 20. Ferrer, S.R. et al. A hierarchical model for studying risk factors for childhood diarrhoea: A case–control study in a middle-income country. Int. J. Epidemiol. 37, 805, 2008. 21. Davies, A.P. et al. Asymptomatic carriage of protozoan parasites in children in day care centers in the United Kingdom. Pediatr. Infect. Dis. J., 28, 838, 2009. 22. Giangaspero, A., Berrilli, F., and Brandonisio, O. Giardia and Cryptosporidium and public health: The epidemiological scenario from the Italian perspective. Parasitol. Res. 101, 1169, 2007. 23. Horman, A. et al. Meta-analysis in assessment of the prevalence and annual incidence of Giardia spp. and Cryptosporidium spp. infections in humans in the Nordic countries. Int. J. Parasitol. 34, 1337, 2004. 24. Almeida, A.A. et al. Genotype analysis of Giardia isolated from asymptomatic children in northern Portugal. J. Eukaryot. Microbiol. 53, S177, 2006. 25. Berrilli, F. et al. Prevalence and genotyping of human isolates of Giardia duodenalis from Albania. Parasitol. Int. 55, 295, 2006. 26. Morales-Espinoza, E.M. et al. Intestinal parasites in children, in highly deprived areas in the border region of Chiapas, Mexico. Salud Publica Mex. 45, 379, 2003. 27. Diaz, E. et al. Epidemiology and control of intestinal parasites with nitazoxanide in children in Mexico. Am. J. Trop. Med. Hyg. 68, 384, 2003. 28. Paniagua, G.L. et al. Two or more enteropathogens are associated with diarrhoea in Mexican children. Ann. Clin. Microbiol. Antimicrob. 6, 17, 2007. 29. Sanchez-Vega, J.T. et al. Cryptosporidiosis and other intestinal protozoan infections in children less than one year of age in Mexico City. Am. J. Trop. Med. Hyg. 75, 1095, 2006. 30. Silva, R.R. et al. Association between nutritional status, environmental and socio-economic factors and Giardia lamblia infections among children aged 6–71 months in Brazil. Trans. R. Soc. Trop. Med. Hyg. 103, 512, 2009. 31. Maia, M.M. et al. Intestinal parasitic infection and associated risk factors, among children presenting at outpatient clinics in Manaus, Amazonas state, Brazil. Ann. Trop. Med. Parasitol. 103, 583, 2009.
32. Korkes, F. et al. Relationship between intestinal parasitic infection in children and soil contamination in an Urban slum. J. Trop. Pediatr., 2008.
33. de Carvalho, T.B., de Carvalho, L.R., and Mascarini, L.M. Occurrence of enteroparasites in day care centers in Botucatu (Sao Paulo State, Brazil) with emphasis on Cryptosporidium sp., Giardia duodenalis and Enterobius vermicularis. Rev. Inst. Med. Trop. Sao Paulo, 48, 269, 2006.
34. Teixeira, J.C., Heller, L., and Barreto, M.L. Giardia duodenalis infection: Risk factors for children living in sub-standard settlements in Brazil. Cad Saude Publica, 23, 1489, 2007.
35. Guignard, S. et al. Prevalence of enteroparasites in a residence for children in the Cordoba Province, Argentina. Eur. J. Epidemiol. 16, 287, 2000.
36. Perez Cordon, G. et al. Prevalence of enteroparasites and genotyping of Giardia lamblia in Peruvian children. Parasitol. Res. 103, 459, 2008.
37. Cabrera, M., Verastegui, M., and Cabrera, R. Prevalence of entero-parasitosis in one Andean community in the Province of Victor Fajardo, Ayacucho, Peru. Rev. Gastroenterol. Peru, 25, 150, 2005.
38. Chaves Mdel, P. et al. Giardia duodenalis prevalence and associated risk factors in preschool and school-age children of rural Colombia. Biomedica, 27, 345, 2007.
39. Dib, H.H., Lu, S.Q., and Wen, S.F. Prevalence of Giardia lamblia with or without diarrhea in South East, South East Asia and the Far East. Parasitol. Res. 103, 239, 2008.
40. Nematian, J., Gholamrezanezhad, A., and Nematian, E. Giardiasis and other intestinal parasitic infections in relation to anthropometric indicators of malnutrition: A large, population-based survey of schoolchildren in Tehran. Ann. Trop. Med. Parasitol. 102, 209, 2008.
41. Hamze, M. et al. Prevalence of infection by intestinal parasites in north Lebanon: 1997–2001. East. Mediterr. Health J. 10, 343, 2004.
42. Foronda, P. et al. Identi�cation of genotypes of Giardia intestinalis of human isolates in Egypt. Parasitol. Res., 103, 1177, 2008.
43. Lalle, M. et al. High genetic polymorphism among Giardia duodenalis isolates from Sahrawi children. Trans. R. Soc. Trop. Med. Hyg., 103, 834, 2009.
44. El Kettani, S., Azzouzi, E.M., and Maata, A. Prevalence of Giardia intestinalis in a farming population using sewage water in agriculture, Settat, Morocco. Med. Mal. Infect. 36, 322, 2006.
45. Perch, M. et al. Seven years’ experience with Cryptosporidium parvum in Guinea-Bissau, West Africa. Ann. Trop. Paediatr. 21, 313, 2001.
46. Gbakima, A.A. et al. Intestinal protozoa and intestinal helminthic infections in displacement camps in Sierra Leone. Afr. J. Med. Sci. 36, 1, 2007.
47. Adams, V.J. et al. Paradoxical helminthiasis and giardiasis in Cape Town, South Africa: Epidemiology and control. Afr. Health Sci. 5, 276, 2005.
48. Pollock, K.G. et al. Giardia surveillance in Scotland, 1988– 2003. Eur. J. Clin. Microbiol. Infect. Dis. 24, 571, 2005.
49. Hoque, E. et al. A descriptive epidemiology of giardiasis in New Zealand and gaps in surveillance data. N Z Med. J. 117, U1149, 2004.
50. Yoder, J.S., and Beach, M.J. Giardiasis surveillance–United States, 2003–2005. MMWR Surveill. Summ. 56, 11, 2007.
51. Hlavsa, M.C., Watson, J.C., and Beach, M.J. Giardiasis surveillance–United States, 1998–2002. MMWR Surveill. Summ. 54, 9, 2005. 52. Furness, B.W., Beach, M.J., and Roberts, J.M. Giardiasis surveillance–United States, 1992–1997. MMWR CDC Surveill. Summ. 49, 1, 2000. 53. Saksirisampant, W. et al. Intestinal parasitic infestations among children in an orphanage in Pathum Thani province. J. Med. Assoc. Thai. 86, S263, 2003. 54. Mohammed Mahdy, A.K. et al. Giardia intestinalis genotypes: Risk factors and correlation with clinical symptoms. Acta Trop. 112, 67, 2009. 55. Sharma, B.K. et al. Prevalence of intestinal parasitic infestation in schoolchildren in the northeastern part of Kathmandu Valley, Nepal. Southeast Asian J. Trop. Med. Public Health 35, 501, 2004. 56. Odoi, A. et al. Geographical and temporal distribution of human giardiasis in Ontario, Canada. Int. J. Health Geogr. 2, 5, 2003. 57. Wongstitwilairoong, B. et al. Intestinal parasitic infections among pre-school children in Sangkhlaburi, Thailand. Am. J. Trop. Med. Hyg. 76, 345, 2007. 58. Vargas, M. et al. Etiology of diarrhea in children less than �ve years of age in Ifakara, Tanzania. Am. J. Trop. Med. Hyg. 70, 536, 2004. 59. Xiao, L., and Fayer, R. Molecular characterization of species and genotypes of Cryptosporidium and Giardia and assessment of zoonotic transmission. Int. J. Parasitol. 38, 1239, 2008. 60. Haque, R. et al. Giardia assemblage A infection and diarrhea in Bangladesh. J. Infect. Dis. 192, 2171, 2005. 61. Eligio-Garcia, L., Cortes-Campos, A., and Jimenez-Cardoso, E. Genotype of Giardia intestinalis isolates from children and dogs and its relationship to host origin. Parasitol. Res. 97, 1, 2005. 62. Chin, A.C. et al. Strain-dependent induction of enterocyte apoptosis by Giardia lamblia disrupts epithelial barrier function in a caspase-3-dependent manner. Infect. Immun. 70, 3673, 2002. 63. Scott, K.G. et al. Intestinal infection with Giardia spp. reduces epithelial barrier function in a myosin light chain kinasedependent fashion. Gastroenterology 123, 1179, 2002. 64. Buret, A.G. et al. Giardia lamblia disrupts tight junctional ZO-1 and increases permeability in non-transformed human small intestinal epithelial monolayers: Effects of epidermal growth factor. Parasitology, 125, 11, 2002. 65. Nash, T.E. Surface antigenic variation in Giardia lamblia. Mol. Microbiol., 45, 585, 2002. 66. Touz, M.C., Conrad, J.T., and Nash, T.E. A novel palmitoyl acyl transferase controls surface protein palmitoylation and cytotoxicity in Giardia lamblia. Mol. Microbiol. 58, 999, 2005. 67. Hiatt, R.A., Markell, E.K., and Ng, E. How many stool examinations are necessary to detect pathogenic intestinal protozoa? Am. J. Trop. Med. Hyg. 53, 36, 1995. 68. Mank, T.G. et al. Sensitivity of microscopy versus enzyme immunoassay in the laboratory diagnosis of giardiasis. Eur. J. Clin. Microbiol. Infect. Dis. 16, 615, 1997. 69. Aldeen, W.E. et al. Comparison of nine commercially available enzyme-linked immunosorbent assays for detection of Giardia lamblia in fecal specimens. J. Clin. Microbiol. 36, 1338, 1998. 70. Garcia, L.S., and Shimizu, R.Y. Evaluation of nine immunoassay kits (enzyme immunoassay and direct �uorescence) for detection of Giardia lamblia and Cryptosporidium parvum in human fecal specimens. J. Clin. Microbiol. 35, 1526, 1997. 71. Zimmerman, S.K., and Needham, C.A. Comparison of conventional stool concentration and preserved-smear methods with Meri�uor Cryptosporidium/Giardia direct immuno�uorescence assay and ProSpecT Giardia EZ microplate assay for detection of Giardia lamblia. J. Clin. Microbiol. 33, 1942, 1995.
72. Chan, R. et al. Evaluation of a combination rapid immunoassay for detection of Giardia and Cryptosporidium antigens. J. Clin. Microbiol. 38, 393, 2000.
73. Aziz, H. et al. A comparison study of different methods used in the detection of Giardia lamblia. Clin. Lab. Sci. 14, 150, 2001.
74. Weiss, J.B., van Keulen, H., and Nash, T.E. Classi�cation of subgroups of Giardia lamblia based upon ribosomal RNA gene sequence using the polymerase chain reaction. Mol. Biochem. Parasitol. 54, 73, 1992.
75. Ey, P.L. et al. Giardia intestinalis: Detection of major genotypes by restriction analysis of gene ampli�cation products. Int. J. Parasitol. 23, 591, 1993.
76. van Keulen, H. et al. A three nucleotide signature sequence in small subunit rRNA divides human Giardia in two different genotypes. J. Eukaryot. Microbiol. 42, 392, 1995.
77. Baruch, A.C., Isaac-Renton, J., and Adam, R.D. The molecular epidemiology of Giardia lamblia: A sequence-based approach, J. Infect. Dis. 174, 233, 1996.
78. Monis, P.T. et al. Molecular genetic analysis of Giardia intestinalis isolates at the glutamate dehydrogenase locus. Parasitology, 112, 1, 1996.
79. Caccio, S.M., De Giacomo, M., and Pozio, E. Sequence analysis of the beta-giardin gene and development of a polymerase chain reaction-restriction fragment length polymorphism assay to genotype Giardia duodenalis cysts from human faecal samples. Int. J. Parasitol. 32, 1023, 2002.
80. Bertrand, I., Albertini, L., and Schwartzbrod, J. Comparison of two target genes for detection and genotyping of Giardia lamblia in human feces by PCR and PCR-restriction fragment length polymorphism. J. Clin. Microbiol. 43, 5940, 2005.
81. Aloisio, F. et al. Severe weight loss in lambs infected with Giardia duodenalis assemblage B. Vet. Parasitol. 142, 154, 2006.
82. Sulaiman, I.M. et al. Distribution of Giardia duodenalis genotypes and subgenotypes in raw urban wastewater in Milwaukee, Wisconsin. Appl. Environ. Microbiol. 70, 3776, 2004.
83. Feng, Y. et al. High intragenotypic diversity of Giardia duodenalis in dairy cattle on three farms. Parasitol. Res. 103, 87, 2008.
84. Traub, R.J. et al. Epidemiological and molecular evidence supports the zoonotic transmission of Giardia among humans and dogs living in the same community. Parasitology, 128, 253, 2004.
85. Amar, C.F. et al. Sensitive PCR-restriction fragment length polymorphism assay for detection and genotyping of Giardia duodenalis in human feces. J. Clin. Microbiol. 40, 446, 2002.
86. Robertson, L.J. et al. Application of genotyping during an extensive outbreak of waterborne giardiasis in Bergen, Norway, during autumn and winter 2004. Appl. Environ. Microbiol. 72, 2212, 2006.
87. Monis, P.T. The importance of systematics in parasitological research. Int. J. Parasitol. 29, 381, 1999.
88. Abe, N., Kimata, I., and Iseki, M. Identi�cation of genotypes of Giardia intestinalis isolates from dogs in Japan by direct sequencing of the PCR ampli�ed glutamate dehydrogenase gene. J. Vet. Med. Sci. 65, 29, 2003.
89. Itagaki, T. et al. Genotyping of Giardia intestinalis from domestic and wild animals in Japan using glutamete dehydrogenase gene sequencing. Vet. Parasitol. 133, 283, 2005. 90. Abe, N., Read, C., Thompson, R.C., and Iseki, M. Zoonotic genotype of Giardia intestinalis detected in a ferret. J. Parasitol. 91, 179, 2005. 91. Homan, W.L. et al. Characterization of Giardia duodenalis by polymerase-chain-reaction �ngerprinting. Parasitol. Res. 84, 707, 1998. 92. Matsubayashi, M., Kimata, I., and Abe, N. Identi�cation of genotypes of Giardia intestinalis isolates from a human and calf in Japan. J. Vet. Med. Sci. 67, 337, 2005. 93. van der Giessen, J.W. et al. Genotyping of Giardia in Dutch patients and animals: A phylogenetic analysis of human and animal isolates. Int. J. Parasitol. 36, 849, 2006. 94. Souza, S.L. et al. Molecular identi�cation of Giardia duodenalis isolates from humans, dogs, cats and cattle from the state of Sao Paulo, Brazil, by sequence analysis of fragments of glutamate dehydrogenase (gdh) coding gene. Vet. Parasitol. 149, 258, 2007. 95. van Keulen, H. et al. Presence of human Giardia in domestic, farm and wild animals, and environmental samples suggests a zoonotic potential for giardiasis. Vet. Parasitol. 108, 97, 2002. 96. Santin, M. et al. Cryptosporidium, Giardia and Enterocytozoon bieneusi in cats from Bogota (Colombia) and genotyping of isolates. Vet. Parasitol. 141, 334, 2006. 97. O’Handley, R.M. et al. Prevalence and genotypic characterisation of Giardia in dairy calves from Western Australia and Western Canada. Vet. Parasitol. 90, 193, 2000. 98. Trout, J.M., Santin, M., and Fayer, R. Prevalence of Giardia duodenalis genotypes in adult dairy cows. Vet. Parasitol. 147, 205, 2007. 99. Appelbee, A.J. et al. Prevalence and genotyping of Giardia duodenalis from beef calves in Alberta, Canada. Vet. Parasitol. 112, 289, 2003. 100. Mahbubani, M.H. et al. Detection of Giardia in environmental waters by immuno-PCR ampli�cation methods. Curr. Microbiol. 36, 107, 1998. 101. Mahbubani, M.H. et al. Differentiation of Giardia duodenalis from other Giardia spp. by using polymerase chain reaction and gene probes. J. Clin. Microbiol. 30, 74, 1992. 102. Volotao, A.C. et al. Genotyping of Giardia duodenalis from human and animal samples from Brazil using beta-giardin gene: A phylogenetic analysis. Acta Trop. 102, 10, 2007. 103. Lalle, M. et al. Genetic heterogeneity at the beta-giardin locus among human and animal isolates of Giardia duodenalis and identi�cation of potentially zoonotic subgenotypes. Int. J. Parasitol. 35, 207, 2005. 104. Ey, P.L. et al. Comparison of genetic groups determined by molecular and immunological analyses of Giardia isolated from animals and humans in Switzerland and Australia. Parasitol. Res. 82, 52, 1996. 105. Hunt, C.L., Ionas, G., and Brown, T.J. Prevalence and strain differentiation of Giardia intestinalis in calves in the Manawatu and Waikato regions of North Island, New Zealand. Vet. Parasitol. 91, 7, 2000. 106. Sogin, M.L. et al. Phylogenetic meaning of the kingdom concept: An unusual ribosomal RNA from Giardia lamblia. Science 243, 75, 1989. 107. van Keulen, H. et al. Nucleotide sequence of the 5.8S and large subunit rRNA genes and the internal transcribed spacer and part of the external spacer from Giardia ardeae. Nucleic Acids Res. 19, 6050, 1991. 108. Lu, S.Q., Baruch, A.C., and Adam, R.D. Molecular comparison of Giardia lamblia isolates. Int. J. Parasitol. 28, 1341, 1998. 109. Berrilli, F. et al. Genotype characterisation of Giardia duodenalis isolates from domestic and farm animals by SSU-rRNA gene sequencing. Vet. Parasitol. 122, 193, 2004.
110. Fayer, R. et al. Detection of Cryptosporidium felis and Giardia duodenalis assemblage F in a cat colony. Vet. Parasitol. 140, 44, 2006.
111. Andrews, R.H. et al. Giardia intestinalis: Electrophoretic evidence for a species complex. Int. J. Parasitol. 19, 183, 1989.
112. Santin, M. et al. First report of Giardia in coyotes (Canis latrans). J. Eukaryot. Microbiol. 50, S709, 2003.
113. Caccio, S.M., and Ryan, U. Molecular epidemiology of giardiasis. Mol. Biochem. Parasitol. 160, 75, 2008.
114. Caccio, S.M. et al. Multilocus genotyping of Giardia duodenalis reveals striking differences between assemblages A and B. Int. J. Parasitol., 2008.
115. Gelanew, T. et al. Molecular characterization of human isolates of Giardia duodenalis from Ethiopia. Acta Trop. 102, 92, 2007.
116. Geurden, T. et al. Multilocus genotyping of Cryptosporidium and Giardia in non-outbreak related cases of diarrhoea in human patients in Belgium. Parasitology, 136, 1161, 2009. 117. Caccio, S.M., and Sprong, H. Giardia duodenalis: Genetic recombination and its implications for taxonomy and molecular epidemiology. Exp. Parasitol., 124, 107, 2009. 118. Lebbad, M. et al. From mouse to moose: Multilocus genotyping of Giardia isolates from various animal species. Vet. Parasitol., 168, 231, 2009. 119. Xiao, L. et al. Host adaptation and host-parasite co-evolution in Cryptosporidium: Implications for taxonomy and public health. Int. J. Parasitol. 32, 1773, 2002. 120. Jiang, J. et al. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl. Environ. Microbiol. 71, 1135, 2005. 121. Smith, H.V. et al. Tools for investigating the environmental transmission of Cryptosporidium and Giardia infections in humans. Trends Parasitol. 22, 160, 2006. 122. Hunter, P.R., and Thompson, R.C. The zoonotic transmission of Giardia and Cryptosporidium. Int. J. Parasitol. 35, 1181, 2005. 8 Chapter 8 - Leishmania
1. WHO. http://www.who.int/Leishmaniasis/Leishmaniasis_ maps/en/index.html. Accessed September 2009.
2. Sánchez-Saldaña, L. et al. Leishmaniasis. Dermatol. Peruana. 14, 82–98, 2004.
3. Oumeish, O.Y. Cutaneous Leishmaniasis: A historical perspective. Clin. Dermatol. 17, 249, 54, 1999.
4. Catania, S. Leishmania viscerale in età pediatrica. Simposio. Controllo della Leishmaniosi: Veterinari e Medici a confronto. Domenica, Roma, pp. 8–9, 2003.
5. Davies, C.R. Effetto di una banda protettiva a base di deltametrina sull’ incidenza di leishmanisi viscerale zoonotica nei bambini. Simposio. Controllo della Leishmaniosi: Veterinari e Medici a confronto. Domenica, Roma, pp. 13–14, 2003.
6. Farrel, J.P. New World Leishmaniasis: The ecology of leishmaniasis and the diversity of leishmanial species in Central and South America, In: Black, S.J., and Seed J.R. (series eds), World Class Parasite: Vol. 4. Leishmania. Kluwer Academic Publishers, Springer-Verlag, New York, 2002.
7. Campino, L.M. Canine reservoirs and Leishmaniasis: Epidemiology and disease. In: Black, S.J., and Seed J.R. (series eds), World Class Parasite: Vol. 4. Leishmania. Kluwer Academic Publishers, Springer-Verlag, New York, 2002.
8. Trager, W. The development of Leishmania donovani in vitro at 37°C, effects of the kind of serum. J. Exp. Med. 97, 177, 1953.
9. Lofgren, R. The structure of Leishmania tropica as revealed by phase and electron microscopy. J. Bacteriol. 60, 617, 1950.
10. Crowther, S., Fulton, J.D., and Joyner, L.P. The metabolism of Leishmania donovani in culture. Biochem. J. 56, 182, 1954.
11. Chang, K.P., and Dwyer, D.M. Leishmania donovani. Hamster macrophage interactions in vitro: Cell entry, intracellular survival, and multiplication of amastigotos. J. Exp. Med. 147, 515, 1978.
12. Magill, A.J. Leishmaniasis, In: Strickland, G.T. (ed.), Hunter’s Tropical Medicine and Emerging and Infectious Diseases. Saunders WB Company, Philadelphia, PA, 2000.
13. Murray, H.W. et al. Advances in leishmaniasis. Lancet 366, 1561, 2005.
14. Kamhawi, S. Phlebotomine sand �ies and Leishmania parasites: Friends or foes. Trends Parasitol. 22, 439, 2006.
15. Cunningham, A.C. Parasitic adaptive mechanisms in infection by Leishmania. Exp. Mol. Pathol. 72, 132, 2002.
16. Chang, K.P., Chaudhuri, G., and Fong, D. Molecular determinants of Leishmania virulence. Annu. Rev. Microbiol. 44, 499, 1990.
17. Cruz, I. et al. Leishmania/HIV co-infections in the second decade. Indian J. Med. Res. 123, 357, 2006.
18. Laison, R., and Saw, J.J. Evolution, classi�cation and geographical distribution. In: Peters, W., and KillickKendrick, R. (eds.), The Lesihmaniases in Biology and Medicine. Academic Press, New York, 1987.
19. Mehlhorn, H. (ed.), Morphology. In Parasitology in Focus: Facts and Trends, 1st edn. Springer Verlag, Berlin, Heidelberg, New York, London, Paris, Tokyo, 1988. 20. Cáceres, O.R., and Montoya, Y.P. Diseño y evaluación de tres oligonucleótidos para la detección de Leishmania por PCR. Rev. Peru Med. Exp. Salud Publica 19, 109, 2002. 21. Leishmaniasis. Módulos Técnico. Serie Documentos Monográ�cos nº 8. O�cina General de Epidemiología (OGE) e Instituto Nacional de Salud (INS), Lima. 2000. 22. Herwaldt, B.L. Leishmaniasis. Lancet 354, 1191, 1999. 23. EI-Hassan, A.M., and Zijlstra, E.E. Leishmaniasis in Sudan. Mucosal leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 95, 519, 2001. 24. Singh, S., Mohapatra, D.P., and Sivakumar, R. Successful replacement of foetal calf serum with human urine for in vitro culture of Leishmania donovani. J. Commun. Dis. 32, 289, 2000. 25. Rosbotham, J.L. et al. Imported mucocutaneous leishmaniasis. Clin. Exp. Dermatol. 21, 288, 1996. 26. Sing, R., and Silvakumar, R. Recent advances in the diagnosis of leishmaniasis. J. Postgrad. Med. 49, 55, 2003. 27. Reithinger, R., and Dujardin, J.C. Molecular diagnosis of leishmaniasis: Current status and future applications. J. Clin. Microbiol. 45, 21, 2007. 28. Weigle, K.A. et al. Leishmanin skin test standardization and evaluation of safety, dose, storage, longevity of reaction and sensitization. Am. J. Trop. Med. Hyg. 44, 260, 1991. 29. Singh, S. New developments in diagnosis of leishmaniasis. Indian J. Med. Res. 123, 311, 2006. 30. Bensoussan, E. et al. Comparison of PCR assays for diagnosis of cutaneous leishmaniasis, J. Clin. Microbiol. 44, 1435, 2006. 31. Sassi, A. et al. Leishmanin skin test lymphoproliferative responses and cytokine production after symptomatic or asymptomatic Leishmania major infection in Tunisia. Clin. Exp. Immunol. 116, 127, 1999. 32. Schallig, H.D. et al. Development and application of “simple” diagnostic tools for visceral leishmaniasis. Med. Microbiol. Immunol. 190, 69, 2001. 33. Schoone, G.J. et al. A fast agglutination-screening test (FAST) for the detection of anti-Leishmania antibodies. Trans. R. Soc. Trop. Med. Hyg. 95, 400, 2001. 34. Montoya, I. et al. Diagnostico molecular para la leishmaniasis. Rev. Perú. Med. Exp. Salud Publica 14, 25, 1997. 35. Jensen, A.T. et al. Serodiagnosis of Leishmania donovani infections: Assessment of enzyme-linked immunosorbent assays using recombinant L. donovani gene B protein (GBP) and a peptide sequence of L. donovani GBP. Trans. R. Soc. Trop. Med. Hyg. 93, 157, 1999. 36. Raj, V.S. et al. Serodiagnosis of leishmaniasis with recombinant ORFF antigen. Am. J. Trop. Med. Hyg. 61, 482, 1999. 37. Bhatia, A. et al. Cloning, characterization and serological evaluation of K9 and K26: Two related hydrophilic antigens of Leishmania chagasi. Mol. Biochem. Parasitol. 102, 249, 1999. 38. Padilla, C.R., and Montoya, Y.P. Caracterización e inmunoreactividad de la proteína acídica ribosomal P2β de L. (V.) braziliensis. Rev. Per. Med. Exp. Salud Publica. 20, 92, 2003. 39. Amorin, A.G. et al. Identi�cation of the C-terminal region of 70 kDa heat shock protein from Leishmania (Viannia) braziliensis a target for the humoral immune response. Cell Stress Chaperones 1, 177, 1996. 40. Singh, S. et al. Diagnostic and prognostic value of K39 recombinant antigen in Indian leishmaniasis J. Parasitol. 81, 1000, 1995. 41. Singh, S., Kumasi, V., and Singh, N. Predicting kala-azar disease manifestations in asymptomatic patients with latent Leishmania donovani infection by detection of antibody against recombinant K39 antigen. Clin. Diagn. Lab. Immunol. 9, 568, 2002.
42. Marín, C. et al. Leishmania infantum: Use of excreted superoxide dismutase in ELISA and Western blotting for serologic diagnostic in naturally infected dogs. Parasitol. Res. 101, 801, 2007.
43. Marín, C. et al. Enzyme-linked immunosorbent assay for superoxide dismutase-excreted antigen in diagnosis of sylvatic and andean cutaneous leishmaniasis of Peru. Am. J. Trop. Med. Hyg. 80, 55, 2009.
44. McCord, J.M., and Fridovich, I., Superoxide dismutase: An enzymatic function in erythrocuprein (hemocupresin). J. Biol. Chem. 244, 6049, 1969.
45. Marín, C., and Sánchez-Moreno, M., The trypanosomiasis diagnosis: A review on the role of the superoxide dismutase as molecular marker. Curr. Res. Microbiol. I, 1, 2009.
46. Marty, P.A. et al. Detection by Westem blot of four antigens characterizing acute clinical leishmaniasis due to Leishmania infantum. Trans. R. Soc. Trop. Med. Hyg. 89, 690, 1995.
47. Mary, C.D. et al. Westem blot ana1ysis of antibodies to Leishmania infantum antigens: Potential of the 14-kD and 16-kD antigens for diagnosis and epidemiologic purposes. Am. J. Trop. Med. Hyg. 47, 764, 1992.
48. Montoya, Y. et al. Recombinant antigens for speci�c and sensitive serodiagnosis of Latin American tegumentary leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 91, 674, 1997.
49. Brito, M.E. et al. Identi�cation of potentially diagnostic Leishmania braziliensis antigens in human cutaneous leishmaniasis by immunoblot análysis. Clin. Diagn. Lab. Immunol. 7, 318, 2000.
50. Senaldi, G. et al. Serological diagnosis of visceral leishmaniasis by a dot-enzyme immunoassay for the detection of a Leishmania donovani-related circulating antigen. J. Immunol. Methods. 193, 9, 2001.
51. Attar, Z.J. et al. Latex agglutination test for the detection of urinary antigens in visceral leishmaniasjs. Acta Trop. 78, 11, 2001.
52. Gavgani, A.S.M., KhademVatan, S., and Ghazanchael, A. KAtex antigen-detection test as a diagnosis tool for latent visceral leishmaniasis cases. African J. Biotechnol. 7, 852, 2008.
53. Tavares, C.A.P., Fernandez, A.P., and Melo, M.N. Molecular diagnosis of leishmaniasis. Expert Rev. Mol. Diagn. 3, 657, 2003. 54. Schallig, H.D., and Oskam, L. Molecu1ar bio1ogical applications in the diagnosis and control of leishmaniasis and parasite identi�cation. Trop. Med. Int. Health. 7, 641, 2002.
55. Gangneux, J.P. et al. Prospective value of PCR ampli�cation and sequencing for diagnosis and typing of Old World Leishmania infections in an area of nonendemicity. J. Clin. Microbiol. 41, 1419, 2003.
56. da Silva, E.S. et al. Diagnosis of human visceral leishmaniasis by PCR using blood samples spotted on �lter paper. Genet. Mol. Res. 3, 251, 2004.
57. Singh, S., Dey, A., and Sivakumar, R. Applications of molecular methods for Leishmania control. Exp. Rev. Mol. Diagn. 5, 251, 2005.
58. De Doncker, S. et al. A new PCR-ELISA for diagnosis of visceral leishmaniasis in blood of HIV-negative subjects. Trans. R. Soc. Trop. Med. Hyg. 99, 25, 2005.
59. Kuboki, N. et al. Loop-mediated isothermal ampli�cation for detection of African trypanosomes. J. Clin. Microbiol. 41, 5517, 2003. 60. Deborggraeve, S. et al. Molecular dipstick test for diagnosis of sleeping sickness. J. Clin. Microbiol. 44, 2884, 2006. 61. Garcia, A.L. et al. American tegumentary leishmaniasis: Antigen-gene polymorphism, taxonomy and clinical pleomorphism. Infect. Genet. Evol. 5, 109, 2005. 62. Strauss-Ayali, D. et al. Polymerase chain reaction using non invasively obtained samples, for the detection of Leishmania infantum DNA in dogs. J. Infect. Dis. 189, l720, 2004. 63. Maurya, R. et al. Evaluation of PCR for diagnosis of Indian kala-azar and assessment of cure. J. Clin. Microbiol. 43, 3038, 2005. 64. Mary, C. et al. Quanti�cation of Leishmania infantum DNA by a real-time PCR assay with high sensitivity. J. Clin. Microbiol. 42, 524, 2004. 65. Martin-Sanchez, J. et al. Detection of Leishmania infantum kinetoplast DNA in peripheral blood from asymptomatic individuals at risk for parenterally transmitted infections: Relationship between polymerase chain reaction results and other Leishmania infection markers. Am. J. Trop. Med. Hyg. 70, 545, 2004. 66. Otero, A.C. et al. Occurrence of Leishmania donovani DNA in donated b1ood from seroreactive Brazi1ian blood donors. Am. J. Trop. Med. Hyg. 62, 128, 2000. 67. de Oliveira, C.I. et al. Clinical utility of polymerase chain reaction-based detection of Leishmania in the diagnosis of American cutaneous leishmaniasis. Clin. Infect. Dis. 37, 49, 2003. 68. Mimori, T. et al. Usefulness of sampling with cotton swab for PCR-diagnosis of cutaneous leishmaniasis in the New World. Acta Trop. 81, 197 (2002) 69. Santiago, M.A. et al. Differentiation between canine cutaneous and visceral leishmaniasis by the detection of immunoglobulin G speci�c for Leishmania (Viannia) braziliensis and Leishmania (Leishmania) chagasi antigens using �ow cytometry. Vet. Parasitol. 154, 341, 2008. 70. Jaffe, L., and Zalis, M., Puri�cation of two Leishmania donovani membrane proteins recognized by sera from patients with visceral leishmaniasis. Mol. Biochem. Parasitol. 27, 53, 1988. 71. Rosário, E.Y. et al. Evaluation of enzyme-linked immunosorbent assay using crude Leishmania and recombinant antigens as a diagnostic marker for canine visceral leishmaniasis. Mem Inst Oswaldo Cruz 100, 197, 2005. 72. Arora, S.K., and Sehgal, S. Identi�cation of major antigens of Leishmania donovani using kala-azar sera. Med. Microbiol. Immunol. 178, 81, 1989. 73. Tonui, W. K. et al. Immunization with Leishmania major exogenous antigens protects susceptible BALB/c mice against challenge infection with L. major. Infect. Immun. 72, 5654, 2004. 74. Mettler, M. et al. Evaluation of enzyme-linked immunosorbent assays, an immuno�uorescent-antibody test, and two rapid tests (immunochromatographic-dipstick and gel tests) for serological diagnosis of symptomatic and asymptomatic Leishmania infections in dogs. J. Clin. Microbiol. 43, 5515, 2005. 75. Barbosa-de-Deus, R. et al. Leishmania major-like antigen for speci�c and sensitive serodiagnosis of human and canine visceral leishmaniasis. Clin. Diagn. Lab. Immunol. 9, 1361, 2002. 76. Rafati, S. et al. Biochemical analysis and immunogenicity of Leishmania major amastigote fractions in cutaneous leishmaniasis. Clin. Exp. Immunol. 110, 203, 1997.
77. Atta, A.M. et al. Antileishmanial IgG and IgE antibodies recognize predominantly carbohydrate epitopes of glycosylated antigens in visceral leishmaniasis. Mem. Inst. Oswaldo Cruz 99, 525, 2004.
78. Kamoun-Essghaier, S. et al. Proteomic approach for characterization of immunodominant membrane-associated 30- to 36-kilodalton fraction antigens of Leishmania infantum promastigotes, reacting with sera from Mediterranean visceral leishmaniasis patients. Clin. Diagn. Lab. Immunol. 12, 310, 2005.
79. Kabiri A.R. et al. Leishmanin skin test in guinea pig with a single puri�ed protein of Leishmania major. Exp. Parasitol. 111, 239–243, 2005.
80. Zeinali, M., Ardestani, S.K., and Kariminia, A. Puri�cation and biochemical characterization of two novel antigens from Leishmania major promastigotes. Korean J. Parasitol. 45, 287, 2007.
81. Ribeiro, F.C. et al. Use of ELISA employing Leishmania (Viannia) braziliensis and Leishmania (Leishmania) chagasi antigens for the detection of IgG and IgG1 and IgG2 subclasses in the diagnosis of American tegumentary leishmaniasis in dogs. Vet. Parasitol. 148, 200, 2007.
82. Gomes-Silva et al. Serological reactivity of different antigenic preparations of Leishmania (Leishmania) amazonensis and the Leishmania braziliensis complex. Rev. Soc. Bras. Med. Trop. 41, 135, 2008.
83. Choudhury, A. et al. Enzyme linked immunosorbant assay in diagnosis of kala-azar in Bhadohi (Varanasi) India. Trans. R. Soc. Trop. Med. Hyg. 84, 363, 1990.
84. Garg, R. et al. Leishmania donovani: Identi�cation of stimulatory soluble antigenic proteins using cured human and hamster lymphocytes for their prophylactic potential against visceral leishmaniasis. Vaccine 24, 2900, 2006.
85. Gupta, S.K. et al. Proteomic approach for identi�cation and characterization of novel immunostimulatory proteins from soluble antigens of Leishmania donovani promastigotes. Proteomics 7, 816, 2007.
86. Jaffe, C.L., and McMahon-Pratt, D., Monoclonal antibodies speci�c for Leishmania tropica I. Characterization of antigens associated with stage- and species-speci�c determinants. J. Immunol. 131, 1987, 1983.
87. Tebourski, F. et al. Identi�cation of an immunodominant 32-kilodalton membrane protein of Leishmania donovani infantum promastigotes suitable for speci�c diagnosis of Mediterranean visceral leishmaniasis J. Clin. Microbiol. 32, 2474, 1994.
88. Kamoun-Essghaier, S. et al. Proteomic approach for characterization of immunodominant membrane-associated 30 to 36 kDa fraction antigens of Leishmania infantum promastigotes reacting with sera from Mediterranean visceral leishmaniasis patients. Clin. Diagn. Lab. Immunol. 12, 310, 2005.
89. Lawrence, F., and Robert-Gero, M. Induction of heat shock and stress proteins in promastigotes of three Leishmania species. PNAS USA 82, 4414, 1985.
90. Nandan, D. et al. A rapid, single-step puri�cation method for immunogenic members of the hsp 70 family: Validation and application. J. Immunol. Methods 176, 255, 1994.
91. Jeronimo, S.M. et al. Identi�cation of Leishmania chagasi antigens recognized by human lymphocytes J. Infect. Dis. 172, 1055, 1995.
92. Ko, Y.G., and Thompson, G.A. Puri�cation of glycosylphosphatidylinositol-anchored proteins by modi�ed Triton X-114 partitioning and preparative gel electrophoresis. Anal. Biochem. 224, 166, 1995. 93. Añez-Rojas, N. et al. Isolation, puri�cation and characterization of GPI-anchored membrane proteins from Trypanosoma rangeli and Trypanosoma cruzi. Acta Trop. 97, 140, 2006. 94. Mukherjee, M., Bhattacharyya, A., and Duttagupta, S. Serodiagnostic and immunoprophylactic potential of a 78 kDa protein of Leishmania donovani of Indian origin. Med Sci Monit. 8, 117, 2002. 95. Rajasekariah, G-H.R. et al. A novel exo-antigen-based ELISA for the detection of canine leishmaniasis. Am. J. Trop. Med. Hyg. 78, 616, 2008. 96. Chenik, M. et al. Approaches for the identi�cation of potential excreted/secreted proteins of Leishmania major parasites. Parasitology 132, 493, 2006. 97. Kubar, J., and Fragaki, K. Recombinant DNA-derived Leishmania proteins: From the laboratory to the �eld. Lancet Infect. Dis. 5, 107, 2005. 98. Kubar, J., and Fragaki, K. Leishmania proteins derived from recombinant DNA: Current status and next steps. Trends Parasitol. 22, 111, 2006. 99. Rafati, A.H. et al. Identi�cation of Leishmania major cysteine proteinases as targets of the immune response in humans. Mol. Biochem. Parasitol. 113, 15, 2001. 100. Pinheiro, P.H. et al. A recombinant cysteine proteinase from Leishmania (Leishmania) chagasi as an antigen for delayedtype hypersensitivity assays and serodiagnosis of canine visceral leishmaniasis. Vet. Parasitol. 26(162), 32, 2009. 101. Burns, J.M. et al. Molecular characterization of a kinesinrelated antigen of Leishmania chagasi that detects speci�c antibody in African and American visceral leishmaniasis. Proc. Natl. Acad. Sci., USA 90, 775, 1993. 102. Fragaki, K. et al. A novel Leishmania infantum nuclear phosphoprotein Lepp12 which stimulates IL1-beta synthesis in THP-1 transfectants. BMC Microbiol. 30, 3, 2003. 103. Suf�a, B. et al. A novel Leishmania infantum recombinant antigen which elicits interleukin 10 production by peripheral blood mononuclear cells of patients with visceral leishmaniasis. Infect. Immun. 68, 630, 2000. 104. Sambrook, J., Fritsch, E.F., and Maniatis, T. Molecular Cloning. A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, 1989. 105. Handman, B., Button, L.L., and McMaster RW, Leishmania major: Production of recombinant gp63, its antigenicity and immunogenicity in mice. Exp. Parasitol. 70, 427, 1990. 106. Soto, M. et al. Multicomponent chimeric antigen for serodiagnosis of canine visceral leishmaniasis. J. Clin. Microbiol. 36, 58, 1998. 107. Gonçalves, A.M., Nehme, N.S., and Morel, C.M. Trypanosomatid characterization by schizodeme analysis, In: Morel, C.M. (ed.), Genes and Antigens of Parasites (A Laboratory Manual). Fundaçao Oswaldo Cruz, Rio de Janeiro, RJ, Brazil, 1984. 108. Sokal, J.E. Measurement of delayed skin-test responses. N. Engl. J. Med. 293, 501, 1975. 109. Gramiccia, M. et al. Leishmaniasis in Sardinia. 5. Leishmanin reaction in the human population of a focus of low endemicity of canine leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 84, 371, 1990. 110. Stocker, J.W., and Heusser, C.H. Methods for binding cells to plastic: Application to a solid-phase radioimmunoassay for cell-surface antigens. J. Immunol. Methods 26, 87, 1979. 111. Bañuls, A.L., Hide, M., and Prugnolle, F. Leishmania and the Leishmaniases: A parasite genetic update and advances in taxonomy, epidemiology and pathogenicity in humans. Adv. Parasitol. 64, 1, 2007.
112. Pampiglione, S. et al. Studies in Mediterranean leishmaniasis. 3. The leishmanin skin test in kala-azar. Trans. R. Soc. Trop. Med. Hyg. 69, 60–68, 1975.
113. Scott, P. et al. Vaccination against cutaneous leishmaniasis in murine model II. Immunological properties of protective and non protective subfractions of a soluble promastigotes extract. J. Immunol. 139, 3118–3125, 1987. 114. Rojas, A. et al. Isolation, puri�cation, characterization and antigenic evaluation of GPI-anchored membrane proteins from Leishmania (Viannia) braziliensis. Acta Trop. 105, 139–144, 2008. 115. Simpson R.J. et al. Peptide mapping and internal sequencing of proteins electroblotted from two-dimentional gels onto polyvinylidene di�uoride membranes. A chromatographic procedure for separating proteins from detergents. J Chromatogr. 476, 345–361, 1989. 9 Chapter 9 - Naegleria
4. De Jonckheere, J.F. Molecular de�nition and ubiquity of species in the genus Naegleria. Protist 155, 89, 2004.
5. De Jonckheere, J.F. Key note lecture: Molecular taxonomy of the genus Naegleria and other vahlkamp�ids. XIth International Meeting on the Biology and Pathogenicity of Free-Living Amoebae Proceedings, Ceské Budejovice, Czech Republic, 2005.
6. Schuster, F. An electron microscope study of the amoebo�agellate, Naegleria gruberi (Schardinger) I. The ameboid and �agellate stage. J. Protozool. 10, 297, 1963.
7. Fulton, C., and Dingle, A.D. Basal bodies but not centrioles, in Naegleria. J. Cell Biol. 51, 826, 1971.
8. Carter, R.F. Description of a Naegleria sp. isolated from two cases of primary amoebic meningoencephalitis, and of the experimental pathological changes induced by it. J. Pathol. 100, 217, 1970.
9. Feldman, M.R. Naegleria fowleri. Fine structural localization of acid phosphatase and heme proteins. Exp. Parasitol. 41, 290, 1977.
10. Patterson, M. et al. Ultrastructure of Naegleria fowleri en�agellation. J. Bacteriol. 147, 217, 1981.
11. Marshall, M.M. et al. Waterborne protozoan pathogens. Clin. Microbiol. Rev. 10, 67, 1997.
12. Werth, J.M., and Kahn A.J. Isolation and preliminary chemical analysis of the cyst wall of the amoeba-�agellate Naegleria gruberi. J. Bacteriol. 94, 1272, 1967.
13. Fulton, C., and Guerrini, A.M. Mitotic synchrony in Naegleria ameba. Exp. Cell Res. 56, 194, 1969.
14. Willmer, E.N. Factors which in�uence the acquisition of �agella by the amoeba. Naegleria gruberi. J. Exp. Biol. 33, 583, 1956.
15. De Jonckheere, J.F., and Brown, S. SSU rDNA analysis reveals the existence of another Naegleria sp. with dividing �agellates: N. robinsoni sp. nov. Eur. J. Protisol. 35, 264, 1999.
16. De Jonckheere, J.F. et al. The amoeba-to-�agellate transformation test is not reliable for the diagnosis of the genus Naegleria. Description of three new Naegleria spp. Protist. 152, 115, 2001.
17. John, D.T., Cole, T.B., and John, R.A. Flagella number among Naegleria �agellates. Folia Parasitol. 38, 289, 1991.
18. Singh, B.N., and Das, S.R. Intranasal infection of mice with �agellate stage of Naegleria aerobia and its bearing on the epidemiology of human meningo-encephalitis. Curr. Sci. 41, 625, 1972.
19. Kyle, D.E., and Noblet, A.D. Vertical distribution of potentially pathogenic free-living amoebae in freshwater lakes. J. Protozool. 32, 99, 1985.
20. Grif�n, J.L. The pathogenic amoebo�agellate Naegleria fowleri environmental isolations, competitors, ecologic interactions, and the �agellate-empty hypothesis. J. Protozool. 30, 403, 1983.
21. Lares-Villa F. A quantitative study of thermophilic Naegleria during summer season, in natural waters used for recreational purposes in Sonora, México. XIth International Meeting on the Biology and Pathogenicity of Free-Living Amoebae Proceedings, Ceské Budejovice, Czech Republic, 2005.
22. Cabanes, P.A. et al. Assessing the risk of primary amoebic meningoencephalitis from swimming in the presence of environmental Naegleria fowleri. Appl. Environ. Microbiol. 67, 2927, 2001.
23. Lugg, R., and Psaila-Savona, P. A legal standard for thermophilic Naegleria in freshwater body in Western Australia. IXth International Meeting on the Biology and Pathogenicity of Free-Living Amoebae Proceedings, Paris, France, p. 291, 2001. 24. Newsome, A.L. et al. Interaction between Naegleria fowleri and Legionella pneumophilia. Infect. Immun. 50, 449, 1985. 25. Jarolim, K.L., McCosh, J.K., and Howard, M.J. The role of blood vessels and lungs in the dissemination of Naegleria fowleri following intranasal inoculation in mice. Folia Parasitol. 49, 183, 2002. 26. Jarolim, K.L. et al. A light microscopy study of the migration of N. fowleri from the nasal submucosa to the central nervous system during the early stage of primary amebic meningoencephalitis in mice. J. Parasitol. 86, 50, 2000. 27. Cervantes-Sandoval, I. et al. Characterization of brain in�ammation during primary amoebic meningoencephalitis. Parasitol. Int. 57, 307, 2008. 28. Galea, I., Bechmann, I., and Perry, V.H. What is immune privilege (not)? Trends. Immunol. 28, 12, 2007. 29. Andersson, P.B., Perry, V.H., and Gordon, S. The acute in�ammatory response to lipopolysaccharide in CNS parenchyma differs from that in other body tissues. Neuroscience, 48, 169, 1992. 30. Andersson, P.B., Perry, V.H., and Gordon, S. Intracerebral injection of proin�ammatory cytokines or leukocyte chemotaxins induces minimal myelomonocytic cell recruitment to the parenchyma of the central nervous system. J. Exp. Med. 176, 255, 1992. 31. Oh, Y.H. et al. Cytopathic changes and pro-in�ammatory cytokines induced by Naegleria fowleri trophozoites in rat microglial cells and protective effects of an anti-Nfa1 antibody. Parasite. Immunol. 27, 453, 2006. 32. De Jonckheere, J.F., Characterization of Naegleria species by restriction endonuclease digestion of whole-cell DNA. Mol. Biochem. Parasitol. 24, 55, 1987. 33. McLaughlin, G.L., Brandt, F.H., and Visvesvara, G.S. Restriction fragment length polymorphisms of the DNA of the selected Naegleria and Acanthamoeba amoebae. J. Clin. Microbiol. 26, 1655, 1988. 34. Kilvington, S., and Beeching, J. Detection of a novel restriction fragment length polymorphism type in Naegleria fowleri (free-living amoeba) isolates from electricity power stations in England and France. Eur. J. Protisol. 33, 186, 1997. 35. De Jonckheere, J.F. Isoenzyme patterns of pathogenic and non-pathogenic Naegleria spp using agarose gel isoelectrofocusing. Ann. Microbiol. 133A, 319, 1982. 36. Moss, D.M. et al. High- resolution polyacrylamide gradient gel electrophoresis (PGGE) of isoenzymes from Naegleria species. J. Protozool. 38, 26, 1988. 37. John, D.T., and Nussbaum, S.L. Naegleria fowleri infection acquired by mice through swimming in amebae-cantaminated water. J. Parasitol. 69, 871, 1983. 38. Schuster, F.L., and Visvesvara G.S. Free-living amoeba as opportunistic and non-opportunistic pathogens of humans and animals. Int. J. Parasitol. 34, 1001, 2004. 39. Mckee, T. et al. Primary amoebic meningoencephalitisGeorgia, 2002. MMWR 52, 962, 2003. 40. Lares-Villa, F. Free-living amoebae infections in Mexico. IXth International Meeting on the Biology and Pathogenicity of Free-Living Amoebae Proceedings, Paris, France, p. 13, 2001. 41. Hannisch, W., and Hallagan, L.F. Primary amebic meningoencephalitis: A review of the clinical literature. Wild. Environ. Med. 8, 211, 1997. 42. Schumacher, D.J., Tien, R.D., and Lane, K. Neuroimaging �ndings in rare amebic infections of the central nervous system. Am. J. Neuroradiol. 16, 930, 1993. 43. Visvesvara, G.S., Moura, H., and Schuster F.L. Pathogenic opportunistic free-living amoebae: Acantamoeba spp., Balamuthia mandrillaris, Naegleria fowleri, and Sappinia diploidea. FEMS Immunol. Med. Microbiol. 50, 1, 2007.
44. Kidney, D.D., and Kim, S.H. CNS infections with free-living amebas: Neuroimaging �ndings. Am. J. Roentgenol. 17, 809, 1998.
45. Martinez, A.J. Free-Living Amebas: Natural History, Prevention, Diagnosis, Pathology and Treatment of Disease, CRC Press, Boca Raton, FL, 156pp, 1985.
46. Visvesvara, G.S., and Maguire, J.H. Pathogenic and opportunistic free-living amebas: Acanthamoeba spp., Balamuthia mandrillaris, Naegleria fowleri, and Sappinia diploidea. In Guerrant, R.L., Walker, D.H., and Weller, P.F. (eds), Tropical Infectious Diseases. Principles, Pathogens, & Practice, Vol. 2, Churchill Livingstone, Philadelphia, PA, pp. 1114–11125, 2006.
47. Behets, J. et al. Detection of Naegleria spp. and Naegleria fowleri: A comparison of �agellation tests, ELISA and PCR. Water. Sci. Technol. 47, 117, 2003.
48. Sparagano, O. et al. Use of monoclonal antibodies to distinguish pathogenic Naegleria fowleri (cysts, trophozoites, or �agellate forms) from other Naegleria species. J. Clin. Microbiol. 31, 2758, 1993.
49. Pelandakis, M., Serre, S., and Pernin, P. Analysis of the 5.8S rRNA gene and the internal transcribed spacers in Naegleria spp. and in N. fowleri. J. Eukaryot, Microbiol. 47, 116, 2000.
50. Réveiller, F., Cabanes, P.A., and Marciano-Cabral, F. Development of a nested PCR assay to detect the pathogenic free-living amoeba Naegleria fowleri. Parasitol. Res. 88, 443, 2002.
51. Qvarnstrom, Y. et al. Multiplex real-time PCR assay for simultaneous detection of Acanthamoeba spp., Balamuthia mandrillaris, and Naegleria fowleri. J. Clin. Microbiol. 44, 3589, 2006.
52. Behets, J. et al. A duplex real-time PCR assay for the quantitative detection of Naegleria fowleri in water samples. Water. Res. 41, 118, 2007.
53. Schild, M. et al. PCR-based diagnosis of Naegleria sp. Infection in formalin-�xed and paraf�n embedded brain sections. J. Clin. Microbiol. 45, 564, 2007.
54. Cervantes-Sandoval, I. et al. Characterization of Naegleria fowleri strains isolated from human cases of primary amoebic meningoencephalitis in Mexico. Rev. Invest. Clin. 59, 342, 2007. 55. Marciano-Cabral, F., and Cabral, G. The immune response to Naegleria fowleri amebae and pathogenesis of infection. FEMS Immunol. Med. Microbiol. 51, 116, 2007. 56. Martínez, A.J., and Visvesvara, G.S. Laboratory diagnosis of pathogen free-living amoebas: Naegleria, Acanthamoeba, and Leptomyxid. Clin. Lab. Med. 11, 861, 1991. 57. Da Rocha-Azevedo, B., Tanowitz, H.B., and MarcianoCabral, F. Diagnosis of infections caused by pathogenic free-living amoebae. Interdiscip. Perspect. Infect. Dis. 2009, 251406, 2009. 58. Kilvington, S., and Beeching, J. Identi�cation and epidemiological typing of Naegleria fowleri with DNA probes. Appl. Environ. Microbiol. 61, 2071, 1995. 59. Zhou, L. et al. Genetic variations in the internal transcribed spacer and mitochondrial small subunit rRNA gene of Naegleria spp. J. Eukaryot. Microbiol. 50, 522, 2003. 60. De Jonckheere, J.F. Sequence variation in the ribosomal internal transcribed spacer, including 5.8S, of Naegleria spp. Protist 149, 221, 1998. 61. Robinson, B.S., Monis, P.T., and Dobson, P.J. Rapid, sensitive, and discriminating identi�cation of Naegleria spp. by real-time PCR and melting-curve analysis. Appl. Environ. Microbiol. 72, 5857, 2006. 62. Moura, H. et al. Detection of protein �ngerprints of emerging and opportunistic pathogens by MALDI-TOF and proteome database search. Am. J. Mass Spectrom. 14, S615, 2003. 63. Guarner, J. et al. Histopathologic spectrum and immunohistochemical diagnosis of amebic meningoencephalitis. Mod. Pathol. 20, 1230, 2007. 64. Chirgwin, J.M. et al. Isolation of biologically active ribonucleic acid from sources enriched in ribonuclease. Biochemistry 18, 5294, 1979. 65. Aviv, H., and Leder, P. Puri�cation and properties of biologically active globin messenger RNA by chromatography on oligothymidylic acid-cellulose. Proc. Natl Acad. Sci. USA, 69, 1408, 1972. 66. Behets, J. et al. Quantitative detection and differentiation of free-living amoeba species using SYBR Green-based real time PCR melting curve analysis. Curr. Microbiol. 53, 506, 2006. 10 Chapter 10 - Pentatrichomonas
7. Tang, P., and Hsin-Sheng L. Pentatrichomonas hominis: Purine salvage pathway. Comp. Biochem. Physiol. 90B, 419, 1988.
8. Felleisen, R.S. Comparative sequence analysis of 5.8S rRNA genes and internal transcribed spacer (ITS) regions of trichomonadid protozoa. Parasitology 115, 111, 1997.
9. Honigberg, B.M. Trichomonads found outside the urogenital tract of humans. In: Honigberg, B.M. (ed.), Trichomonads Parasitic in Humans, Chapter 19. Springer-Verlag, New York, 1990.
10. Giboda, M. Intestinal parasites in Kampuchea, Takeo province. J. Hyg. Epidemiol. Microbiol. Immunol. 29, 377, 1985.
11. Cancrini, G. et al. Intestinal parasites in the Camiri, Gutierrez and Boyuibe areas, Santa Cruz Department, Bolivia. Parasitologia 30, 263, 1988.
12. Subbannayya, K. et al. Entamoeba histolytica and other parasitic infections in south Kanara district, Karnataka. J. Commun. Dis. 21, 207, 1989.
13. Ali-Shtayeh, M.S. et al. Prevalence and seasonal �uctuations of intestinal parasitic infections in the Nablus area, West Bank of Jordan. Ann. Trop. Med. Parasitol. 83, 67, 1989.
14. Yang, C.-R. et al. Diarrhoea surveillance in children aged under 5 years in a rural area of Hebei province, China. J. Diarrhoeal Dis. Res. 8, 155, 1990.
15. Chunge, R.N. et al. Longitudinal study of young children in Kenya: Intestinal parasitic infection with special reference to Giardia lamblia, its prevalence, incidence and duration, and its association with diarrhoea and with other parasites. Acta Trop. 50, 39, 1991.
16. Okafor, C.N., and Azubike, C.N. Studies in intestinal parasitic disease agents in stools of people in a rural area of Nigeria. West. Afr. J. Med. 11, 106, 1992.
17. Ogunsanya, T.I., Rotimi, V.O., and Adenuga, A. A study of the aetiological agents of childhood diarrhoea in Lagos, Nigeria. J. Med. Microbiol. 40, 10, 1994. 18. Henry, M.C. et al. Community survey of diarrhoea in children under 5 years in Kinshasa, Zaire. Ann. Soc. Belg. Med. Trop. 75, 105, 1995.
19. Sherchand, J.B., Larsson, S., and Shrestha, M.P. Intestinal parasites in children and adults with and without abdominal discomfort from the Kathmandu area of Nepal. Trop. Gastroenterol. 17, 15, 1996.
20. Saksirisampant, W. et al. Intestinal parasitic infestations among children in an orphanage in Pathum Thani province, J. Med. Assoc. Thai. 86, S263, 2003.
21. Addy, P.A., Antepim, G., and Frimpong, E.H. Prevalence of pathogenic Escherichia coli and parasites in infants with diarrhoea in Kumasi, Ghana, East. Afr. Med. J. 81, 353, 2004.
22. Degerli, S., Ozcelik, S., and Celiksoz, A. The distribution of intestinal parasites in patients presenting at the Parasitology Laboratory of the Cumhuriyet University. Turkiye Parazitol. Derg. 29, 116, 2005.
23. Culha, G. The distribution of patients with intestinal parasites presenting at the Parasitology Laboratory of the Mustafa Kemal University Medical Faculty. Turkiye Parazitol. Derg. 30, 302, 2006.
24. Celik, T. et al. Incidence of intestinal parasites among primary school children in Malatya. Turkiye Parazitol. Derg. 30, 35, 2006.
25. El Shazly, A.M. et al. Intestinal parasites in Dakahlia governorate, with different techniques in diagnosing protozoa. J. Egypt. Soc. Parasitol. 36, 1023, 2006.
26. Alaofé, H. et al. Intestinal parasitic infections in adolescent girls from two boarding schools in southern Benin. Trans. R. Soc. Trop. Med. Hyg. 103, 452, 2008. 27. Haghighi, A. et al. Frequency of enteric protozoan parasites among patients with gastrointestinal complaints in medical centers of Zahedan, Iran. Trans. R. Soc. Trop. Med. Hyg. 103, 452, 2009. 28. Nasiri, V. et al. Intestinal parasitic infections among inhabitants of Karaj city, Tehran Province, Iran in 2006–2008. Korean J. Parasitol. 47, 265, 2009. 29. Mancilla-Ramirez, J., and Gonzalez-Yunes, R. Diarrhea associated with Trichomonas hominis in a newborn infant. Bol. Med. Hosp. Infant. Mex. 46, 623, 1989. 30. Chunge, C.N. et al. Pathogenicity of Trichomonas hominis in Kenya. East. Afr. Med. J. 65, 532, 1988. 31. Hegner, R. Differential reactions of species and strains of trichomonad �agellates to changes of the environment. Am. J. Hyg. 15, 513, 1932. 32. Khan, A.R., and Huq, F. Disease agents carried by �ies in Dacca city. Bangladesh Med. Res. Counc. Bull. 4, 86, 1978. 33. Caruso, G. Human trichomoniasis. Acta Med. Ital. Mal. Infett. Parassit. 15, 231, 1960. 34. Dobbel, C. Research on the intestinal protozoa of monkeys and man. VI Experiments with the trichomonads of man and macaques. Parasitology 24, 531, 1934. 35. Jongwutiwes, S., Silachamroon, U., and Putaporntip, C. Pentatrichomonas hominis in empyema thoracis, Trans. R. Soc. Trop. Med. Hyg. 94, 185, 2000. 36. Masur, H., Hook, E. III, and Armstrong, D.A. Trichomonas species in a mixed microbial meningitis. JAMA 236, 1978, 1976. 37. Jakobsen, E.B., Friis-Moller, A., and Friis, J. Trichomonas species in a subhepatic abscess. Eur. J. Clin. Microbiol. 6, 296 (1987). 38. Crucitti, T. et al. Detection of Pentatrichomonas hominis DNA in biological specimens by PCR. Lett. Appl. Microbiol. 38, 510, 2004. 39. Stabler, R.M., Feo, L.G., and Rakoff, A.E. Implantation of intestinal trichomonads (T. hominis) into the human vagina. Am. J. Trop. Med. Hyg. 34, 114, 1941. 40. Stabler, R.M., and Feo, L.G. Survival time of intravaginally implanted Trichomonas hominis. Am. J. Trop. Med. Hyg. 22, 633, 1942. 41. Crucitti, T. et al. Trichomonas vaginalis is highly prevalent in adolescent girls, pregnant women, and commercial sex workers in Ndola, Zambia. Sex. Transm. Dis. 37, 223, 2010. 42. Gooikin, J.L. et al. Molecular characterization of Trichomonads from feces of dogs with diarrhea. J. Parasitol. 91, 939, 2005. 43. Gookin, J.L., Stauffer, S.H., and Levy, M.G. Identi�cation of Pentatrichomonas hominis in feline fecal samples by polymerase chain reaction assay. Vet. Parasitol. 145, 11, 2007. 44. Gookin, J.L. et al. Optimization of a species-speci�c polymerase chain reaction assay for identi�cation of Pentatrichomonas hominis in canine fecal specimens. Am. J. Vet. Res. 68, 783, 2007. 45. Shaio, M.-F., and Chen, J.-R. Immunoglobulin M-dependent classical complement pathway activation in killing of Pentatrichomonas hominis. Infect. Immun. 57, 902, 1989. 46. Lo, H.S., and Shaio, M. A method for the isolation of Trichomonas hominis in axenic culture. Ann. Trop. Med. Parasitol. 75, 567, 1981. 47. Romatowski, J. Pentatrichomonas hominis infection in four kittens. JAVMA 8, 1270, 2000. 48. Levy, M.G. et al. Tritrichomonas foetus and not Pentatrichomonas hominis is the etiologic agent of feline trichomonal diarrheae. J. Parasitol. 89, 99, 2003. 11 Chapter 11 - Trichomonas
17. Fiori, P.L., Rappelli, P., and Addis, M.F., The �agellated parasite Trichomonas vaginalis: New insights into cytopathogenicity mechanisms. Microb. Pathog. 1, 149, 1999.
18. Graves, A., and Gardner, W.A., Pathogenicity of Trichomonas vaginalis. Clin. Obstet. Gynecol. 36, 145, 1993.
19. de Carli, G.A. et al., Laboratory diagnosis of Trichomonas vaginalis among women attending a venereal diseases control division. First report. Rev. Latinoam. Microbiol. 29, 301, 1987.
20. Fouts, A.C., and Kraus, S.J., Trichomonas vaginalis: Reevaluation of its clinical presentation and laboratory diagnosis. J. Infect. Dis. 141, 137, 1993.
21. Weinberger, M.W., and Harger, J.H., Accuracy of the Papanicolaou smear in the diagnosis of asymptomatic infection with Trichomonas vaginalis. Obstet. Gynecol. 82, 425, 1993.
22. Schwebke, J.R., Morgan, S.C., and Pinson, G.B., Validity of self-obtained vaginal specimens for diagnosis of trichomoniasis. J. Clin. Microbiol. 35, 1618, 1997.
23. Schwebke, J., Venglarik, M., and Morgan, S., Delayed versus immediate bedside inoculation of culture media for diagnosis of vaginal trichomonosis. J. Clin. Microbiol. 37, 2369, 1999.
24. Tabrizi, S.N. et al., A self-administered technique for the detection of sexually transmitted diseases in remote communities. J. Infect. Dis. 176, 289, 1997.
25. Wiese, W. et al., A meta-analysis of the Papanicolaou smear and wet mount for the diagnosis of vaginal trichomoniasis. Am. J. Med. 108, 301, 2000.
26. Kissinger, P.J. et al., Vaginal swabs versus lavage for detection of Trichomonas vaginalis and bacterial vaginosis among HIVpositive women. Sex. Transm. Dis. 32, 227, 2005.
27. Garber, G.E. et al., Cell culture compared with broth for detection of Trichomonas vaginalis. J. Clin. Microbiol. 25, 1275, 1987. 28. Krieger, J.N. et al., Diagnosis of trichomoniasis. Comparison of conventional wet-mount examination with cytologic studies, cultures, and monoclonal antibody staining of direct specimens. JAMA 259, 1223, 1988.
29. Gelbart, S.M. et al., Comparison of Diamond’s medium modi�ed and Kupferberg medium for detection of Trichomonas vaginalis. J. Clin. Microbiol. 27, 1095, 1989.
30. Schmid, G.P. et al., Evaluation of six media for the growth of Trichomonas vaginalis from vaginal secretions. J. Clin. Microbiol. 27, 1230, 1989.
31. Boeke, A.J., Dekker, J.H., and Peerbooms, P.G., A comparison of yield from cervix versus vagina for culturing Candida albicans and Trichomonas vaginalis. Genitourin. Med. 69, 41, 1993.
32. Draper, D. et al., Detection of Trichomonas vaginalis in pregnant women with the InPouch TV culture system. J. Clin. Microbiol. 31, 1016, 1993.
33. Borchardt, K.A. et al., A comparison of the sensitivity of the InPouch TV, Diamond’s and Trichosel media for detection of Trichomonas vaginalis. Genitourin. Med. 73, 297, 1997.
34. Levi, M.H. et al., Comparison of the InPouch TV culture system and Diamond’s modi�ed medium for detection of Trichomonas vaginalis. J. Clin. Microbiol. 35, 3308, 1997.
35. Ohlemeyer, C.L. et al., Diagnosis of Trichomonas vaginalis in adolescent females: InPouch TV culture versus wet-mount microscopy. J. Adolesc. Health 22, 205, 1998.
36. Gilbert, R. et al., Cytopathogenic effect of Trichomonas vaginalis on human vaginal epithelial cells cultured in vitro. Infect. Immun. 68, 4200, 2000. 37. Thomason, J.L. et al., Comparison of four methods to detect Trichomonas vaginalis. J. Clin. Microbiol. 26, 1869, 1988. 38. Rompalo, A.M. et al., Evaluation of use of a single intravaginal swab to detect multiple sexually transmitted infections in active-duty military women. Clin. Infect. Dis. 33, 1455, 2001. 39. Beal, C. et al., The plastic envelope method, a simpli�ed technique for culture diagnosis of trichomoniasis. J. Clin. Microbiol. 30, 2265, 1992. 40. Kurth, A. et al., Performance of a new, rapid assay for detection of Trichomonas vaginalis. J. Clin. Microbiol. 42, 2940, 2004. 41. Mason, P.R., Serodiagnosis of Trichomonas vaginalis infection by the indirect �uorescent antibody test. J. Clin. Pathol. 32, 1211, 1979. 42. Alderete, J.F., Enzyme linked immunosorbent assay for detecting antibody to Trichomonas vaginalis: Use of whole cells and aqueous extract as antigen. Br. J. Vener. Dis. 60, 164, 1984. 43. Smith, R.F., Detection of Trichomonas vaginalis in vaginal specimens by direct immuno�uorescence assay. J. Clin. Microbiol. 24, 1107, 1986. 44. Watt, R.M. et al., Rapid assay for immunological detection of Trichomonas vaginalis. J. Clin. Microbiol. 24, 551, 1986. 45. Yule, A. et al., Detection of Trichomonas vaginalis antigen in women by enzyme immunoassay. J. Clin. Pathol. 40, 566, 1987. 46. Carney, J.A. et al., New rapid latex agglutination test for diagnosing Trichomonas vaginalis infection. J. Clin. Pathol. 41, 806, 1988. 47. Lisi, P.J. et al., Monoclonal-antibody-based enzyme-linked immunosorbent assay for Trichomonas vaginalis. J. Clin. Microbiol. 26, 1684, 1988. 48. Bickley, L.S. et al., Comparison of direct �uorescent antibody, acridine orange, wet mount, and culture for detection of Trichomonas vaginalis in women attending a public sexually transmitted diseases clinic. Sex. Transm. Dis. 16, 127, 1989. 49. Gombosová, A., and Valent, M., Dot-immunobinding assay with monoclonal antibody for detection of Trichomonas vaginalis in clinical specimens. Genitourin. Med. 66, 447, 1990. 50. Romia, S.A., and Othman, T.A., Detection of antitrichomonal antibodies in sera and cervical secretions in trichomoniasis. J. Egypt Soc. Parasitol. 21, 373, 1991. 51. Sharma, P. et al., A comparison of wet mount, culture and enzyme linked immunosorbent assay for the diagnosis of trichomoniasis in women. Trop. Geogr. Med. 43, 257, 1991. 52. DeMeo, L.R. et al., Evaluation of a deoxyribonucleic acid probe for the detection of Trichomonas vaginalis in vaginal secretions. Am. J. Obstet. Gynecol. 174, 1339, 1996. 53. Heine, R.P. et al., Polymerase chain reaction analysis of distal vaginal specimens: A less invasive strategy for detection of Trichomonas vaginalis. Clin. Infect. Dis. 24, 985, 1997. 54. Lin, P.R., Shaio, M.F., and Liu, J.Y., One-tube, nested-PCR assay for the detection of Trichomonas vaginalis in vaginal discharges. Ann. Trop. Med. Parasitol. 91, 61, 1997. 55. Shaio, M.F., Lin, P.R., and Liu, J.Y., Colorimetric one-tube nested PCR for detection of Trichomonas vaginalis in vaginal discharge. J. Clin. Microbiol. 35, 132, 1997. 56. Ryu, J. et al., Diagnosis of trichomoniasis by polymerase chain reaction. Yonsei Med. J. 40, 56, 1999. 57. van der Schee, C. et al., Improved diagnosis of Trichomonas vaginalis infection by PCR using vaginal swabs and urine specimens compared to diagnosis by wet mount microscopy, culture, and �uorescent staining. J. Clin. Microbiol. 37, 4127, 1999.
58. Lawing, L., Hedges, S., and Schwebke, J., Detection of trichomonosis in vaginal and urine specimens from women by culture and PCR. J. Clin. Microbiol. 38, 3585, 2000.
59. Patel, S.R. et al., Systematic review of diagnostic tests for vaginal trichomoniasis. Infect. Dis. Obstet. Gynecol. 8, 248, 2000.
60. Knox, J. et al., Evaluation of self-collected samples in contrast to practitioner- collected samples for detection of Chlamydia trachomatis, Neisseria gonorrhoeae, and Trichomonas vaginalis by polymerase chain reaction among women living in remote areas. Sex. Transm. Dis. 29, 647, 2002.
61. Schwebke, J., and Lawing, L., Improved detection by DNA ampli�cation of Trichomonas vaginalis in males. J. Clin. Microbiol. 40, 3681, 2002.
62. Wendel, K.A. et al., Trichomonas vaginalis polymerase chain reaction compared with standard diagnostic and therapeutic protocols for detection and treatment of vaginal trichomoniasis. Clin. Infect. Dis. 35, 576, 2002.
63. Kaydos-Daniels, S. et al., Validation of a urine-based PCRenzyme-linked immunosorbent assay for use in clinical reseach settings to detect Trichomonas vaginalis in men. J. Clin. Microbiol. 41, 318, 2003.
64. Pillay, A. et al., Comparison of a TaqMan-based real-time polymerase chain reaction with conventional tests for the detection of Trichomonas vaginalis. Sex. Transm. Infect. 83, 126, 2007.
65. Rojas, L., Fraga, J., and Sariego, I., Genetic variability between Trichomonas vaginalis isolates and correlation with clinical presentation. Infect. Genet. Evol. 4, 53, 2004.
66. Smith, K.S. et al., Comparison of conventional testing to polymerase chain reaction in detection of Trichomonas vaginalis in indigenous women living in remote areas. Int. J. STD AIDS 16, 811, 2005.
67. Hardick, J. et al., Comparison between the Gen-Probe transcription-mediated ampli�cation Trichomonas vaginalis research assay and real-time PCR for Trichomonas vaginalis detection using a Roche LightCycler instrument with female self-obtained vaginal swab samples and male urine samples. J. Clin. Microbiol. 44, 4197, 2006.
68. Radonjic, I.V. et al., Diagnosis of Trichomonas vaginalis infection: The sensitivities and speci�cities of microscopy, culture and PCR assay. Eur. J. Obst. Gynecol. Reprod. Biol. 126, 116, 2006.
69. Upcroft, J.A. et al., Genotyping Trichomonas vaginalis. Int. J. Parasitol. 36, 821, 2006.
70. Munson, E. et al., Trichomonas vaginalis transcriptionmediated ampli�cation-based analyte-speci�c reagent and alternative target testing of primary clinical vaginal saline suspensions. Diagn. Microbiol. Infect. Dis. 68, 66, 2010.
71. Nye, M.B., Schwebke, J.R., and Body, B.A., Comparison of APTIMA Trichomonas vaginalis transcription-mediated ampli�cation to wet mount microscopy, culture, and polymerase chain reaction for diagnosis of trichomoniasis in men and women. Am. J. Obst. Gynecol. 200, 188.e1, 2009.
72. Riley, D.E. et al., Development of a polymerase chain reaction based diagnosis of Trichomonas vaginalis. J. Clin. Microbiol. 30, 465, 1992.
73. Jordan, J.A., Lowery, D., and Trucco, M., Taqman-based detection of Trichomonas vaginalis DNA from female genital specimens. J. Clin. Microbiol. 39, 3819, 2001. 74. Katiyar, S.K., and Edlind, T.D., β-Tubulin genes of Trichomonas vaginalis. Mol. Biochem. Parasitol. 64, 33, 1994. 75. Madico, G. et al., Diagnosis of Trichomonas vaginalis infection by PCR using vaginal swab samples. J. Clin. Microbiol. 36, 3205, 1998. 76. Paces, J., Urbankova, V., and Urbanek, P., Cloning and characterization of a repetitive DNA sequence speci�c for Trichomonas vaginalis. Mol. Biochem. Parasitol. 54, 247, 1992. 77. Kengne, P. et al., Trichomonas vaginalis repeated DNA target for highly sensitive and speci�c polymerase chain reaction diagnosis. Cell Mol. Biol. 40, 819, 1994. 78. Mayta, H. et al., 18S ribosomal DNA-based PCR for diagnosis of Trichomonas vaginalis. J. Clin. Microbiol. 38, 2683, 2000. 79. Alderete, J.F. et al., Cloning and molecular characterization involved in Trichomonas vaginalis cytoadherence. Mol. Microbiol. 17, 69, 1995. 80. Jeremias, J. et al., Detection of Trichomonas vaginalis using the polymerase chain reaction in pregnant and non-pregnant women. Infect. Dis. Obstet. Gynecol. 2, 16, 1994. 81. Crucitti, T. et al., Comparison of culture and different PCR assays for detection of Trichomonas vaginalis in self collected vaginal swab specimens. Sex. Transm. Infect. 79, 393, 2003. 82. Caliendo, A.M. et al., Real-time PCR improves detection of Trichomonas vaginalis infection compared with culture using self-collected vaginal swabs. Infect. Dis. Obstet. Gynecol. 13, 145, 2005. 83. Schirm, J. et al., Trichomonas vaginalis detection using realtime TaqMan PCR. J. Microbiol. Methods 68, 243, 2007. 84. Depuydt, C.E. et al., Epidemiology of Trichomonas vaginalis and human papilloma virus infection detected by real-time PCR in �anders. Gynecol. Obstet. Invest. 70, 273, 2010. 85. Diaz, N. et al., Rapid detection of coinfections by Trichomonas vaginalis, Mycoplasma hominis, and Ureaplasma urealyticum by a new multiplex polymerase chain reaction. Diagn. Microbiol. Infect. Dis. 67, 30, 2010. 86. López-Villaseñor, I. et al., Trichomonas vaginalis ribosomal DNA: Analysis of the intergenic region and mapping of the transcription start point. Mol. Biochem. Parasitol. 137, 175, 2004. 87. Vatanshenassan, M. et al. Trichomonas vaginalis: Investigation of a novel diagnostic method in urine samples using cysteine proteinase 4 gene and PCR technique. Exp. Parasitol. 126, 187, 2010. 88. Simpson, P. et al., Real-time PCRs for detection of Trichomonas vaginalis beta-tubulin and 18S rRNA genes in female genital specimens. J. Med. Microbiol. 56, 772, 2007. 89. Hardick, J. et al., Use of the Roche LightCycler Instrument in a real-time PCR for Trichomonas vaginalis in urine samples from females and males. J. Clin. Microbiol. 41, 5619, 2003. 90. Simões-Barbosa, A. et al., Trichomonas vaginalis: Intrastrain polymorphisms within the ribosomal intergenic spacer do not correlate with clinical presentation. Exp. Parasitol. 110, 108, 2005. 12 Chapter 12 - Trypanosoma
9. Deborggraeve, S. et al. Molecular analysis of archived blood slides reveals an atypical human Trypanosoma infection. Diagn. Microbiol. Infect. Dis. 61, 428–433, 2008.
10. Joshi, P.P. et al. Human trypanosomiasis caused by Trypanosoma evansi in India: The �rst case report. Am. J. Trop. Med. Hyg. 73, 491–495, 2005.
11. Sarataphan, N. et al. Diagnosis of a Trypanosoma lewisilike (Herpetosoma) infection in a sick infant from Thailand. J. Med. Microbiol. 56, 1118–1121, 2007.
12. Kennedy, P.G. Diagnostic and neuropathogenesis issues in human African trypanosomiasis. Int. J. Parasitol. 36, 505– 512, 2006.
13. Yun, O. et al. Feasibility, drug safety, and effectiveness of etiological treatment programs for Chagas disease in Honduras, Guatemala, and Bolivia: 10-year experience of medecins sans frontieres. PLoS Negl. Trop. Dis. 3, e488, 2009.
14. Prata, A. Clinical and epidemiological aspects of Chagas disease. Lancet Infect. Dis. 1, 92–100, 2001.
15. Marin-Neto, J.A. et al. Pathogenesis of chronic Chagas heart disease. Circulation 115, 1109–1123, 2007.
16. Chappuis, F. et al. Options for �eld diagnosis of human African trypanosomiasis. Clin. Microbiol. Rev. 18, 133–146, 2005.
17. Buscher, P. et al. Improved models of mini anion exchange centrifugation technique (mAECT) and modi�ed single centrifugation (MSC) for sleeping sickness diagnosis and staging. PLoS. Negl. Trop. Dis. 3, e471, 2009.
18. Woo, P.T. The haematocrit centrifuge technique for the diagnosis of African trypanosomiasis. Acta Trop. 27, 384–386, 1970.
19. Woo, P.T. Evaluation of the haematocrit centrifuge and other techniques for the �eld diagnosis of human trypanosomiasis and �lariasis. Acta Trop. 28, 298–303, 1971.
20. Zillmann, U. et al. Improved performance of the anionexchange centrifugation technique for studies with human infective African trypanosomes. Acta Trop. 62, 183–187, 1996.
21. Magnus, E. et al. A card-agglutination test with stained trypanosomes (C.A.T.T.) for the serological diagnosis of T. b. gambiense trypanosomiasis. Ann. Soc. Belg. Med. Trop. 58, 169–176, 1978.
22. Truc, P. et al. Evaluation of the micro-CATT, CATT/ Trypanosoma brucei gambiense, and LATEX/T b gambiense methods for serodiagnosis and surveillance of human African trypanosomiasis in West and Central Africa. Bull. World Health Organ 80, 882–886, 2002.
23. Greenwood, B.M. et al. Cerebrospinal-�uid IgM in patients with sleeping-sickness. Lancet, 2, 525–527, 1973.
24. Lejon, V. et al. Neuro-in�ammatory risk factors for treatment failure in “early second stage” sleeping sickness patients treated with pentamidine. J. Neuroimmunol. 144, 132–138, 2003.
25. Lejon, V. et al. Novel markers for treatment outcome in late-stage Trypanosoma brucei gambiense trypanosomiasis. Clin. Infect. Dis. 47, 15–22, 2008.
26. Chippaux, J.P. et al. Sensitivity and speci�city of Chagas StatPak test in Bolivia. Trop. Med. Int. Health, 14, 732–735, 2009.
27. Ferreira, A.W. et al. Laboratory diagnosis of Chagas’ heart disease. Sao Paulo Med. J. 113, 767–771, 1995.
28. Mullis, K.B. et al. Speci�c synthesis of DNA in vitro via a polymerase-catalyzed chain reaction. Methods Enzymol. 155, 335–350, 1987.
29. Gibson, W. Resolution of the species problem in African trypanosomes. Int. J. Parasitol. 37, 829–838, 2007.
30. Xong, H.V. et al. A VSG expression site-associated gene confers resistance to human serum in Trypanosoma rhodesiense. Cell, 95, 839–846, 1998. 31. Welburn, S.C. et al. Identi�cation of human-infective trypanosomes in animal reservoir of sleeping sickness in Uganda by means of serum-resistance-associated (SRA) gene. Lancet 358, 2017–2019, 2001. 32. Radwanska, M. et al. Novel primer sequences for polymerase chain reaction-based detection of Trypanosoma brucei gambiense. Am. J. Trop. Med. Hyg. 67, 289–295, 2002. 33. Picozzi, K. et al. A multiplex PCR that discriminates between Trypanosoma brucei brucei and zoonotic T. b. rhodesiense, Exp. Parasitol. 118, 41–46, 2008. 34. Mugasa, C.M. et al. Diagnostic accuracy of molecular ampli�cation tests for human African trypanosomiasis— Systematic review. PLoS. Negl. Trop. Dis. 6, e1438, 2012. 35. Vallejo, G.A. et al. Species speci�c detection of Trypanosoma cruzi and Trypanosoma rangeli in vector and mammalian hosts by polymerase chain reaction ampli�cation of kinetoplast minicircle DNA. Acta Trop. 72, 203–212, 1999. 36. Velazquez, M. et al. Trypanosoma cruzi: An analysis of the minicircle hypervariable regions diversity and its in�uence on strain typing. Exp. Parasitol. 120, 235–241, 2008. 37. Gil, J. et al. Comparison of a PCR test based on the histone H2A/SIRE genes with classical serological tests for the diagnosis of chronic Chagas disease in Colombian patients. Biomedica. 27(Suppl 1), 83–91, 2007. 38. Desquesnes, M. et al. Detection and identi�cation of Trypanosoma of African livestock through a single PCR based on internal transcribed spacer 1 of rDNA. Int. J. Parasitol. 31, 610–614, 2001. 39. Cox, A. et al. A PCR based assay for detection and differentiation of African trypanosome species in blood. Exp. Parasitol. 111, 24–29, 2005. 40. Adams, E.R. et al. Trypanosome identi�cation in wild tsetse populations in Tanzania using generic primers to amplify the ribosomal RNA ITS-1 region. Acta Trop. 100, 103–109, 2006. 41. Schijman, A.G. et al. Differential detection of Blastocrithidia triatomae and Trypanosoma cruzi by ampli�cation of 24 s alpha ribosomal RNA genes in faeces of sylvatic triatomine species from rural northwestern Argentina. Acta Trop. 99, 50–54, 2006. 42. Hamilton, P.B. et al. A novel, high-throughput technique for species identi�cation reveals a new species of tsetse-transmitted trypanosome related to the Trypanosoma brucei subgenus. Trypanozoon. Infect. Genet. Evol. 8, 26–33, 2008. 43. Deborggraeve, S. et al. Molecular dipstick test for diagnosis of sleeping sickness. J. Clin. Microbiol. 44, 2884–2889, 2006. 44. Deborggraeve, S. et al. T. cruzi OligoC-TesT: a Simpli�ed and standardized polymerase chain reaction format for diagnosis of Chagas disease. PLoS. Negl. Trop. Dis. 3, e450, 2009. 45. Stothard, J.R. et al. On the molecular taxonomy of Trypanosoma cruzi using riboprinting. Parasitology 117(Pt 3), 243–247, 1998. 46. Bastrenta, B. et al. Restriction fragment length polymorphism of 195 bp repeated satellite DNA of Trypanosoma cruzi supports the existence of two phylogenetic groups. Mem. Inst. Oswaldo Cruz 94, 323–328, 1999. 47. Luna-Marin, K.P. et al. ITS-RFLP- and RAPD-based genetic variability of Trypanosoma cruzi I, human and vector strains in Santander, Colombia. Parasitol. Res. 105, 519–528, 2009. 48. Campos, R.F. et al. Comparative analysis by polymerase chain reaction ampli�ed minicircles of kinetoplast DNA of a stable strain of Trypanosoma cruzi from Sao Felipe, Bahia, its clones and subclones: Possibility of predominance of a principal clone in this area. Mem. Inst. Oswaldo Cruz 94, 23–29, 1999.
49. Delespaux, V. et al. PCR-RFLP using Ssu-rDNA ampli�cation: Applicability for the diagnosis of mixed infections with different trypanosome species in cattle. Vet. Parasitol. 117, 185–193, 2003.
50. Becker, S. et al. Real-time PCR for detection of Trypanosoma brucei in human blood samples. Diagn. Microbiol. Infect. Dis. 50, 193–199, 2004.
51. Cummings, K.L. et al. Rapid quantitation of Trypanosoma cruzi in host tissue by real-time PCR. Mol. Biochem. Parasitol. 129, 53–59, 2003.
52. Freitas, J.M. et al. Real time PCR strategy for the identi�cation of major lineages of Trypanosoma cruzi directly in chronically infected human tissues. Int. J. Parasitol. 35, 411–417, 2005.
53. Piron, M. et al. Development of a real-time PCR assay for Trypanosoma cruzi detection in blood samples. Acta Trop. 103, 195–200, 2007.
54. Duffy, T. et al. Accurate real-time PCR strategy for monitoring bloodstream parasitic loads in Chagas disease patients. PLoS. Negl. Trop. Dis. 3, e419, 2009.
55. Hamilton, P.B. et al. Phylogenetic analysis reveals the presence of the Trypanosoma cruzi clade in African terrestrial mammals. Infect. Genet. Evol. 9, 81–86, 2009.
56. Hamilton, P.B. et al. Trypanosomes are monophyletic: Evidence from genes for glyceraldehyde phosphate dehydrogenase and small subunit ribosomal RNA. Int. J. Parasitol. 34, 1393–1404, 2004.
57. Sajid, M. et al. Cysteine proteases of parasitic organisms. Mol. Biochem. Parasitol. 120, 1–21, 2002.
58. Cortez, A.P. et al. Cathepsin L-like genes of Trypanosoma vivax from Africa and South America—Characterization, relationships and diagnostic implications. Mol. Cell Probes. 23, 44–51, 2009.
59. Marcili, A. et al. Comparative phylogeography of Trypanosoma cruzi TCIIc: New hosts, association with terrestrial ecotopes, and spatial clustering. Infect. Genet. Evol. 9, 1265–1274, 2009.
60. Da Silva, F.M. et al. Phylogeny, taxonomy and grouping of Trypanosoma rangeli isolates from man, triatomines and sylvatic mammals from widespread geographical origin based on SSU and ITS ribosomal sequences. Parasitology 129, 549–561, 2004.
61. Metzker, M.L. Emerging technologies in DNA sequencing. Genome Res. 15, 1767–1776, 2005.
62. Deiman, B. et al. Characteristics and applications of nucleic acid sequence-based ampli�cation (NASBA). Mol. Biotechnol. 20, 163–179, 2002.
63. Mugasa, C.M. et al. Detection of Trypanosoma brucei parasites in blood samples using real-time nucleic acid sequence-based ampli�cation. Diagn. Microbiol. Infect. Dis. 61, 440–445, 2008. 64. Mugasa, C.M. et al. Nucleic acid sequence-based ampli�cation with oligochromatography for detection of Trypanosoma brucei in clinical samples. J. Clin. Microbiol. 47, 630–635, 2009. 65. Notomi, T. et al. Loop-mediated isothermal ampli�cation of DNA. Nucleic Acids Res. 28, E63, 2000. 66. Kuboki, N. et al. Loop-mediated isothermal ampli�cation for detection of African trypanosomes. J. Clin. Microbiol. 41, 5517–5524, 2003. 67. Thekisoe, O.M. et al. Species-speci�c loop-mediated isothermal ampli�cation (LAMP) for diagnosis of trypanosomosis. Acta Trop. 102, 182–189, 2007. 68. Njiru, Z.K. et al. Loop-mediated isothermal ampli�cation (LAMP) method for rapid detection of Trypanosoma brucei rhodesiense. PLoS. Negl. Trop. Dis. 2, e147, 2008. 69. Njiru, Z.K. et al. African trypanosomiasis: Sensitive and rapid detection of the sub-genus Trypanozoon by loop-mediated isothermal ampli�cation (LAMP) of parasite DNA. Int. J. Parasitol. 38, 589–599, 2008. 70. Thekisoe, O.M. et al. Stability of loop-mediated isothermal ampli�cation (LAMP) reagents and its ampli�cation ef�ciency on crude trypanosome DNA templates. J. Vet. Med. Sci. 71, 471–475, 2009. 71. Gibson, W. Species-speci�c probes for the identi�cation of the African tsetse-transmitted trypanosomes. Parasitology 136, 1501–1507, 2009. 72. Harris, E. et al. Detection of Trypanosoma brucei spp. in human blood by a nonradioactive branched DNA-based technique. J. Clin. Microbiol. 34, 2401–2407, 1996. 73. Radwanska, M. et al. Direct detection and identi�cation of African trypanosomes by �uorescence in situ hybridization with peptide nucleic acid probes. J. Clin. Microbiol. 40, 4295–4297, 2002. 74. Lorger, M. et al. Targeting the variable surface of African trypanosomes with variant surface glycoproteinspeci�c, serum-stable RNA aptamers. Eukaryot. Cell 2, 84–94, 2003. 75. Ulrich, H. et al. In vitro selection of RNA aptamers that bind to cell adhesion receptors of Trypanosoma cruzi and inhibit cell invasion. J. Biol. Chem. 277, 20756–20762, 2002. 76. Deborggraeve, S. et al. Molecular diagnostics for sleeping sickness: What is the bene�t for the patient? Lancet Infect. Dis. 10, 433–439, 2010. 77. Boom, R. et al. Rapid and simple method for puri�cation of nucleic acids. J. Clin. Microbiol. 28, 495–503, 1990. 78. Kaneko, H. et al. Tolerance of loop-mediated isothermal ampli�cation to a culture medium and biological substances. J. Biochem. Biophys. Methods 70, 499–501, 2007.
Section II
Ciliophora and Apicomplex 13 Chapter 13 - Babesia
1. Homer, M.J. et al., Babesiosis. Clin. Microbiol. Rev., 13, 451, 2000.
2. Vannier, E., Gewurz, B.E., and Krause, P.J., Human Babesiosis. Infect. Dis. Clin. North Am., 22, 469, 2008.
3. Vannier, E., and Krause, P.J., Update on Babesiosis. Interdiscip Perspect Infect Dis., 2009, 984568, 2009.
4. Babes, V., Sur l’hemoglobinurie bacterienne du boeuf. C. R. Acad. Sci., 107, 692, 1888.
5. M’Fadyean, J., and Stockman, S., A new species of piroplasm found in the blood of British cattle. J. Comp. Pathol., 24, 340, 1911.
6. Skrabalo, Z., and Deanovic, Z., Piroplasmosis in man; report of a case. Doc. Med. Geograph. Trop., 9, 11, 1957.
7. Hunfeld, K.-P., Hildebrandt, A., and Gray, J.S., Babesiosis: Recent insights into an ancient disease. Int. J. Parasitol., 38, 1219, 2008.
8. Allsopp, M.T., and Allsopp, B.A., Molecular sequence evidence for the reclassi�cation of some Babesia species. Ann. N. Y. Acad. Sci., 1081, 509, 2006.
9. Kogut, S.J. et al., Babesia microti, upstate New York. Emerg. Infect. Dis., 11, 476, 2005. 10. Shih, C.-M. et al., Human babesiosis in Taiwan: Asymptomatic infection with a Babesia microti-like organism in a Taiwanese woman. J. Clin. Microbiol., 35, 450, 1997. 11. Wei, Q. et al., Human babesiosis in Japan: Isolation of Babesia microti-like parasites from an asymptomatic transfusion donor and from a rodent from an area where babesiosis is endemic. J. Clin. Microbiol., 39, 2178, 2001. 12. Rios, L., Alvarez, G., and Blair, S., Serological and parasitological study and report of the �rst case of human babesiosis in Colombia. Rev. Soc. Bras. Med. Trop., 36, 493, 2003. 13. Marathe, A. et al., Human babesiosis—a case report. Indian J. Med. Microbiol., 23, 267, 2005. 14. Hildebrandt, A. et al., First con�rmed autochthonous case of human Babesia microti infection in Europe. Eur. J. Clin. Microbiol. Infect. Dis., 26, 595, 2007. 15. Zintl, A. et al., Babesia divergens, a bovine blood parasite of veterinary and zoonotic importance. Clin. Microbiol. Rev., 16, 622, 2003. 16. Herwaldt, B.L. et al., A fatal case of babesiosis in Missouri: Identi�cation of another piroplasm that infects humans. Ann. Intern. Med., 124, 643, 1996. 17. Herwaldt, B.L. et al., Babesia divergens-like Infection, Washington State. Emerg. Infect. Dis., 10, 622, 2004. 18. Conrad, P.A. et al., Description of Babesia duncani n.sp. (Apicomplexa: Babesiidae) from humans and its differentiation from other piroplasms. Int. J. Parasitol., 36, 779, 2006. 19. Herwaldt, B.L. et al., Endemic babesiosis in another eastern state: New Jersey. Emerg. Infect. Dis., 9, 184, 2003. 20. Haselbarth, K. et al., First case of human babesiosis in Germany—clinical presentation and molecular characterisation of the pathogen. Int. J. Med. Microbiol., 297, 197, 2007. 21. Krause, P.J. et al., Persistent and relapsing babesiosis in immunocompromised patients. Clin. Infect. Dis., 46, 370, 2008. 22. Herwaldt, B.L. et al., Molecular characterization of a nonBabesia divergens organism causing zoonotic babesiosis in Europe. Emerg. Infect. Dis., 9, 942, 2003. 23. Armstrong, P.M. et al., Diversity of Babesia infecting deer ticks (Ixodes dammini). Am. J. Trop. Med. Hyg., 58, 739, 1998. 24. Kjemtrup, A.M., and Conrad, P.A., Human babesiosis: An emerging tick-borne disease. Int. J. Parasitol., 30, 1323, 2000. 25. Holman, P.J., Phylogenetic and biologic evidence that Babesia divergens is not endemic in the United States. Ann. New York Acad. Sci., 1081, 518, 2006. 26. Blaschitz, M. et al., Babesia species occurring in Austrian Ixodes ricinus ticks. Appl. Env. Microbiol., 74, 4841, 2008. 27. Rodgers, S.E., and Mather, T.N., Human Babesia microti incidence and Ixodes scapularis distribution, Rhode Island, 1998–2004. Emerg. Infect. Dis., 13, 633, 2007. 28. Cieniuch, S., Stan´czak, J., and Ruczaj, A., The �rst detection of Babesia EU1 and Babesia canis canis in Ixodes ricinus ticks (Acari, Ixodidae) collected in urban and rural areas in northern Poland. Pol. J. Microbiol., 58, 231, 2009. 29. Reye, A.L. et al., Prevalence and seasonality of tick-borne pathogens in questing Ixodes ricinus ticks from Luxembourg. Appl. Env. Microbiol., 76, 2923, 2010. 30. Eskow, E.S. et al., Southern extension of the range of human babesiosis in the Eastern United States. J. Clin. Microbiol., 37, 2051, 1999. 31. Gubernot, D.M. et al., Babesia infection through blood transfusions: Reports received by the US Food and Drug Administration, 1997–2007. Clin. Infect. Dis., 48, 25, 2009. 32. Gray, J. et al., Transmission studies of Babesia microti in Ixodes ricinus ticks and gerbils. J. Clin. Microbiol., 40, 1259, 2002.
33. Ortiz, J.M., and Eagle, R.C., Jr., Ocular �ndings in human babesiosis (Nantucket fever). Am. J. Ophthalmol., 93, 307, 1982.
34. Foppa, I.M. et al., Entomologic and serologic evidence of zoonotic transmission of Babesia microti, eastern Switzerland. Emerg. Infect. Dis., 8, 722, 2002.
35. Okabayashi, T. et al., Detection of Babesia microti-like parasite in �lter paper-absorbed blood of wild rodents. J. Vet. Med. Sci., 64, 145, 2002.
36. Kim, J.-Y. et al., First case of human babesiosis in Korea: Detection and characterization of a novel type of Babesia sp. (KO1) similar to ovine Babesia. J. Clin. Microbiol., 45, 2084, 2007.
37. Benach, J.L., and Habicht, G.S., Clinical characteristics of human babesiosis. J. Infect. Dis., 144, 481, 1981.
38. Bush, J.B. et al., Human babesiosis—a preliminary report of 2 suspected cases in South Africa. South Afr. Med. J., 78, 699, 1990.
39. Goren�ot, A. et al., Human babesiosis. Ann. Trop. Med. Parasitol., 92, 489, 1998.
40. Beattie, J.F., Michelson, M.L., and Holman, P.J., Acute babesiosis caused by Babesia divergens in a resident of Kentucky. N. Engl. J. Med., 347, 697, 2002.
41. Fox, L.M. et al., Neonatal babesiosis: Case report and review of the literature. Pediatr. Infect. Dis. J., 25, 169, 2006.
42. Krause, P.J. et al., Diagnosis of babesiosis: Evaluation of a serologic test for the detection of Babesia microti antibody. J. Infect. Dis., 169, 923, 1994.
43. Berman, K.H. et al., Fatal case of babesiosis in postliver transplant patient. Transplantation, 87, 452, 2009.
44. Froberg, M.K. et al., Case report: Spontaneous splenic rupture during acute parasitemia of Babesia microti. Ann. Clin. Lab. Sci., 38, 390, 2008.
45. Hemmer, R.M., Ferrick, D.A., and Conrad, P.A., Role of T cells and cytokines in fatal and resolving experimental babesiosis: Protection in TNFRp55 − /− mice infected with the human Babesia WA1 parasite. J. Parasitol., 86, 736, 2000.
46. Krause, P.J. et al., Comparison of PCR with blood smear and inoculation of small animals for diagnosis of Babesia microti parasitemia. J. Clin. Microbiol., 34, 2791, 1996.
47. Healy, G.R., and Ruebush, T.K., II Morphology of Babesia microti in human blood smears. Am. J. Clin. Pathol., 73, 107, 1980.
48. Sun, T., Tenenbaum, M.J., and Greenspan, J., Morphologic and clinical observations in human infection with Babesia microti. J. Infect. Dis., 148, 239, 1983.
49. Ryan, R. et al., Diagnosis of babesiosis using an immunoblot serologic test. Clin. Diagn. Lab. Immunol., 8, 1177, 2001.
50. Gubbels, J.M. et al., Simultaneous detection of bovine Theileria and Babesia species using reverse line blot hybridization. J. Clin. Microbiol., 37, 1782, 1999. 51. Skotarczak, B., and Cichocka, A., Isolation and ampli�cation by polymerase chain reaction DNA of Babesia microti and Babesia divergens in ticks in Poland. Ann. Agric. Environ. Med., 8, 187, 2001. 52. Pieniazek, N., Sawczuk, M., and Skotarczak, B., Molecular identi�cation of Babesia parasites isolated from Ixodes ricinus ticks collected in northwestern Poland. J. Parasitol., 92, 32, 2006. 53. Holman, P.J. et al., Ribosomal RNA analysis of Babesia odocoilei isolates from farmed reindeer (Rangifer tarandus tarandus) and elk (Cervus elaphus canadensis) in Wisconsin. Parasitol. Res., 91, 378, 2003. 54. Persing, D.H. et al., Detection of Babesia microti by polymerase chain reaction. J. Clin. Microbiol., 30, 2097, 1992. 56. Centeno-Lima, S. et al., A fatal case of human babesiosis in Portugal: Molecular and phylogenetic analysis. Trop. Med. Int. Health, 8, 760, 2003. 57. Caccio, S. et al., The beta-tubulin gene of Babesia and Theileria parasites is an informative marker for species discrimination. Int. J. Parasitol., 30, 1181, 2000. 58. Zamoto, A. et al., Epizootiologic survey for Babesia microti among small wild mammals in northeastern Eurasia and a geographic diversity in the beta-tubulin gene sequences. J. Vet. Med. Sci., 66, 785, 2004. 59. Casati, S. et al., Presence of potentially pathogenic Babesia sp. for human in Ixodes ricinus in Switzerland. Ann. Agric. Environ. Med., 13, 65, 2006. 60. Denes, E. et al., Management of Babesia divergens babesiosis without a complete course of quinine treatment. Eur. J. Clin. Microbiol. Infect. Dis., 18, 672, 1999. 61. Krause, P.J. et al., Atovaquone and azithromycin for the treatment of babesiosis. N. Engl. J. Med., 343, 1454, 2000. 62. Raju, M. et al., Atovaquone and azithromycin treatment for babesiosis in an infant. Pediatr. Infect. Dis. J., 26, 181, 2007. 63. Weiss, L.M., Wittner, M., and Tanowitz, H.B., The treatment of babesiosis. N. Engl. J. Med., 344, 773, 2001. 64. Vyas, J.M., Telford, S.R., and Robbins, G.K., Treatment of refractory Babesia microti infection with atovaquone-proguanil in an HIV-infected patient: Case report. Clin. Infect. Dis., 45, 1588, 2007. 65. Holman, P.J. et al., Comparative infectivity of Babesia divergens and a zoonotic Babesia divergens-like parasite in cattle. Am. J. Trop. Med. Hyg., 73, 865, 2005. 66. Saito-Ito, A. et al., Transfusion-acquired, autochthonous human babesiosis in Japan: Isolation of Babesia microti-like parasites with hu-RBC-SCID mice. J. Clin. Microbiol., 38, 4511, 2000. 14 Chapter 14 - Balantidium
1. Adl, S.M. et al., The new higher level classi�cation of eukaryotes with emphasis on the taxonomy of protists. J. Eukaryot. Microbiol., 52, 399, 2005.
2. Grim, J.N., and Buonanno, F., A re-description of the ciliate genus and type species, Balantidium entozoon. Eur. J. Protistol., 45, 174, 2009.
3. Grim, J.N., Description of somatic kineties and vestibular organization of Balantidium jocularum, sp. n., possible taxonomic implications for the class Litostomatea and the genus Balantidium. Acta Protozool., 32, 37, 1993.
4. Grim, J.N., Clements, K.D., and By�eld, T., New species of Balantidium and Paracichlidotherus (Ciliophora) inhabiting the intestines of four surgeon�sh species from the Tuvalu Islands, Paci�c Ocean. J. Eukaryot. Microbiol., 49, 146, 2002.
5. Li, M. et al., First report of two Balantidium species from the Chinese giant salamander, Andrias davidianus: Balantidiuim sinensis Nie 1935 and Balantidium andianusis n. sp. Parasitol. Res., 102, 605, 2008.
6. Schuster, F.L., and Ramirez-Avila, L., Current world status of Balantidium coli. Clin. Microbiol. Rev., 21, 626, 2008.
7. Skotarczak, B., Ultrastructural and cytochemical identi�cation of peroxisomes in Balantidium coli, Ciliophora. Folia Biol., 45, 117, 1997.
8. Hindsbo, O. et al., Age-dependent occurrence of the intestinal ciliate Balantidium coli in pigs at a Danish research farm. Acta Vet. Scand., 41, 79, 2000.
9. Yang, Y. et al., Diarrhoea in piglets and monkeys experimentally infected with Balantidium coli isolated from human faeces. J. Trop. Med. Hyg., 98, 69, 1995.
10. Nakauchi, K., The prevalence of Balantidium coli infection in �fty-six mammalian species. J. Vet. Med. Sci., 61, 63, 1999.
11. Cox, F.E., Human balantidiasis in Iran: Are camels reservoir hosts? Trends Parasitol., 21, 553, 2005.
12. Headley, S.A., Kummala, E., and Sukura, A., Balantidium coli-infection in a Finnish horse. Vet. Parasitol., 158, 129, 2008.
13. Bilal, C.Q. et al., Prevalence and chemotherapy of Balantidium coli in cattle in the River Ravi region, Lahore (Pakistan). Vet. Parasitol., 163, 15, 2009.
14. Pomajbíková, K. et al., Discrepancies in the occurrence of Balantidium coli between wild and captive African great apes. J. Parasitol., 96, 1139, 2010.
15. Karanis, P., Kourenti, C., and Smith, H., Waterborne transmission of protozoan parasites: A worldwide review of outbreaks and lessons learnt. J. Water Health, 5, 1, 2007.
16. Esteban, J.-G. et al., Balantidiasis in Aymara children from the northern Bolivian altiplano. Am. J. Trop. Med. Hyg., 59, 922, 1998.
17. Maleky, F., Case report of Balantidium coli in human from south of Tehran, Iran. Indian J. Med. Sci., 52, 201, 1998.
18. Baskerville, L., Ahmed, Y., and Ramchand, S., Balantidium colitis. Report of a case. Am. J. Dig. Dis., 15, 727, 1970.
19. Francesco, R., and Giannotti, T., Balantidium coli in cervicovaginal cytology. A case report. Pathologica, 75, 439, 1983.
20. Currie, A.R., Human balantidiasis. A case report. S. Afr. J. Surg., 28, 23, 1990.
21. Dodd, L.G., Balantidium coli infestation as a cause of acute appendicitis. J. Infect. Dis., 163, 1392, 1991.
22. Cermeño, J.R. et al., Balantidium coli in a HIV-infected patient with chronic diarrhea. AIDS, 17, 941, 2003. 23. Barsoum, R.S., Parasitic infections in organ transplantation. Exp. Clin. Transplant., 2, 258, 2004. 24. Agapov, M., Balantidiasis in a patient with suspected Crohn’s disease. Endoscopy, 38, 655, 2006. 25. Ladas, S.D. et al., Invasive balantidiasis presented as chronic colitis and lung involvement. Dig. Dis. Sci., 34, 1621, 1989. 26. Oberhuber, G., Karpitschka, G., and Stolte, M., Balantidium coli: A rare cause of colonic ulcer. Eur. J. Gastroenterol. Hepatol., 5, 755, 1993. 27. Gonzalez de Canales Simon, P. et al., Colonic balantidiasis. Gastroenterol. Hepatol., 23, 129, 2000. 28. Dorfman, S., Rangel, O., and Bravo, L.G., Balantidiasis: Report of a fatal case with appendicular and pulmonary involvement. Trans. R. Soc. Trop. Med. Hyg., 78, 833, 1984. 29. Clyti, E. et al., A case of coinfection by Balantidium coli and HIV in French Guiana. Bull. Soc. Pathol. Exot., 91, 309, 1998. 30. Vasilakopolou, A. et al., Balantidium coli pneumonia in an immunocompromised patient. Scand. J. Infect. Dis., 35, 144, 2003. 31. Ferry, T. et al., Severe peritonitis due to Balantidium coli acquired in France. Eur. J. Clin. Microbiol. Infect. Dis., 23, 393, 2004. 32. Sharma, S., and Harding, G., Necrotizing lung infection caused by the protozoan Balantidium coli. Can. J. Infect. Dis., 14, 163, 2003. 33. Anargyrou, K. et al., Pulmonary Balantidium coli infection in a leukemic patient. Am. J. Hematol., 73, 180, 2003. 34. Koopowitz, A. et al., Balantidium coli-induced pulmonary haemorrhage with iron de�ciency. S. Afr. Med. J., 100, 534, 2010. 35. Yazar, S. et al., Dysentery caused by Balantidium coli in a patient with non-Hodgkin’s lymphoma from Turkey. World J. Gastroenterol., 10, 458, 2004. 36. Maino, A. et al., Urinary balantidiasis: Diagnosis at a glance by urine sediment examination. J. Nephrol., 23, 732, 2010. 37. Clark, C.G., and Diamond, L.S., Methods for cultivation of luminal parasitic protists of clinical importance. Clin. Microbiol. Rev., 15, 329, 2002. 38. Lynn, D.H., and Sogin, M.L., Assessment of phylogenetic relationships among ciliated protists using partial ribosomal RNA sequences derived from reverse transcripts. Biosystems, 21, 249, 1988. 39. Lynn, D.H., Morphology or molecules: How do we identify the major lineages of ciliates (Phylum Ciliophora)? Eur. J. Protistol., 39, 356, 2003. 40. Strüder-Kypke, M.C. et al., Molecular phylogeny of litosome ciliates (Ciliophora, Litostomatea) with emphasis on freeliving haptorian genera. Protist, 157, 261, 2006. 41. Strüder-Kypke, M.C., Kornilova, O.A., and Lynn, D.H., Phylogeny of trichostome ciliates (Ciliophora, Litostomatea) endosymbiontic in the Yakut horse (Equus caballus). Eur. J. Protistol., 43, 319, 2007. 42. Gao, S. et al., Phylogeny of six genera of the subclass Haptoria (Ciliophora, Litostomatea) inferred from sequences of the gene coding for small subunit ribosomal RNA. J. Eukaryot. Microbiol., 55, 562, 2008. 43. Nilles-Bije, M.L., and Rivera, W.L., Ultrastructural and molecular characterization of Balantidium coli isolated in the Philippines. Parasitol. Res., 106, 387, 2010. 44. Medlin, L. et al., The characterization of enzymatically ampli�ed eukaryotic 16S-like rRNA-coding regions. Gene, 71, 491, 1988. 15 Chapter 15 - Cryptosporidium
1. Fayer, R. et al. Examination of naturally exposed bottlenose dolphins (Tursiops truncatus) for microsporidia, Cryptosporidium, and Giardia. J. Parasitol. 94, 143, 2008.
2. Rose, J.B. et al. Survey of potable water supplies for Cryptosporidium and Giardia. Environ. Sci. Technol. 25, 1393, 1991.
3. Fayer, R. et al. Epidemiology of Cryptosporidium: Trans mission, detection and identi�cation. Int. J. Parasitol. 30, 1305, 2000.
4. Karanis, P. et al. Waterborne transmission of protozoan parasites: A worldwide review of outbreaks and lessons learnt. J. Water Health 5, 1, 2007.
5. Levine, N.D. Taxonomy and review of the coccidian genus Cryptosporidium (protozoa, apicomplexa). J. Protozool. 31, 94, 1984.
6. Hijjawi, N.S. et al. Successful in vitro cultivation of Cryptosporidium andersoni: Evidence for the existence of novel extracellular stages in the life cycle and implications for the classi�cation of Cryptosporidium. Int. J. Parasitol. 32, 1719, 2002.
7. Carreno, R.A. et al. Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitol. Res. 85, 899, 1999.
8. Hijjawi, N.S. et al. Complete development of Cryptosporidium parvum in host cell-free culture. Int. J. Parasitol. 34, 769, 2004.
9. Leander, B.S. et al. Molecular phylogeny and surface morphology of Colpodella edax (Alveolata): Insights into the phagotrophic ancestry of apicomplexans. J. Eukaryot. Microbiol. 50, 334, 2003.
10. Rosales, M.J. et al. Extracellular like-gregarine stages of Cryptosporidium parvum. Acta Trop. 95, 74, 2005.
11. Templeton, T.J. et al. Genome sequence survey for Ascogregarina taiwanensis supports evolutionary af�liation, but metabolic diversity between a gregarine and Cryptosporidium. Mol. Biol. Evol. 27, 235, 2010.
12. Fayer, R. et al. Cryptosporidium ryanae n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). Vet. Parasitol. 156, 191, 2008.
13. Fayer, R. and Santin, M. Cryptosporidium xiaoi n. sp. (Apicomplexa: Cryptosporidiidae) in sheep (Ovis aries). Vet. Parasitol. 164, 192, 2009.
14. Fayer, R. Taxonomy and species delimitation in Cryptosporidium. Exp. Parasitol. 124, 90, 2010.
15. Xiao, L. Molecular epidemiology of cryptosporidiosis: An update. Exp. Parasitol. 124, 80, 2010.
16. Cama, V.A. et al. Cryptosporidium species and genotypes in HIV-positive patients in Lima, Peru. J. Eukaryot. Microbiol. 50, S531, 2003.
17. Xiao, L., and Fayer, R. Molecular characterisation of species and genotypes of Cryptosporidium and Giardia and assessment of zoonotic transmission. Int. J. Parasitol. 38, 1239, 2008.
18. Xiao, L. et al. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl. Environ. Microbiol. 70, 891, 2004.
19. Warren, C.A., and Guerrant, R.L. Clinical disease and pathology. In: Fayer, R., and Xiao, L. (eds), Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL, p. 235, 2008.
20. Xiao, L. and Feng, Y. Zoonotic cryptosporidiosis. FEMS Immunol. Med. Microbiol. 52, 309, 2008.
21. Morgan, U. et al. Molecular characterization of Cryptosporidium isolates obtained from human immunode�ciency virus-infected individuals living in Switzerland, Kenya, and the United States. J. Clin. Microbiol. 38, 1180, 2000. 22. Guyot, K. et al. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 39, 3472, 2001. 23. Gargala, G. Drug treatment and novel drug target against Cryptosporidium. Parasite 15, 275, 2008. 24. Farthing, M.J. Clinical aspects of human cryptosporidiosis, Contrib. Microbiol. Immunol. 6, 50, 2000. 25. Okhuysen, P.C. and Chappell, C.L. Cryptosporidium virulence determinants—Are we there yet? Int. J. Parasitol. 32, 517, 2002. 26. Liu, J. et al. Inhibition of apoptosis in Cryptosporidium parvum-infected intestinal epithelial cells is dependent on survivin. Infect. Immun. 76, 3784, 2008. 27. Wang, C. et al. Induced susceptibility of host is associated with an impaired antioxidant system following infection with Cryptosporidium parvum in Se-de�cient mice. PLoS One 4, 4628, 2009. 28. Carmolli, M. et al. De�cient serum mannose-binding lectin levels and MBL2 polymorphisms increase the risk of single and recurrent Cryptosporidium infections in young children. J. Infect. Dis. 200, 1540, 2009. 29. Fall, A. et al. Morphology is not a reliable tool for delineating species within Cryptosporidium. J. Parasitol. 89, 399, 2003. 30. O’Donoghue, P.J. Cryptosporidium and cryptosporidiosis in man and animals. Int. J. Parasitol. 25, 139, 1995. 31. Baxby, D. et al. The development and performance of a simple, sensitive method for the detection of Cryptosporidium oocysts in faeces. J. Hyg. (Lond), 93, 317, 1984. 32. Ma, P. and Soave, R. Three-step stool examination for cryptosporidiosis in 10 homosexual men with protracted watery diarrhoea. J. Infect. Dis. 147, 824, 1983. 33. Henricksen, S.A. and Pohlenz, J.F.L. Staining of cryptospordia by a modi�ed Ziehl-Neelsen technique. Acta Vet. Scand. 22, 594, 1981. 34. Pohjola, S. et al. Dimethylsulphoxide-Ziehl-Neelsen staining technique for the detection of cryptosporidial oocysts. Vet. Rec. 115, 442, 1984. 35. Smith, H.V. et al. An analysis of staining methods for the detection of Cryptosporidium spp. oocysts in water-related samples. Parasitology, 99, 323, 1989. 36. Moodley, D. et al. Cryptosporidium infections in children in Durban. Seasonal variation, age distribution and disease status. S. Afr. Med. J. 79, 295, 1991. 37. Chichino, G. et al. New rapid staining methods of Cryptosporidium oocysts in stools. J. Protozool. 38, 212, 1991. 38. Elliot, A. et al. Improved staining method for detecting Cryptosporidium oocysts in stools using malachite green. J. Gen. Appl. Microbiol. 45, 139, 1999. 39. Casemore, D.P. et al. Laboratory diagnosis of cryptosporidiosis. J. Clin. Pathol. 38, 1337, 1985. 40. Garcia, L.S. et al. Techniques for the recovery and identi�cation of Cryptosporidium oocysts from stool specimens. J. Clin. Microbiol. 18, 185, 1983. 41. Arrowood, M.J. and Sterling, C.R. Comparison of conventional staining methods and monoclonal antibody-based methods for Cryptosporidium oocyst detection. J. Clin. Microbiol. 27, 1490, 1989. 42. Geurden, T. et al. Prevalence and molecular characterisation of Cryptosporidium and Giardia in lambs and goat kids in Belgium. Vet. Parasitol. 155, 142, 2008. 43. Ortega, Y.R. et al. Ef�cacy of gaseous chlorine dioxide as a sanitizer against Cryptosporidium parvum, Cyclospora cayetanensis, and Encephalitozoon intestinalis on produce. J. Food Prot. 71, 2410, 2008.
44. Smith, A.L. Diagnostics. In: Fayer, R. and Xiao, L. (eds), Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL, p. 173, 2008.
45. Wang, Z. et al. Detection and genotyping of Entamoeba histolytica, Entamoeba dispar, Giardia lamblia and Cryptosporidium parvum by oligonucleotide microarray. J. Clin. Microbiol. 42, 3262, 2004.
46. Smith, H.V. et al. A microscopic system with a dual band �lter for the simultaneous enumeration of Cryptosporidium parvum oocysts and sporozoites. Water Res. 37, 2525, 2003.
47. Smith, H.V. and Ronald, A. Cryptosporidium: The analytical challenge. In: Smith, H.V. and Thompson, K. (eds), Cryptosporidium: The Analytical Challenge. The Royal Society of Chemistry, Cambridge, p. 1, 2001.
48. Millar, C.B. et al. Cryptosporidium in foodstuffs—An emerging aetiological route of human foodborne illness. Trends Food Sci. Technol. 13, 168, 2002.
49. Paton, C.A. et al. Immunomagnetisable separation for the recovery of Cryptosporidium sp. oocysts. In Clark, S.A. et al. (eds), Rapid Detection Assays for Food and Water. The Royal Society of Chemistry, Cambridge, U.K., p. 38, 2001.
50. Ferrari, B.C. et al. Comparison of Cryptosporidium-speci�c and Giardia-speci�c monoclonal antibodies for monitoring water samples. Water Res. 33, 1611, 1999.
51. Weber, R. et al. Threshold of detection of Cryptosporidium oocysts in human stool specimens: Evidence for low sensitivity of current diagnostic methods, J. Clin. Microbiol. 29, 1323, 1991.
52. Webster, K.A. et al. Detection of Cryptosporidium parvum oocysts in faeces: Comparison of conventional coproscopical methods and the polymerase chain reaction. Vet. Parasitol. 61, 5, 1996.
53. Kang, J.M. et al. Identi�cation and characterization of a mitochondrial iron-superoxide dismutase of Cryptosporidium parvum. Parasitol. Res. 103, 787, 2008. 54. Campbell, G.A. and Mutharasan, R. Near real-time detection of Cryptosporidium parvum oocyst by IgM-functionalized piezoelectric-excited millimeter-sized cantilever biosensor, Biosens. Bioelectron. 23, 1039, 2008.
55. Fayer, R. et al. Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). J. Parasitol. 91, 624, 2005.
56. Amar, C.F.L. et al. Blinded application of microscopy, bacteriological culture, immunoassays and PCR to detect gastrointestinal pathogens from faecal samples of patients with community-acquired diarrhoea. Eur. J. Clin. Microbiol. Infect. Dis. 23, 529, 2004.
57. Amar, C.F. et al. Detection and identi�cation by real time PCR/RFLP analyses of Cryptosporidium species from human faeces. Lett. Appl. Microbiol. 38, 217, 2004.
58. Sulaiman, I.M. et al. Evaluation of Cryptosporidium parvum genotyping techniques. Appl. Environ. Microbiol. 65, 4431, 1999.
59. Sturbaum, G.D. et al. Species-speci�c, nested PCRrestriction fragment length polymorphism detection of single Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 2665, 2001.
60. Xiao, L. et al. Phylogenetic analysis of Cryptosporidium parasites based on the small-subunit rRNA gene locus. Appl. Environ. Microbiol. 65, 1578, 1999.
61. Xiao, L. et al. Molecular characterization of Cryptosporidium oocysts in samples of raw surface water and wastewater. Appl. Environ. Microbiol. 67, 1097, 2001.
62. Ryan, U. et al. Identi�cation of novel Cryptosporidium genotypes from the Czech Republic. Appl. Environ. Microbiol. 69, 4302, 2003. 63. Jiang, J. and Xiao, L. An evaluation of molecular diagnostic tools for the detection and differentiation of human-pathogenic Cryptosporidium spp. J. Eukaryot. Microbiol. 50(Suppl), 542, 2003. 64. Loeb, K.R. et al. High-throughput quantitative analysis of hepatitis B virus DNA in serum using the TaqMan �uorogenic detection system. Hepatology 32, 626, 2000. 65. Higgins, J.A. et al. Real-time PCR for the detection of Cryptosporidium parvum. J. Microbiol. Methods 47, 323, 2001. 66. Garces-Sanchez, G. et al. Evaluation of two methods for quanti�cation of hsp70 mRNA from the waterborne pathogen Cryptosporidium parvum by reverse transcription real-time PCR in environmental samples. Water Res. 43, 2669, 2009. 67. Monis, P.T. et al. Emerging technologies for the detection and genetic characterization of protozoan parasites. Trends Parasitol. 21, 340, 2005. 68. Rasmussen, J.P. et al. Use of DNA melting simulation software for in silico diagnostic assay design: Targeting regions with complex melting curves and con�rmation by real-time PCR using intercalating dyes. BMC Bioinformat. 8, 107, 2007. 69. Xiao, L. et al. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72, 2004. 70. Gatei, W. et al. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect. Genet. Evol. 7, 197, 2007. 71. Caccio, S. et al. Large sequence variation at two microsatellite loci among zoonotic (genotype C) isolates of Cryptosporidium parvum. Int. J. Parasitol. 31, 1082, 2001. 72. Widmer, G. et al. Genotyping of Cryptosporidium parvum with microsatellite markers. Methods Mol. Biol. 268, 177, 2004. 73. Feng, X. et al. Extensive polymorphism in Cryptosporidium parvum identi�ed by multilocus microsatellite analysis. Appl. Environ. Microbiol. 66, 3344, 2000. 74. Strong, W.B. et al. Cloning and sequence analysis of a highly polymorphic Cryptosporidium parvum gene encoding a 60-kilodalton glycoprotein and characterization of its 15- and 45-kilodalton zoite surface antigen products. Infect. Immun. 68, 4117, 2000. 75. Plutzer, J. and Karanis, P. Genetic polymorphism in Cryptosporidium species: An update. Vet. Parasitol. 165, 187, 2009. 76. Leav, B.A. et al. Analysis of sequence diversity at the highly polymorphic Cpgp40/15 locus among Cryptosporidium isolates from human immunode�ciency virus-infected children in South Africa. Infect. Immun. 70, 3881, 2002. 77. Alves, M. et al. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J. Clin. Microbiol. 41, 2744, 2003. 78. Chalmers, R.M. et al. Direct comparison of selected methods for genetic categorisation of Cryptosporidium parvum and Cryptosporidium hominis species. Int. J. Parasitol. 35, 397, 2005. 79. Arrowood, M.J. and Donaldson, K. Improved puri�cation methods for calf-derived Cryptosporidium parvum oocysts using discontinuous sucrose and cesium chloride gradients. J. Eukaryot. Microbiol. 43, 89S, 1996. 80. Meloni, B.P. and Thompson, R.C. Simpli�ed methods for obtaining puri�ed oocysts from mice and for growing Cryptosporidium parvum in vitro. J. Parasitol. 82, 757, 1996. 81. Peng, M.M. et al. Molecular epidemiology of cryptosporidiosis in children in Malawi. J. Eukaryot. Microbiol. 50, S557, 2003. 82. Striepen, B. and Kissinger, J.C. Genomics meets transgenics in search of the elusive Cryptosporidium drug target. Trends Parasitol. 20, 355, 2004.
83. Foo, C. et al. Novel Cryptosporidium genotype in wild Australian mice (Mus domesticus). Appl. Environ. Microbiol. 73, 7693, 2007.
84. Sulaiman, I.M., Lal, A.A., and Xiao, L. Molecular phylogeny and evolutionary relationships of Cryptosporidium parasites at the actin locus. J. Parasitol. 88, 388, 2002.
85. Cacciò, S. et al. Genetic polymorphism at the beta-tubulin locus among human and animal isolates of Cryptosporidium parvum. FEMS Microbiol. Lett. 170, 173, 1999. Erratum in: FEMS Microbiol. Lett. 173, 273, 1999.
86. Spano, F. et al. PCR-RFLP analysis of the Cryptosporidium oocyst wall protein (COWP) gene discriminates between C. wrairi and C. parvum, and between C. parvum isolates of human and animal origin. FEMS Microbiol. Lett. 150, 209, 1997.
87. Xiao, L. et al. Sequence differences in the diagnostic target region of the oocyst wall protein gene of Cryptosporidium parasites. Appl. Environ. Microbiol. 66, 5499, 2000.
88. Gibbons, C.L., and Awad-El-Kariem, F.M. Nested PCR for the detection of Cryptosporidium parvum. Parasitol. Today 15, 345, 1999.
89. Morgan, U.M. et al. Phylogenetic relationships among isolates of Cryptosporidium: Evidence for several new species. J. Parasitol. 85, 1126, 1999.
90. Giles, M. et al. A multiplex allele speci�c polymerase chain reaction (MAS-PCR) on the dihydrofolate reductase gene for the detection of Cryptosporidium parvum genotypes 1 and 2. Parasitology 125, 35, 2002.
91. Sulaiman, I.M. et al. Phylogenetic relationships of Cryptosporidium parasites based on the 70-kilodalton heat shock protein (HSP70) gene. Appl. Environ. Microbiol. 66, 2385, 2000.
92. Feng, Y. et al. 90-kilodalton heat shock protein, Hsp90, as a target for genotyping Cryptosporidium spp. known to infect humans. Eukaryot. Cell 8, 478, 2009. 93. Carraway, M., Tzipori, S., and Widmer, G. Identi�cation of genetic heterogeneity in the Cryptosporidium parvum ribosomal repeat. Appl. Environ. Microbiol. 62, 712, 1996. 94. Carraway, M., Tzipori, S., and Widmer, G. A new restriction fragment length polymorphism from Cryptosporidium parvum identi�es genetically heterogeneous parasite populations and genotypic changes following transmission from bovine to human hosts. Infect. Immun. 65, 3958, 1997. 95. Spano, F. et al. Cryptosporidium parvum: PCR-RFLP analysis of the TRAP-C1 (thrombospondin-related adhesive protein of Cryptosporidium-1) gene discriminates between two alleles differentially associated with parasite isolates of animal and human origin. Exp. Parasitol. 90, 195, 1998. 96. Awad-el-Kariem, F.M., Warhurst, D.C., and McDonald, V. Detection and species identi�cation of Cryptosporidium oocysts using a system based on PCR and endonuclease restriction. Parasitology 109, 19, 1994. 97. Johnson, D.W. et al. Development of a PCR protocol for sensitive detection of Cryptosporidium oocysts in water samples. Appl. Environ. Microbiol. 61, 3849, 1995. 98. Leng, X., Mosier, D.A., and Oberst, R.D. Differentiation of Cryptosporidium parvum, C. muris, and C. baileyi by PCRRFLP analysis of the 18S rRNA gene. Vet Parasitol. 62, 1–7, 1996. 99. Bonnin, A. et al. Genotyping human and bovine isolates of Cryptosporidium parvum by polymerase chain reactionrestriction fragment length polymorphism analysis of a repetitive DNA sequence. FEMS Microbiol. Lett. 137, 207, 1996. 100. Morgan, U.M. et al. Differentiation between human and animal isolates of Cryptosporidium parvum using rDNA sequencing and direct PCR analysis. J. Parasitol. 83, 825, 1997. 16 Chapter 16 - Cyclospora
1. Eberhard, M.L. and Arrowood, M.J., Cyclospora spp., Curr. Opin. Infect. Dis., 15, 519, 2002.
2. Curry, A., and Smith, H.V., Emerging pathogens: Isospora, Cyclospora and microsporidia, Parasitology, 117, S143, 1998.
3. Herwaldt, B.L., Cyclospora cayetanensis: A review, focusing on the outbreaks of cyclosporiasis in the 1990s, Clin. Infect. Dis., 31, 1040, 2000. 4. Ortega, Y.R., Gilman, R.H., and Sterling, C.R., A new coccidian parasite (Apicomplexa: Eimeriidae) from humans, J. Parasitol., 80, 625, 1994. 5. Shields, J.M., and Olson, B.H., Cyclospora cayetanensis: A review of an emerging parasitic coccidian, Int. J. Parasitol., 33, 371, 2003. 6. Lainson, R., The genus Cyclospora (Apicomplexa: Eimeriidae), with a description of Cyclospora schneideri n.sp. in the snake Anilius scytale scytale (Aniliidae) from Amazonian Brazil—A review, Mem. Inst. Oswaldo Cruz, 100, 103, 2005. 7. Smith, H.V. et al., Sporulation of Cyclospora sp. oocysts, Appl. Environ. Microbiol., 63, 1631, 1997. 8. Ortega, Y.R., Sterling, C.R., and Gilman, R.H., Cyclospora cayetanensis, Adv. Parasitol., 40, 399, 1998. 9. Chacin-Bonilla, L. et al., Cyclospora cayetanensis infections among diarrheal patients from Venezuela, Am. J. Trop. Med. Hyg., 65, 351, 2001. 10. Chacin-Bonilla, L. et al., Prevalence and pathogenic role of Cyclospora cayetanensis in a Venezuelan community, Am. J. Trop. Med. Hyg., 68, 304, 2003. 11. Fleming, C.A. et al., A foodborne outbreak of Cyclospora cayetanensis at a wedding: Clinical features and risk factors for illness, Arch. Intern. Med., 158, 1121, 1998. 12. Mead, P.S., Food-related illness and death in the United States, Emerg. Infect. Dis., 5, 607, 1999. 13. Crist, A. et al., Outbreak of cyclosporiasis associated with snow peas—Pennsylvania, 2004, MMWR, 53, 876, 2004. 14. Gascon, J. et al., Cyclosporiasis: A clinical and epidemiological study in travellers with imported Cyclospora cayetanensis infection, Med. Clin. (Barc.), 116, 461, 2001. 15. Ashford, R.W., Occurrence of an undescribed coccidian in man in Papua New Guinea, Ann. Trop. Med. Parasitol., 73, 497, 1979. 16. Nunez, F.A. et al., The �rst report in Cuba of human intestinal infection by Cyclospora cayetanensis, Ortega, 1993, Rev. Cubana Med. Trop., 47, 211, 1995. 17. Clarke, S.C., and McIntyre, M., The incidence of Cyclospora cayetanensis in stool samples submitted to a district general hospital, Epidemiol. Infect., 117, 189, 1996. 18. Madico, G. et al., Epidemiology and treatment of Cyclospora cayetanensis infection in Peruvian children, Clin. Infect. Dis., 24, 977, 1997. 19. Graczyk, T.K., Ortega, Y.R., and Conn, D.B., Recovery of waterborne oocysts of Cyclospora cayetanensis by Asian freshwater clams (Corbicula °uminea), Am. J. Trop. Med. Hyg., 59, 928, 1998. 20. Bern, C. et al., Epidemiologic studies of Cyclospora cayetanensis in Guatemala, Emerg. Infect. Dis., 5, 766, 1999. 21. Fryauff, D.J. et al., Cyclospora cayetanensis among expatriate and indigenous populations of West Java, Indonesia, Emerg. Infect. Dis., 5, 585, 1999. 22. Sherchand, J.B. and Cross, J.H., Emerging pathogen Cyclospora cayetanensis infection in Nepal, Southeast Asian J. Trop. Med. Public. Health, 32, S143, 2001. 23. Wang, K.X. et al., Cyclospora cayetanensis in Anhui, China, World J. Gastroenterol., 8, 1144, 2002. 24. Lopez, A.S. et al., Outbreak of cyclosporiasis associated with basil in Missouri in 1999, Clin. Infect. Dis., 32, 1010, 2001. 25. Nimri, L.F., Cyclospora cayetanensis and other intestinal parasites associated with diarrhea in a rural area of Jordan, Int. Microbiol., 6, 131, 2003.
26. Zerpa, R., Uchima, N., and Huicho, L., Cyclospora cayetanensis associated with watery diarrhoea in Peruvian patients, J. Trop. Med. Hyg., 98, 325, 1995.
27. Marshall, M.M. et al., Waterborne protozoan pathogens, Clin. Microbiol. Rev., 10, 67, 1997.
28. Wright, M.S., and Collins, P.A., Waterborne transmission of Cryptosporidium, Cyclospora and Giardia, Clin. Lab. Sci., 10, 287, 1997.
29. Sherchand, J.B. et al., Study of Cyclospora cayetanensis in health care facilities, sewage water and green leafy vegetables in Nepal, Southeast Asian J. Trop. Med. Public Health, 30, 58, 1999.
30. Sterling, C.R., and Ortega, Y.R., Cyclospora: An enigma worth unraveling, Emerg. Infect. Dis., 5, 48, 1999.
31. Mota, P., Rauch, C.A., and Edberg, S.C., Microsporidia and Cyclospora: Epidemiology and assessment of risk from the environment, Crit. Rev. Microbiol., 26, 69, 2000.
32. Dowd, S.E. et al., Con�rmed detection of Cyclospora cayetanesis, Encephalitozoon intestinalis and Cryptosporidium parvum in water used for drinking, J. Water Health, 1, 117, 2003.
33. Mans�eld, L.S., and Gajadhar, A.A., Cyclospora cayetanensis, a food- and waterborne coccidian parasite, Vet. Parasitol., 126, 73, 2004.
34. Eberhard, M.L. et al., Cyclospora cayetanensis infections in Haiti: A common occurrence in the absence of watery diarrhea, Am. J. Trop. Med. Hyg., 60, 584, 1999.
35. Chalmers, R.M., Nichols, G., and Rooney, R., Foodborne outbreaks of cyclosporiasis have arisen in North America. Is the United Kingdom at risk? Commun. Dis. Public Health, 3, 50, 2000.
36. Ho, A.Y. et al., Outbreak of cyclosporiasis associated with imported raspberries, Philadelphia, Pennsylvania, 2000, Emerg. Infect. Dis., 8, 783, 2000.
37. Lopez, F.A. et al., Molecular characterization of Cyclosporalike organisms from baboons, J. Infect. Dis., 179, 670, 1999.
38. Döller, P.C. et al., Cyclosporiasis outbreak in Germany associated with the consumption of salad, Emerg. Infect. Dis., 8, 992, 2002.
39. Connor, B.A., Reidy, J., and Soave, R., Cyclosporiasis: Clinical and histopathologic correlates, Clin. Infect. Dis., 28, 1216, 1999.
40. Ortega, Y.R. et al., Isolation of Cryptosporidium parvum and Cyclospora cayetanensis from vegetables collected in markets of an endemic region in Peru, Am. J. Trop. Med. Hyg., 57, 683, 1997.
41. Eberhard, M.L, Pieniazek, N.J., and Arrowood, M.J., Laboratory diagnosis of Cyclospora infections, Arch. Pathol. Lab. Med., 121, 792, 1997.
42. Visvesvara, G.S. et al., Uniform staining of Cyclospora oocysts in fecal smears by a modi�ed safranin technique with microwave heating, J. Clin. Microbiol., 35, 730, 1997.
43. Varma, M. et al., Detection of Cyclospora cayetanensis using a quantitative real-time PCR assay, J. Microbiol. Methods, 53, 27, 2003.
44. Abou el Naga, I.F., Studies on a newly emerging protozoal pathogen: Cyclospora cayetanensis, J. Egypt. Soc. Parasitol., 29, 575, 1999. 45. Ortega, Y.R. et al., Pathologic and clinical �ndings in patients with cyclosporiasis and a description of intracellular parasite life-cycle stages, J. Infect. Dis., 176, 1584, 1997.
46. Robertson, L.J., Gjerde, B., and Campbell, A.T., Isolation of Cyclospora oocysts from fruits and vegetables using lectincoated paramagnetic beads, J. Food Prot., 63, 1410, 2000. 47. Jinneman, K.C. et al., Template preparation for PCR and RFLP of ampli�cation products for the detection and identi�cation of Cyclospora sp. and Eimeria spp. oocysts directly from raspberries, J. Food Prot., 61, 1497, 1998. 48. Orlandi, P.A., and Lampel, K.A., Extraction-free, �lter-based template preparation for rapid and sensitive PCR detection of pathogenic parasitic protozoa, J. Clin. Microbiol., 38, 2271, 2000. 49. Sturbaum, G.D. et al., Species-speci�c, nested PCRrestriction fragment length polymorphism detection of single Cryptosporidium parvum oocysts, Appl. Environ. Microbiol., 67, 2665, 2001. 50. Dalton, C. et al., Viability of Giardia intestinalis cysts and viability and sporulation state of Cyclospora cayetanensis oocysts determined by electrorotation, Appl. Environ. Microbiol., 67, 586, 2001. 51. Quintero-Betancourt, W., Peele, E.R., and Rose, J.B., Cryptosporidium parvum and Cyclospora cayetanensis: A review of laboratory methods for detection of these waterborne parasites, J. Microbiol. Methods, 49, 209, 2002. 52. Shields, J.M., and Olson, B.H., PCR-restriction fragment length polymorphism method for detection of Cyclospora cayetanensis in environmental waters without microscopic con�rmation, Appl. Environ. Microbiol., 69, 4662, 2003. 53. Steele, M., Unger, S., and Odumeru, J., Sensitivity of PCR detection of Cyclospora cayetanensis in raspberries, basil, and mesclun lettuce, J. Microbiol. Methods, 54, 277, 2003. 54. Veiweij, J.J. et al., Detection of Cyclospora cayetanensis in travellers returning from the tropics and subtropics using microscopy and real-time PCR, Int. J. Med. Microbiol., 293, 199, 2003. 55. Mundaca, C.C. et al., Use of PCR to improve diagnostic yield in an outbreak of cyclosporiasis in Lima, Peru, Trans. R. Soc. Trop. Med. Hyg., 102, 712, 2008. 56. Relman, D.A. et al., Molecular phylogenetic analysis of Cyclospora, the human intestinal pathogen, suggests that it is closely related to Eimeria species, J. Infect. Dis., 173, 440, 1996. 57. Jinneman, K.C. et al., An oligonucleotide-ligation assay for the differentiation between Cyclospora and Eimeria spp. polymerase chain reaction ampli�cation products. J. Food Prot., 62, 682, 1999. 58. Sturbaum, G.D. et al., Detection of Cyclospora cayetanensis in wastewater, Appl. Environ. Microbiol., 64, 2284, 1998. 59. Eberhard, M.L. et al., Morphologic and molecular characterization of new Cyclospora species from Ethiopian monkeys: C. cercopitheci sp. n., C. colobi sp. n., and C. papionis sp. n, Energ. Inf. Dis., 5, 651, 1999. 60. Lopez, A.S. et al., Epidemiology of Cyclospora cayetanensis and other intestinal parasites in a community in Haiti, J. Clin. Microbiol., 41, 2047, 2003. 61. Chu, D.M. et al., Detection of Cyclospora cayetanensis in animal fecal isolates from Nepal using an FTA �lter-base polymerase chain reaction method, Am. J. Trop. Med. Hyg., 71, 373, 2004. 62. Orlandi, P.A. et al., Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR, Appl. Environ. Microbiol., 69, 4806, 2003. 63. Hussein, E.M., Molecular identi�cation of Cyclospora spp. using multiplex PCR from diarrheic children compared to others conventional methods, J. Egypt Soc. Parasitol., 37, 585, 2007.
64. Hussein, E.M. et al., Real-time PCR and �ow cytometry in detection of Cyclospora oocysts in fecal samples of symptomatic and asymptomatic pediatrics patients, J. Egypt Soc. Parasitol., 37, 151, 2007.
65. Adam, R.D. et al., Intervening transcribed spacer region 1 variability in Cyclospora cayetanensis, J. Clin. Microbiol., 38, 2339, 2000.
66. Olivier, C. et al., Sequence variability in the �rst internal transcribed spacer region within and among Cyclospora species is consistent with polyparasitism, Int. J. Parasitol., 31, 1475, 2001.
67. Lalonde, L.F., and Gajadhar, A.A., Highly sensitive and speci�c PCR assay for reliable detection of Cyclospora cayetanensis oocysts, Appl. Environ. Microbiol., 74, 4354, 2008.
68. Vesey, G. et al., A new method for the concentration of Cryptosporidium oocysts from water, J. Appl. Bacteriol., 75, 82, 1993.
69. da Silva, A.J. et al., Fast and reliable extraction of protozoan parasite DNA from fecal specimens, Mol. Diagn., 4, 57, 1999.
70. Ortega, Y.R., and Sanchez, R., Update on Cyclospora cayetanensis, a food-borne and waterborne parasite, Clin. Microbiol. Rev., 23, 218, 2010.
71. Chacín-Bonilla, L., Epidemiology of Cyclospora cayetanensis: A review focusing in endemic areas, Acta Trop., 115, 181, 2010.
72. Helmy, M.M., Cyclospora cayetanensis: A review, focusing on some of the remaining questions about cyclosporiasis, Infect. Disord. Drug Targets, 10, 368, 2010. 73. Riner, D.K. et al., Intragenomic sequence variation of the ITS-1 region within a single �ow-cytometry-counted Cyclospora cayetanensis oocysts, J. Parasitol., 96, 914, 2010. 74. Taniuchi, M. et al., Multiplex polymerase chain reaction method to detect Cyclospora, Cystoisospora, and Microsporidia in stool samples, Diagn. Microbiol. Infect. Dis., 71, 386, 2011. 75. Masucci, L. et al., First Italian case of cyclosporiasis in an immunocompetent woman: Local acquired infection, New Microbiol., 31, 281, 2008. 76. Naito, T. et al., Cyclospora infection in an immunocompetent patient in Japan, Jpn. J. Infect. Dis., 62, 57, 2009. 77. Ozdamar, M., Hakko, E., and Turkoglu, S., High occurrence of cyclosporiasis in Istanbul, Turkey, during a dry and warm summer. Parasit. Vectors, 3, 39, 2010. 78. Insulander, M. et al., A foodborne outbreak of Cyclospora infection in Stockholm, Sweden, Foodborne Pathog. Dis., 7, 1585, 2010. 79. Hall, R.L., Jones, J.L., and Herwaldt, B.L., Surveillance for laboratory-con�rmed sporadic cases of cyclosporiasis—United States, 1997-2008, MMWR Surveil.l Summ., 60, 1, 2011. 80. Iqbal, J. et al., Cyclospora cayetanensis: First report of imported and autochthonous infections in Kuwait, J. Infect. Dev. Ctries., 5, 383, 2011. 81. Milord, F. et al., Cyclospora cayetanensis: A description of clinical aspects of an outbreak in Quebec, Canada. Epidemiol. Infect., 140, 626, 2012. 82. Eberhard, M.L. et al., Attempts to establish experimental Cyclospora cayetanensis infection in laboratory animals, J. Parasitol., 86, 577, 2000. 17 Chapter 17 - Isospora
1. Wenyon, C.M. Coccidiosis of cats and dogs and the status of the Isospora of man. Ann. Trop. Med. Parasitol. 17, 231, 1923.
2. Elsdon-Dew, R. Isospora natalensis (sp. Nov.) in man. J. Trop. Med. Hyg. 56, 149, 1953.
3. Ravenel, J.M., Suggs, J.L., and Legerton, C.W. Human coccidiosis. Recurrent diarrhea of 26 years duration due to Isospora belli: A case report. J. S. Carolina Med. Assoc. 72, 217, 1976.
4. Jongwutiwes, S., Sampatanukul, P., and Putaporntip, C. Recurrent isosporiasis over a decade in an immunocompetent host successfully treated with pyrimethamine. Scand. J. Infect. Dis. 34, 859, 2002.
5. Franzen, C. et al. Taxonomic position of the intestinal protozoan parasite Isospora belli as based on ribosomal RNA sequences. Parasitol. Res. 86, 669, 2000.
6. Morrison. D.A. et al. The current status of the small subunit rRNA phylogeny of the coccidia (Sporozoa). Int. J. Parasitol. 34, 501, 2004.
7. Jongwutiwes, S. et al. Morphologic and molecular characterization of Isospora belli oocysts from patients in Thailand. Am. J. Trop. Med. Hyg. 77, 107, 2007.
8. Dubey, J.P., and Frenkel, J.K. Extra-intestinal stages of Isospora felis and I. rivolta (Protozoa: Eimeriidae) in cats. J. Protozool. 19, 89, 1972.
9. Restrepo, C., Macher, A.M., and Radany, E.H. Disseminated extraintestinal isosporiasis in a patient with acquired immune de�ciency syndrome. Am. J. Clin. Pathol. 87, 536, 1987.
10. Michiels, J.F. et al. Intestinal and extraintestinal Isospora belli infection in an AIDS patient. A second case report. Pathol. Res. Pract. 190, 1089, 1994.
11. Lindsay, D.S., Dubey, J.P., and Blagburn, B.L. Biology of Isospora spp. from humans, nonhuman primates, and domestic animals. Clin. Microbiol. Rev. 10, 19, 1997.
12. Tamura, K. et al. MEGA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596, 2007. 13. Beaver, P.C., Jung, R.C., and Cupp, E.W. Coccidia, Microsporidia and Pneumocystis. In: Beaver, P.C., Jung, R.C., and Cupp, E.W. (eds), Clinical Parasitology, 9th edn. Lea and Febiger, Philadelphia, PA, pp. 149–173, 1984. 14. Zaman, V. Observations on human Isospora. Trans. R. Soc. Trop. Med. Hyg. 62, 556, 1968. 15. Matsui, T. et al. Infectivity and sporogony of Caryosporatype oocyst of Isospora rivolta obtained by heating. Parasitol. Res., 79, 599, 1993. 16. Ferguson, D.J.P. et al. Ultrastructural observations on multiplication of Cystoisospora (Isospora) felis by endodyogeny. Z. Parasitenkd. 63, 289, 1980. 17. Matuschka, F.R. Ultrastructural evidence of endodyogeny in Isospora suis from pigs. Z. Parasitenkd. 67, 27, 1982. 18. Velásquez, J.N. et al. Isosporosis and unizoite tissue cysts in patients with acquired immunode�ciency syndrome. Hum. Pathol. 32, 500, 2001. 19. Frenkel, J.K. et al. Isospora belli infection: Observation of unicellular cysts in mesenteric lymphoid tissues of a Brazilian patient with AIDS and animal inoculation. J. Eukaryot. Microbiol. 50(Suppl), 682, 2003. 20. Lindsay, D.S. et al. Examination of extraintestinal tissue cysts of Isospora belli. J. Parasitol. 83, 620, 1997. 21. Faust, E.C. et al. Human isosporosis in the Western hemisphere. Am. J. Trop. Med. Hyg. 10, 343, 1961. 22. Henry, M.C. et al. Parasitological observations of chronic diarrhoea in suspected AIDS adult patients in Kinshasa (Zaire). Trans. R. Soc. Trop. Med. Hyg. 80, 309, 1986. 23. DeHovitz, J.A. et al. Clinical manifestations and therapy of Isospora belli infection in patients with the acquired immunode�ciency syndrome. N. Engl. J. Med. 315, 87, 1986. 24. Certad, G. et al. Isosporiasis in Venezuelan adults infected with human immunode�ciency virus: Clinical characterization. Am. J. Trop. Med. Hyg. 69, 217, 2003. 25. Lainson, R. and da Silva, B.A. Intestinal parasites of some diarrhoeic HIV-seropositive individuals in North Brazil, with particular reference to Isospora belli Wenyon, 1923 and Dientamoeba fragilis Jepps & Dobell, 1918. Mem. Inst. Oswaldo. Cruz. 94, 611, 1999. 26. Sorvillo, F.J. et al. Epidemiology of isosporiasis among persons with acquired immunode�ciency syndrome in Los Angeles County. Am. J. Trop. Med. Hyg. 53, 656, 1995. 27. Brandborg, L.L., Goldberg, S.B., and Breidenbach, W.C. Human coccidiosis: A possible cause of malabsorption. N. Engl. J. Med. 283, 1306, 1970. 28. Trier, J.S. et al. Chronic intestinal coccidiosis in man: Intestinal morphology and response to treatment. Gastroenterol. 66, 923, 1974. 29. Matsubayashi, H. and Nozawa, T. Experimental infection of Isospora hominis in man. Am. J. Trop. Med. 28, 633, 1948. 30. McCracken, A.W. Natural and laboratory-acquired infection by Isospora belli. South. Med. J. 65, 800, 1972. 31. Syrkis, I. et al. A case of severe human coccidiosis in Israel. Israel. J. Med. Sci., 1, 373, 1975. 32. Pape, J.W., Verdier, R.I., and Johnson, W.D. Jr. Treatment and prophylaxis of Isospora belli infection in patients with the acquired immunode�ciency syndrome. N. Engl. J. Med., 320, 1044, 1989. 33. de Otazu, R.D. et al. Endometrial coccidiosis. J. Clin. Pathol., 57, 1104, 2004. 34. Herwaldt, B.L. Cyclospora cayetanensis: A review, focusing on the outbreaks of cyclosporiasis in the 1990s. Clin. Infect. Dis. 31, 1040, 2000.
35. Garcia, L.S. Collection, preservation, and shipment of fecal specimens. In: Garcia, L.S. (ed), Diagnostic Medical Parasitology, 4th edn. ASM Press, Washington, DC, pp. 723–740, 2001.
36. Jongwutiwes, S. et al. Simple method for long-term copropreservation of Cryptosporidium oocysts for morphometric and molecular analysis. Trop. Med. Int. Health. 7, 257, 2002.
37. Garcia, L.S. Macroscopic and microscopic examination of fecal specimens. In: Garcia, L.S. (ed), Diagnostic Medical Parasitology, 4th edn. ASM Press, Washington, DC, pp. 741–785, 2001.
38. Nime, F.A. et al. Acute enterocolitis in a human being infected with the protozoan Cryptosporidium. Gastroenterol. 70, 592, 1976.
39. Meisel, J.L. et al. Overwhelming watery diarrhea associated with a Cryptosporidium in an immunosuppressed patient. Gastroenterol. 70, 1156, 1976.
40. Ritchie, L. An ether sedimentation technique for routine stool examinations. Bull. U.S. Army Med. Dept. 8, 326, 1948.
41. Young, K.H. et al. Ethyl acetate as a substitute for diethyl ether in the formalin-ether sedimentation technique. J. Clin. Microbiol., 10, 852, 1979.
42. Garcia, L.S. Sputum, aspirates, and biopsy material. In: Garcia, L.S. (ed), Diagnostic Medical Parasitology, 4th edn. ASM Press, Washington, DC, pp. 809–828, 2001.
43. Faust, E.C. et al. A critical study of clinical laboratory technics for the diagnosis of protozoan cysts and helminth eggs in feces. Am. J. Trop. Med. 18, 169, 1938. 44. Ma, P., and Soave, R. Three-step stool examination for cryptosporidiosis in 10 homosexual men with protracted watery diarrhea. J. Infect. Dis. 147, 824, 1983. 45. Varea, M. et al. Fuchsin �uorescence and auto�uorescence in Cryptosporidium, Isospora and Cyclospora oocysts. Int. J. Parasitol. 28, 1881, 1998. 46. Daugschies, A. et al. Auto�uorescence microscopy for the detection of nematode eggs and protozoa, in particular Isospora suis, in swine feces. Parasitol. Res. 87, 409, 2001. 47. Bialek, R. et al. Comparison of auto�uorescence and iodine staining for detection of Isospora belli in feces. Am. J. Trop. Med. Hyg. 67, 304, 2002. 48. Sun, T. et al. Light and electron microscopic identi�cation of Cyclospora species in the small intestine. Evidence of the presence of asexual life cycle in human host. Am. J. Clin. Pathol. 105, 216, 1996. 49. Müller, A. et al. Detection of Isospora belli by polymerase chain reaction using primers based on small-subunit ribosomal RNA sequences. Eur. J. Clin. Microbiol. Infect. Dis. 19, 631, 2000. 50. Joachim, A. et al. Detection of Isospora suis (Biester and Murray 1934) in piglet faeces: Comparison of microscopy and PCR. J. Vet. Med. B. 51, 140, 2004. 51. QIAamp DNA Stool Mini Kit Handbook. For DNA puri�cation from stool samples, August 2001. 52. QIAGEN DNeasy ® Blood & Tissue Handbook. For puri�cation of total DNA from animal tissue, July 2006. 18 Chapter 18 - Plasmodium
TABLE 18.4
Primers and Probes for Real-Time PCR Identification of Five Human-Infecting Plasmodium Species
Primer Sequence (5′–3′)
Pan-species forward GTT AAG GGA GTG AAG ACG ATC AGA TA
Pan-species reverse AAC CCA AAG ACT TTG ATT TCT CAT AAG
P. falciparum 18S rRNA forward ATT GCT TTT GAG AGG TTT TGT TAC TTT
P. falciparum 18S rRNA forward GCT GTA GTA TTC AAA CAC AAT GAA CTC AA
P. ovale 18S rRNA forward CCG ACT AGG TTT TGG ATG AAA GAT TTT T
P. ovale 18S rRNA reverse CAA CCC AAA GAC TTT GAT TTC TCA TAA
P. malariae 18S rRNA forward AGT TAA GGG AGT GAA GAC GAT CAG A
P. malariae 18S rRNA reverse CAA CCC AAA GAC TTT GAT TTC TCA TAA
P. falciparum LDH forward ACG ATT TGG CTG GAG CAG AT
P. falciparum LDH reverse TCT CTA TTC CAT TCT TTG TCA CTC TTT C
Human GAPDH forward CCT CCC GCT TCG CTC TCT
Human GAPDH reverse GCT GGC GAC GCA AAA GA MGB Probe
Pan-species VIC-TCG TAA TCT TAA CCA TAA AC
P. falciparum FAM-CAT AAC AGA CGG GTA GTC AT
P. ovale VIC-CGA AAG GAA TTT TCT TAT T
P. malariae FAM-ATG AGT GTT TCT TTT AGA TAG C
Human GAPDH VIC-CCT CCT GTT CGA CAG TCA GCC GC TaqMan Probe
P. falciparum LDH FAM-GTA ATA GTA ACA GCT GGA TTT ACC AAG GCC CCA-TAMRA
7. Collins, W.E., and Jeffery, G.M., Plasmodium ovale: Parasite and disease. Clin. Microbiol. Rev., 18, 570, 2005.
8. Collins, W.E., and Jeffery, G.M., Plasmodium malariae: Parasite and disease. Clin. Microbiol. Rev., 20, 579, 2007.
9. Win, T.T. et al., Wide distribution of Plasmodium ovale in Myanmar. Trop. Med. Int. Health, 7, 231, 2002.
10. Jongwutiwes, S. et al., Naturally acquired Plasmodium knowlesi malaria in human, Thailand. Emerg. Infect. Dis., 10, 2211, 2004.
11. Singh, B. et al., A large focus of naturally acquired Plasmodium knowlesi infections in human beings. Lancet, 363, 1017, 2004.
12. Vythilingam, I. et al., Natural transmission of Plasmodium knowlesi to humans by Anopheles latens in Sarawak, Malaysia. Trans. R. Soc. Trop. Med. Hyg., 100, 1087, 2006.
13. Luchavez, J. et al., Human infections with Plasmodium knowlesi, the Philippines. Emerg. Infect. Dis., 14, 811, 2008.
14. White, N.J., Plasmodium knowlesi: The �fth human malaria parasite. Clin. Infect. Dis., 46, 172, 2008.
15. Daneshvar, C. et al., Clinical and laboratory features of human Plasmodium knowlesi infection. Clin. Infect. Dis., 49, 852, 2009.
16. Van den Eede, P. et al., Human Plasmodium knowlesi infections in young children in central Vietnam. Malar. J., 8, 249, 2009.
17. Oddoux, O. et al., Identi�cation of the �ve human Plasmodium species including P. knowlesi by real-time polymerase chain reaction. Eur. J. Clin. Microbiol. Infect. Dis., 30, 597, 2011.
18. Sulistyaningsih, E. et al., Diagnostic dif�culties with Plasmodium knowlesi infection in humans. Emerg. Infect. Dis., 16, 1033, 2010.
19. Escalante, A.A. et al., The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochrondrial genome. Proc. Natl. Acad. Sci. USA, 95, 8124, 1988.
20. Greenwood, B., The molecular epidemiology of malaria. Trop. Med. Int. Health, 7, 1012, 2002.
21. Krief, S. et al., On the diversity of malaria parasites in African apes and the origin of Plasmodium falciparum from bonobos. PloS Pathog., 6, e1000765, 2010.
22. Liu, W. et al., Origin of the human malaria parasite Plasmodium falciparum in gorillas. Nature, 23, 420, 2010.
23. Ricklefs, R.E., and Outlaw, D.C., A molecular clock for malaria parasites. Science, 329, 226, 2010.
24. Zhou, M. et al., High prevalence of Plasmodium malariae and Plasmodium ovale in malaria patients along the ThaiMyanmar border, as revealed by acridine staining and PCRbased diagnoses. Top. Med. Int. Health, 3, 304, 1998.
25. Muehlenbachs, A. et al., An unusual presentation of placental malaria: A single persisting nidus of sequestered parasites. Hum. Pathol., 38, 520, 2007.
26. Bruneel, F. et al., Severe imported falciparum malaria: A cohort study in 400 critically ill adults. PLoS One, 5, e13236, 2010.
27. Sawai, H. et al., Lineage-speci�c positive selection at the merozoite surface protein 1 (msp1) locus of Plasmodium vivax and related simian malaria parasites. Evol. Biol., 10, 52, 2010.
28. Shiff, C.J., Premij, Z., and Minjas, J.N., The rapid ParaSight ™ -F test. A new diagnostic tool for Plasmodium falciparum infection. Trans. R. Soc. Trop. Med. Hyg., 87, 29, 1993.
29. Warhurst, D.C., and Williams, J.E., Laboratory diagnosis of malaria. J. Clin. Pathol., 49, 533, 1996.
30. Moody, A., Rapid diagnostic tests for malaria parasites. Clin. Microbiol. Rev., 15, 66, 2002. 31. Rakotonirina, H. et al., Accuracy and reliability of malaria diagnostic techniques for guiding febrile outpatient treatment in malaria-endemic countries. Am. J. Trop. Med. Hyg., 78, 217, 2008. 32. Kawamoto, F., and Billingsley, P.F., Rapid diagnosis of malaria by �uorescence microscopy. Parasitol. Today, 8, 69, 1992. 33. Nkrumah, B. et al., Comparison of the novel Partec rapid malaria test to the conventional Giemsa stain and the gold standard real-time PCR. J. Clin. Microbiol., 48, 2925, 2010. 34. Rantala, A.M. et al., Comparison of real-time PCR and microscopy for malaria parasite detection in Malawian pregnant women. Malar. J., 9, 269, 2010. 35. Matsumoto, Y., Matsuda, S., and Yoshida, Y., Ultrastructure of erythrocytic stages of Plasmodium ovale in humans. Am. J. Trop. Med. Hyg., 35, 689, 1986. 36. Atkinson, C.T. et al., Ultrastructure of the erythrocytic stages of Plasmodium malariae. J. Protozool., 34, 267, 1987. 37. Millet, P. et al., In vitro cultivation of exoerythrocytic stages of the human malaria parasite Plasmodium malariae. Am. J. Trop. Med. Hyg., 38, 470, 1988. 38. McKenzie, F.E., Collins, W.E., and Jeffery, G.M., Plasmodium malariae blood-stage dynamics. J. Parasitol., 87, 626, 2001. 39. Lee, K.-S. et al., Morphological features and differential counts of Plasmodium knowlesi parasites in naturally acquired human infections. Malar. J., 8, 73, 2009. 40. Ndao, M. et al., Comparison of blood smear, antigen detection, and nested-PCR methods for screening refugees from regions where malaria is endemic after a malaria outbreak in Quebec, Canada. J. Clin. Microbiol., 42, 2694, 2004. 41. Khairnar, K. et al., Multiplex real-time quantitative PCR, microscopy and rapid diagnostic immuno-chromatographic tests for the detection of Plasmodium spp: Performance, limit of detection analysis and quality assurance. Malar. J., 8, 284, 2009. 42. Beier, M.S. et al., Identi�cation of malaria species by ELISA in sporozoite and oocyst infected Anopheles from western Kenya. Am. J. Trop. Med. Hyg., 39, 323, 1988. 43. Collins, F.H. et al., Monoclonal antibody-based enzymelinked immunosorbent assay (ELISA) for detection of Plasmodium malariae sporozoites in mosquitoes. Am. J. Trop. Med. Hyg., 38, 283, 1988. 44. Relf, W.A. et al., Diagnosis of Plasmodium vivax malaria using a speci�c deoxyribonucleic acid probe. Trans. R. Soc. Trop. Med. Hyg., 84, 630, 1990. 45. McLaughlin, G.L. et al., Evaluation of non-isotopic DNA assay kit for diagnosing Plasmodium falciparum malaria in Madagascar. Am. J. Trop. Med. Hyg., 48, 211, 1993. 46. Contamin, H. et al., PCR typing of �eld isolates of Plasmodium falciparum. J. Clin. Microbiol., 33, 944, 1995. 47. Oliveira, D.A. et al., Polymerase chain reaction and a liquidphase nonisotopic hybridization for species-speci�c and sensitive detection of malaria infection. Am. J. Trop. Med. Hyg., 52, 139, 1995. 48. Stoffels, S.A.W., Docters van Leeuwen, W.M., and Post, R.J., Detection of Plasmodium-infectious mosquitoes by polymerase chain reaction without dissection of the salivary glands. Med. Vet. Entomol., 9, 433, 1995. 49. Vu, T.T. et al., Screening donor blood for malaria by polymerase chain reaction. Trans. R. Soc. Trop. Med. Hyg., 89, 44, 1995. 50. Kawamoto, F. et al., Sequence variation in the 18S rRNA gene, a target for PCR-based malaria diagnosis, in Plasmodium ovale from southern Vietnam. Microbiology, 34, 2287, 1996.
51. Singh, B. et al., Detection of malaria in Malaysia by nested polymerase chain reaction ampli�cation of dried blood spots on �lter papers. Trans. R. Soc. Trop. Med. Hyg., 90, 519, 1996.
52. Singh, B. et al., A genus- and species-speci�c nested polymerase chain reaction malaria detection assay for epidemiologic studies. Am. J. Trop. Med. Hyg., 60, 687, 1999.
53. Cox-Singh, J. et al., Increased sensitivity of malaria detection by nested polymerase chain reaction using simple sampling and DNA extraction. Int. J. Parasitol., 27, 1575, 1997.
54. Tahar, R., Ringwald, P., and Basco, L.K., Diagnosis of Plasmodium malariae infection by the polymerase chain reaction. Trans. R. Soc. Trop. Med. Hyg., 91, 410, 1997.
55. Rubio, J.M. et al., Usefulness of seminested multiplex PCR in surveillance of imported malaria in Spain. J. Clin. Microbiol., 37, 3260, 1999.
56. Rubio, J.M. et al., Alternative polymerase chain reaction method to identify Plasmodium species in human blood samples: The semi-nested multiplex malaria PCR (SnM-PCR). Trans. R. Soc. Trop. Med. Hyg., S199, 96, 2002.
57. Manca, N. et al., Search for malaria parasites by PCR and Southern blot in patients with imported malaria in Italy. New Microbiol., 23, 339, 2000.
58. Perandin, F. et al., Usefulness of genus-speci�c PCR and Southern blot species-speci�c hybridization for the detection of imported malaria cases in Italy. New Microbiol., 24, 69, 2001.
59. Padley, D. et al., Use of a rapid, single-round, mutiplex PCR to detect malarial parasites and identify the species present. Ann. Trop. Med. Parasitol., 97, 131, 2003.
60. Berry, A. et al., Contribution of PCR-based methods to diagnosis and management of imported malaria. Med. Trop. (Mars), 65, 176, 2005.
61. McNamara, D.T. et al., Diagnosing infection levels of four human malaria parasite species by a polymerase chain reaction/ligase detection reaction �uorescent microsphere-based assay. Am. J. Trop. Med. Hyg., 74, 413, 2006.
62. Boonma, P. et al., Comparison of three molecular methods for the detection and speciation of Plasmodium vivax and Plasmodium falciparum. Malar. J., 6, 124, 2007.
63. Gama, B.E. et al., Real-time PCR versus conventional PCR for malaria parasite detection in low-grade parasitemia. Exp. Parasitol., 116, 427, 2007.
64. Mens, P.F. et al., Molecular diagnosis of malaria in the �eld: development of a novel 1-step nucleic acid lateral �ow immunoassay for the detection of all 4 human Plasmodium spp. and its evaluation in Mbita, Kenya. Diagn. Microbiol. Infect. Dis., 61, 421, 2008.
65. Safeukui, I. et al., Evaluation of FRET real-time PCR assay for rapid detection and differentiation of Plasmodium species in returning travellers and migrants. Malar. J., 7, 70, 2008.
66. Veron, V., Simon, S., and Carme, B., Multiplex real-time PCR detection of P. falciparum, P. vivax and P. malariae in human blood samples. Exp. Parasitol., 121, 346, 2009.
67. Bourgeois, N. et al., Comparison of three real-time PCR methods with blood smears and rapid diagnostic test in Plasmodium sp. infection. Clin. Microbiol. Infect., 16, 1305, 2010.
68. Cnops, L. et al., Giemsa-stained thick blood �lms as a source of DNA for Plasmodium species-speci�c real-time PCR. Malar. J., 9, 370, 2010.
69. Mixson-Hayden, T., Lucchi, N.W., and Udhayakumar, V., Evaluation of three PCR-based diagnostic assays for detecting mixed Plasmodium infection. BMC Res. Notes, 3, 88, 2010. 70. Ta, T.T. et al., First case of detection of Plasmodium knowlesi in Spain by real time PCR in a traveller from Southeast Asia. Malar. J., 9, 219, 2010. 71. Lal, A.A. et al., Ribosomal RNA-based diagnosis of Plasmodium falciparum malaria. Mol. Biochem. Parasitol., 36, 67, 1989. 72. Goman, M., Mons, B., and Scaife, J., The complete sequence of a Plasmodium malariae SSU rRNA gene and its comparison to other plasmodial SSU rRNA genes. Mol. Biochem. Parasitol., 45, 281, 1991. 73. Snounou, G. et al., Identi�cation of the four human malaria parasite species in �eld samples by the polymerase chain reaction and detection of a high prevalence of mixed infections. Mol. Biochem. Parasitol., 58, 283, 1993. 74. Waters, A.P., The ribosomal RNA genes of Plasmodium. Adv. Parasitol., 34, 33, 1994. 75. Tanomsing, N. et al., Accurate and sensitive detection of Plasmodium species in humans by use of the dihydrofolate reductase-thymidylate synthase linker region. J. Clin. Microbiol., 48, 3735, 2010. 76. Arai, M. et al., Detection of Plasmodium falciparum in human blood by a nested polymerase chain reaction. Am. J. Trop. Med. Hyg., 51, 617, 1994. 77. Patsoula, E. et al., A single-step, PCR-based method for the detection and differentiation of Plasmodium vivax and P. falciparum. Ann. Trop. Med. Parasitol., 97, 15, 2003. 78. Whiley, D.M. et al., Detection and differentiation of Plasmodium species by polymerase chain reaction and colorimetric detection in blood samples of patients with suspected malaria. Diagn. Microbiol. Infect. Dis., 49, 25, 2004. 79. Rosanas-Urgell, A. et al., Comparison of diagnostic methods for the detection and quanti�cation of the four sympatric Plasmodium species in �eld samples from Papua New Guinea. Malar. J., 9, 361, 2010. 80. Imwong, M. et al., Spurious ampli�cation of Plasmodium vivax small subunit RNA gene using the primers currently used to detect P. knowlesi. J. Clin. Microbiol., 47, 4173, 2009. 81. Cnops, L., Jacobs, J., and Van Esbroeck, M., Validation of a four-primer real-time PCR as a diagnostic tool for single and mixed Plasmodium infections. Clin. Microbiol. Infect., 17, 1101, 2011. 82. Genc, A., Eroglu, F., and Koltas, I.S., Detection of Plasmodium vivax by nested PCR and real-time PCR. Korean J. Parasitol., 48, 99, 2010. 83. Kimura, M. et al., Species-speci�c PCR detection of malaria parasites by microtiter plate hybridization: clinical study with malaria patients. J. Clin. Microbiol., 33, 2342, 1995. 84. de Monbrison, F. et al., Simultaneous identi�cation of the four human Plasmodium species and quanti�cation of Plasmodium DNA load in human blood by real-time polymerase chain reaction. Trans. R. Soc. Trop. Med. Hyg., 97, 387, 2003. 85. Rougemont, M. et al., Detection of four Plasmodium species in blood from humans by 18S rRNA gene subunit-based and species-speci�c real-time PCR assays. J. Clin. Microbiol., 42, 5636, 2004. 86. Mangold, K.A. et al., Real-time PCR for detection and identi�cation of Plasmodium spp. J. Clin. Microbiol., 43, 2435, 2005. 87. Babady, N.E. et al., Detection of Plasmodium knowlesi by real-time polymerase chain reaction. Am. J. Trop. Med. Hyg., 81, 516, 2009.
88. Shokoples, S.E. et al., Multiplexed real-time PCR assay for discrimination of Plasmodium species with improved sensitivity for mixed infections. J. Clin. Microbiol., 47, 975, 2009.
89. Divis, P.C. et al., A TaqMan real-time PCR assay for the detection and quantitation of Plasmodium knowlesi. Malar. J., 9, 344, 2010.
90. Dormond, L. et al., Multiplex real-time PCR for the diagnosis of malaria: Correlation with microscopy. Clin. Microbiol. Infect., 17, 469, 2011.
91. Witney, A.A. et al., Determining liver stage parasite burden by real time quantitative PCR as a method for evaluating pre-erythrocytic malaria vaccine ef�cacy. Mol. Biochem. Parasitol., 118, 233, 2001. 92. Nwakanma, D.C. et al., Quantitative detection of Plasmodium falciparum DNA in saliva, blood, and urine. J. Infect. Dis., 199, 1567, 2009. 93. Lucchi, N.W. et al., Real-time �uorescence loop mediated isothermal ampli�cation for the diagnosis of malaria. PLoS One, 5, e13733, 2010. 94. Taylor, S.M. et al., High-throughput pooling and real-time PCR-based strategy for malaria detection. J. Clin. Microbiol., 48, 512, 2010. 19 Chapter 19 - Sarcocystis
1. Levine, N.D. 1986. The taxonomy of Sarcocystis (Protozoa, Apicomplexa) species. J. Parasitol. 72, 372.
2. Frenkel, J.K., and Smith, D.D. 2003. Determination of the genera of cyst-forming coccidia. Parasitol. Res. 91, 384.
3. Rommel, M., and Heydorn, A.O. 1972. Contributions to the life cycle of Sarcosporidia. 3. Isospora hominis (Railliet and Lucet, 1891) Wenyon, 1923, the sporocyst of the Sarcosporidia of cattle and swine. Berl. Munch. Tierarztl. Wochenschr. 85, 143.
4. Heydorn, A.O. et al. 1975. Proposal for a new nomenclature of the Sarcosporidia. Z. Parasitenkd. 48, 73.
5. Levine, N.D. 1977. Nomenclature of Sarcocystis in the ox and sheep and of fecal Coccidia of the dog and cat. J. Parasitol. 63, 36.
6. Dubey, J.P. 1976. Review of Sarcocystis of domestic animals and of other Coccidia of cats and dogs. J. Am. Vet. Med. Assoc. 169, 1061.
7. Dubey, J.P., Speer, C.A., and Fayer, R. 1989. Sarcocystosis of Animals and Man, Chapters 1 and 10. CRC Press, Boca Raton, USA.
8. Dubey, J.P., and Lindsay, D.S. 2006. Neosporosis, toxoplasmosis, and sarcocystosis in ruminants. Vet. Clin. North Am. Food Anim. Pract. 22, 645.
9. Mehlhorn, H., and Heydorn, A.O. 1978. The Sarcosporidia (Protozoa, Sporozoa): Life cycle and �ne structure. Adv. Parasitol. 16, 43.
10. Dubey, J.P., and Fayer, R. 1983. Zoonosis in practice—sarcocystosis. Br. Vet. J. 139, 371.
11. Mehlhorn, H. 1988. Parasitology in Focus. Springer Verlag Berlin Heidelberg, Germany, 32.
12. Heydorn, A.O., Mehlhorn, H., and Gestrich, R. 1975. Light and electron microscope studies on cysts of Sarcocystis fusiformis in the muscles of calves infected experimentally with oocysts and sporocysts of Isospora hominis Railliet et Lucet, 1891. 2. Fine structure of metrocytes and merozoites. Zentralbl. Bakteriol. Orig. A. 232, 373.
13. Bottner, A., Charleston, W.A.G., and Hopcroft, D. 1987. The structure and identity of macroscopically visible Sarcocystis cysts in cattle. Vet. Parasitol. 24, 35.
14. Dubey, J.P., Fayer, R., and Speer, C.A. 1988. Experimental Sarcocystis hominis infection in cattle—Lesions and ultrastructure of sarcocysts. J. Parasitol. 74, 875.
15. Dubey, J.P., Speer, C.A., and Charleston, W.A.G. 1989. Ultrastructural differentiation between sarcocysts of Sarcocystis hirsuta and Sarcocystis hominis. Vet. Parasitol. 34, 153.
16. Odening, K. et al. 1995. On the morphological diagnostics and host speci�city of the Sarcocystis species of some domesticated and wild bovini (cattle, banteng and bison). Appl. Parasitol. 36, 161.
17. Heydorn, A.O. 1977. Sarkosporidien en�ziertes �eisch als mogliche krankheitsurache fur den menschen. Arch. Lebenmittelhyg. 28, 27. 18. Pena, H.F., de, J., Ogassawara, S., and Sinhorini, I.L. 2001. Occurrence of cattle Sarcocystis species in raw kibbe from Arabian food establishments in the city of São Paulo, Brazil, and experimental transmission to humans. J. Parasitol. 87, 1459. 19. Gauert, B., Jungmann, R., and Hiepe, T. 1983. Beziehungen zwischen intestinalstöringen und parasitären darminfektionen des menschen unter besonderer berücksichtigung von Sarcocystis-befall. Dt. Gesundh.–Wesen. 38, 62. 20. Tungtrongchitr, A. et al. 2007. The potential usefulness of the modi�ed Kato thick smear technique in the detection of intestinal sarcocystosis during �eld surveys. Southeast Asian J. Trop. Med. Public Health 38, 232. 21. Bottner, A. et al. 1987. The prevalence and identity of Sarcocystis in beef cattle in New Zealand. Vet. Parasitol. 24, 157. 22. van Knapen, F., Bouwman, D., and Greve, E. 1987. Study on the incidence of Sarcocystis spp in Dutch cattle using various methods. Tijdschr. Diergeneeskd. 112, 1095. 23. Vercruysse, J., Fransen, J., and Vangoubergen, M. 1989. The prevalence and identity of Sarcocystis cysts in cattle in Belgium. J. Vet. Med. B Infect. Dis. Vet. Public Health 36, 148. 24. Fortier, G. et al. 1993. Prevalence of muscular bovine sarcocystosis in the Calvados (Normandy-France). Rec. Med. Vet. Ec. Alfort. 169, 779. 25. Woldemeskel, M., and Gebreab, F. 1996. Prevalence of sarcocysts in livestock of Northwest Ethiopia. J. Vet. Med. B Infect. Dis. Vet. Public Health 43, 55. 26. Latif, B.M.A. et al. 1999. Prevalence of Sarcocystis spp in meat-producing animals in Iraq. Vet. Parasitol. 84, 85. 27. De Bosschere, H., and Ducatelle, R. 2001. Inverse correlation between myositis eosinophilica and number of Sarcocystis cystozoits in heart of cattle. Vlaams Diergen. Tijdschr. 70, 118. 28. Vangeel, L. et al. 2007. Molecular-based identi�cation of Sarcocystis hominis in Belgian minced beef. J. Food Prot. 70, 1523. 29. Pritt, B. et al. 2008. Detection of Sarcocystis parasites in retail beef: A regional survey combining histological and genetic detection methods. J. Food Prot. 71, 2144. 30. Hinaidy, H.K., and Supperer, R. 1979. Incidence of Sarcosporidia of pigs in Austria. Wien. Tierarztl. Monatsschr. 66, 281. 31. Damriyasa, I.M. et al. 2004. Cross-sectional survey in pig breeding farms in Hesse, Germany: Seroprevalence and risk factors of infections with Toxoplasma gondii, Sarcocystis spp and Neospora caninum in sows. Vet. Parasitol. 126, 271. 32. Omata, Y. et al. 1993. Survey of Sarcocystis spp infection in slaughtered pigs in East Hokkaido, Japan. J. Protozool. Res. 3, 29. 33. Dubey, J.P., and Powell, E.C. 1994. Prevalence of Sarcocystis in sows from Iowa. Vet. Parasitol. 52, 151. 34. Stephan, R., Tholen, R., and Meier, D. 1998. Greenish discolorations of beef muscle tissue and their assessment according to meat hygiene regulations. A case report. Tierarztl. Prax. Ausg. Grobtiere Nutztiere. 26, 31. 35. Saleque, A., Juyal, P.D., and Bhatia, B.B. 1990. Effect of temperature on the infectivity of Sarcocystis miescheriana cysts in pork. Vet. Parasitol. 36, 343. 36. Savini, G., Robertson, I.D., and Dunsmore, J.D. 1996. Viability of the sporocysts of Sarcocystis cruzi after exposure to different temperatures and relative humidities. Vet. Parasitol. 67, 153. 37. Dubey, J.P. et al. 2002. Effects of high temperature and disinfectants on the viability of Sarcocystis neurona sporocysts. J. Parasitol. 88, 1252.
38. Beaver, P.C., Gadgil, K., and Morera, P. 1979. Sarcocystis in man: A review and report of �ve cases. Am. J. Trop. Med. Hyg. 28, 819.
39. Wong, K.T., and Pathmanathan, R. 1992. High prevalence of human skeletal-muscle sarcocystosis in South-East Asia. Trans. R. Soc. Trop. Med. Hyg. 86, 631.
40. Mehrotra, R. et al. 1996. Diagnosis of human Sarcocystis infection from biopsies of the skeletal muscle. Pathology 28, 281. 41. Arness, M.K. et al. 1999. An outbreak of acute eosinophilic myositis attributed to human Sarcocystis parasitism. Am. J. Trop. Med. Hyg. 61, 548.
42. Fayer, R. 2004. Sarcocystis spp in human infections. Clin. Microbiol. Rev. 17, 894.
43. Pamphlett, R., and Odonoghue, P. 1990. Sarcocystis infection of human muscle. Aust. N. Z. J. Med. 20, 705.
44. Bunyaratvej, S., and Unpunyo, P. 1992. Combined Sarcocystis and gram-positive bacterial infections. A possible cause of segmental enterocolitis in Thailand. J. Med. Assoc. Thai. 75, 38.
45. Xiang, Z. et al. 2009. Non-invasive methods for identifying oocysts of Sarcocystis spp from de�nitive hosts. Parasitol. Int. 58, 293
46. Lele, V.R., Dhopavkar, P.V., and Kher, A. 1986. Sarcocystis infection in man (a case report). Indian J. Pathol. Microbiol. 29, 87.
47. Crum-Cian�one, N.F. 2008. Bacterial, fungal, parasitic, and viral myositis. Clin. Microbiol. Rev. 21, 473.
48. Velasquez, J.N. et al. 2008. Systemic sarcocystosis in a patient with acquired immune de�ciency syndrome. Hum. Pathol. 39, 1263.
49. Heckeroth, A.R., and Tenter, A.M. 2007. Sarcocystosis, Protozoal Abortion in Farm Ruminants: Guidelines for Diagnosis and Control. Oxfordshire, UK, 172.
50. Tadros, W., Laarman, J.J., and van den Eijk, A.A. 1974. The demonstration of antibodies to Sarcocystis fusiformis antigen in sera of Isospora hominis carriers, using the indirect �uorescence technique. Z. Parasitenkd. 43, 221.
51. Wong, K.T. et al. 1994. Light microscopic and threedimensional morphology of the human muscular sarcocyst. Parasitol. Res. 80, 138.
52. Tenter, A.M. 1995. Current Research on Sarcocystis species of domestic-animals. Int. J. Parasitol. 25, 1311.
53. Gajadhar, A.A., Marquardt, W.C., and Blair, C.D. 1992. Development of a model ribosomal RNA hybridization assay for the detection of Sarcocystis and other coccidia. Can. J. Vet. Res. 56, 208.
54. Zarlenga, D.S., and Higgins, J. 2001. PCR as a diagnostic and quantitative technique in veterinary parasitology. Vet. Parasitol. 101, 215.
55. MacPherson, J.M., and Gajadhar, A.A. 1993. Ribosomal RNA sequences for the speci�c detection of Toxoplasma gondii by hybridization assay. Mol. Cell. Probes 7, 97.
56. Heckeroth, A.R., and Tenter, A.M. 1999. Development and validation of species-speci�c nested PCRs for diagnosis of acute sarcocystiosis in sheep. Int. J. Parasitol. 29, 1331.
57. Macpherson, J.M., and Gajadhar, A.A. 1994. Speci�c ampli�cation of Sarcocystis cruzi DNA using a randomly primed polymerase chain reaction assay. Vet. Parasitol. 55, 267.
58. Guclu, F., Aldem, O.S., and Guler, L. 2004. Differential identi�cation of cattle Sarcocystis spp by random ampli�ed polymorphic DNA polymerase chain reaction (RAPD-PCR). Rev. Med. Vet. 155, 440. 59. Neefs, J.M. et al. 1991. Compilation of small ribosomal-subunit RNA sequences. Nucleic Acids Res. 19, 1987. 60. Tenter, A.M., and Johnson, A.M. 1997. Phylogeny of the tissue cyst-forming coccidia. Adv. Parasitol. 39, 69. 61. Fischer, S., and Odening, K. 1998. Characterization of bovine Sarcocystis species by analysis of their 18S ribosomal DNA sequences. J. Parasitol. 84, 50. 62. Prichard, R., and Tait, A. 2001. The role of molecular biology in veterinary parasitology. In: 18th International Conference of the 20 Chapter 20 - Toxoplasma
1. Schmidt, G.D., and Roberts, L.S. Foundations of Parasitology, 7th edn. McGraw-Hill, Boston, USA, 2005.
2. Nicolle, C., and Manceaux, L. Sur une infection à corps de Leishman (ou organisms voisins) du gondi. C. R. Seances Acad. Sci. 147, 763–766, 1908.
3. Dubey, J.P. The history and life cycle of Toxoplasma gondii. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 1–17, 2007
4. Petersen, E., and Liesenfeld, O. Clinical disease and diagnostics. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 81–100, 2007.
5. Buxton, D. et al. Toxoplasma gondii and ovine toxoplasmosis: New aspects of an old story. Vet. Parasitol. 149, 25–28, 2007.
6. Dubey, J.P., and Beattie, C.P. Toxoplasmosis of Animal and Man. CRC Press, Boca Raton, FL, USA, pp. 1–120, 1988.
7. Levine, N.D. Taxonomy of the sporozoa. J. Parasitol. 56 (sect. II, part 1), 208–209, 1963.
8. Jones, J.L., and Dubey, J.P. Waterborne toxoplasmosis— recent developments. Exp. Parasitol. 124, S10–S25, 2010.
9. Bogitsh, B.J., Carter, C.E., and Oeltmann, T.N. Human Parasitology, 3rd edn. Elsevier, Academic Press, London, UK, 2005.
10. Dubey, J.P., Lindsay, D.S., and Speer, C.A. Structures of Toxoplasma gondii tachyzoites, bradyzoites and sporozoites, and biology and development of tissue cysts. Clin. Microbiol. Rev. 11, 267–299, 1998.
11. Frenkel, J.K., Ruiz, A., and Chinchilla, M. Soil survival of Toxoplasma oocysts in Kansas and Costa Rica. Am. J. Trop. Med. Hyg. 24, 439–443, 1975.
12. Frenkel, J.K., and Dubey, J.P. Effects of freezing on the viability of Toxoplasma oocysts. J. Parasitol. 59, 587–588, 1973. 13. Dubey, J.P. Toxoplasma gondii survival under de�ned temperatures. J. Parasitol. 84, 862–865, 1998a.
14. Lindsay, D.S., Blagburn, B.L., and Dubey, J.P. Survival of nonsporulated Toxoplasma gondii oocysts under refrigerator conditions. Vet. Parasitol. 103, 309–313, 2002.
15. Lindsay, D.S. et al. Sporulation and survival of Toxoplasma gondii oocysts in seawater. J. Eukaryot. Microbiol. 50, S687–S688, 2003.
16. Ferguson, D.J.P., and Dubremetz, J.F. The ultrastructure of Toxoplasma gondii. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 19–48, 2007.
17. Ferguson, D.J., Hutchison, W.M., and Pettersen, E. Tissue cyst rupture in mice chronically infected with Toxoplasma gondii. An immunocytochemical and ultrastructural study. Parasitol. Res. 75, 599–603, 1989.
18. Weiss, L.M., and Kim, K. Bradyzoite development. In: Weiss, L.M., and Kim, K. (eds.), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 133–152, 2007.
19. Weiss, L.M., and Dubey, J.P. Toxoplasmosis: A history of clinical observations. Int. J. Parasitol. 39, S895–S901, 2009.
20. Luft, B.J., and Remington, J.S. Toxoplasmic encephalitis in AIDA. Clin. Infect. Dis. 15, 211–222, 1992.
21. Soete, M. et al. Toxoplasma gondii: Kinetics of bradyzoite–tachyzoite interconversion in vitro. Exp. Parasitol. 76, 259–264, 1993. 22. Odaert, H. et al. Stage conversion of Toxoplasma gondii in mouse brain during infection and immunodepression. Parasitol. Res. 82, 28–31, 1996. 23. Schaap, D. et al. Vaccination against toxoplasmosis: Current status and future prospects. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 721–752, 2007. 24. Morley, E.K. et al. Signi�cant familial differences in the frequency of abortion and Toxoplasma gondii infection within a �ock of Charollais sheep. Parasitology 131, 181– 185, 2005. 25. Duncanson, P. et al. High levels of congenital transmission of Toxoplasma gondii in a commercial sheep �ock. Int. J. Parasitol. 31, 1699–1703, 2001. 26. Frenkel, J.K., Dubey, J.P., and Miller, N.L. Toxoplasma gondii in cats: Fecal stages identi�ed as coccidian oocysts. Science 167, 893–896, 1970. 27. Dubey, J.P., and Frenkel, J.K. Cyst-induced toxoplasmosis in cats. J. Protozool. 19, 155–177, 1972. 28. Speer, C.A. et al. Time lapse video microscopy and ultrastructure of penetrating sporozoites, types 1 and 2 parasitophorous vacuoles, and the transformation of sporozoites to tachyzoites of the VEG strain of Toxoplasma gondii. J. Parasitol. 83, 565–574, 1997. 29. Speer, C.A., and Dubey, J.P. Ultrastructural differentiation of Toxoplasma gondii schizonts (types B to E) and gamonts in the intestines of cats fed bradyzoites. Int. J. Parasitol. 35, 193–206, 2004. 30. Dubey, J.P. History of discovery of the life cycle of Toxoplasma gondii. Int. J. Parasitol. 39, 877–882, 2009a. 31. Dubey, J.P. Oocyst shedding by cats fed isolated bradyzoites and comparison of bradyzoites of the VEG strain Toxoplasma gondii to cats and mice. J. Parasitol. 87, 215– 219, 2001. 32. Dubey, J.P. Duration of immunity to shedding of Toxoplasma gondii oocysts by cats. J. Parasitol. 81, 410–415, 1995. 33. Dubey, J.P. Comparative infectivity of oocysts and bradyzoites of Toxoplasma gondii for intermediate (mice) and de�nitive (cats) hosts. Vet. Parasitol. 140, 69–75, 2006. 34. OIE, 2009. Toxoplasmosis. In: Manual of Diagnostic Tests and Vaccines for Terrestrial Animals 2009. OIE, Terrestrial Manual 2008. Updated: August 14, 2009. http://www.oie.int/ eng/normes/mmanual/a_summry.htm. 35. Latkany, P. Ocular disease due to Toxoplasma gondii. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan—Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 101–131, 2007. 36. Dubey, J.P. Effect of freezing on the infectivity of Toxoplasma cysts to cats. J. Am. Vet. Med. Assoc. 165, 534–536, 1974. 37. Kotula, A.W. et al. Effect of freezing on infectivity of Toxoplasma gondii tissue cysts in pork. J. Food Prot. 54, 687–690, 1991. 38. Hill, D.E. et al. Effect of commonly used enhancement solutions on the viability of Toxoplasma gondii tissue cysts in pork loin. J. Food Protect. 67, 2230–2233, 2004. 39. Dubey, J.P. Toxoplasmosis in pigs—The last 20 years. Vet. Parasitol. 164, 89–103, 2009b. 40. Dubey, J.P. Re-examination of resistance of Toxoplasma gondii tachyzoites and bradyzoites to pepsin and trypsin digestion. Parasitology 116, 43–50, 1998b. 41. Bowie, W.R. et al. Outbreak of toxoplasmosis associated with municipial drinking water. The BC Toxoplasma Investigation Team. Lancet 350, 173–177, 1997. 42. de Moura, L. et al. Waterborne outbreak of toxoplasmosis, Brazil, from �eld to gene. Emerg. Infect. Dis. 12, 326–329, 2006.
43. Howe, D.K., Summers, B.C., and Sibley, L.D. Acute virulence in mice is associated with markers on chromosome VIII in Toxoplasma gondii. Infect. Immun. 64, 5193–5198, 1996.
44. Dardé, M.L., Ajzenberg, D., and Smith, J. Population structure and epidemiology of Toxoplasma gondii. In: Weiss, L.M., and Kim, K. (eds), Toxoplasma gondii. The Model Apicomplexan— Perspectives and Methods. Elsevier Ltd., Academic Press, London, UK, pp. 49–80, 2007.
45. Howe, D.K., and Sibley, L.D. Toxoplasma gondii comprises three clonal lineages: Correlation of parasite genotype with human disease. J. Infect. Dis. 172, 1561–1566, 1995.
46. Velmurugan, G.V., Dubey, J.P., and Su, C. Genotyping studies of Toxoplasma gondii isolates from Africa revealed that the archetypal clonal lineages predominate as in North America and Europe. Vet. Parasitol. 155, 314–318, 2008.
47. Ajzenberg, D. et al. Genetic diversity, clonality and sexuality in Toxoplasma gondii. Int. J. Parasitol. 34, 1185–1196, 2004.
48. Dubey, J.P., and Su, C. Population biology of Toxoplasma gondii: What’s out and where did they come from. Mem. Inst. Oswaldo Cruz. 104, 190–195, 2009.
49. Mikaelian, I. et al. Toxoplasmosis in beluga whales (Delphinapterus leucas) from the St. Lawrence estuary: Two case reports and a serological survey. J. Comp. Path. 122, 73–76, 2000.
50. Dubey, J.P. et al. Toxoplasma gondii, Neospora caninum, Sarcocystis neurona, and Sarcocystis canis-like infections in marine mammals. Vet. Parasitol. 116, 275–296, 2003.
51. Measures, L.N. et al. Seroprevalence of Toxoplasma gondii in Canadian pinnipeds. J. Wildl. Dis., 40, 294–300, 2004.
52. Prestrud, K.W. et al. Serosurvey for Toxoplasma gondii in arctic foxes and possible sources of infection in the high Arctic of Svalbard. Vet. Parasitol. 150, 6–12, 2007.
53. Jensen, S.-K. et al. The prevalence of Toxoplasma gondii in polar bears and their marine mammal prey; evidence for a marine transmission pathway? Polar Biol., 33(5), 599–606, 2009.
54. Rah, H. et al. Serosurvey of selected zoonotic agents in polar bears (Ursus maritimus). Vet. Rec. 156, 7–13, 2005.
55. Oksanen, A. et al. Prevalence of antibodies against Toxoplasma gondii in polar bears (Ursus maritimus) from Svalbard and East Greenland. J. Parasitol. 95, 89–94, 2009.
56. Sørensen, K.K. et al. Acute toxoplasmosis in three wild arctic foxes (Alopex lagopus) from Svalbard; one with co-infections of Salmonella Enteritidis PT1 and Yersinia pseudotuberculosis serotype 2b. Res. Vet. Sci. 78, 161–167, 2005.
57. McDonald, J.C. et al. An outbreak of toxoplasmosis in pregnant women in northern Quebec. J. Infect. Dis. 161, 769–774, 1990.
58. Messier, V. et al. Seroprevalence of Toxoplasma gondii among Nunavik Inuit (Canada). Zoonoses and Public Health 56, 188–197, 2009.
59. Johnson, C.K. et al. Prey choice and habitat use drive sea otter pathogen exposure in a resource-limited coastal system. PNAS 106, 2242–2247, 2009.
60. Vikøren, T. et al. Prevalence of Toxoplasma gondii antibodies in wild red deer, roe deer, moose, and reindeer from Norway. Vet. Parasitol. 120, 159–169, 2004.
61. Tenter, A.M., Heckeroth, A.R., and Weiss, L.M. Toxoplasma gondii: From animals to humans. Int. J. Parasitol. 30, 1217– 1258, 2000.
62. Dorny, P., Praet, N., Deckers, N., and Gabriel, S. Emerging food-borne parasites. Vet. Parasitol. 163, 196–206, 2009. 63. Dubey, J.P., and Jones, J.L. Toxoplasma gondii infection in humans and animals in the United States. Int. J. Parasitol. 38, 1257–1278, 2008. 64. Araujo, F.G. Depletion of L3T4 (CD4+) T lymphocytes prevents development of resistance to Toxoplasma gondii in mice. Infect. Immun. 59, 1614–1619, 1991. 65. Gazzinelli, R.T. et al. Synergistic role of CD4+ and CD8+ T lymphocytes in IFN-γ production and protective immunity induced by an attenuated Toxoplasma gondii vaccine. J. Immunol. 146, 286–292, 1991. 66. Mordue, D.G. et al. Acute toxoplasmosis leads to lethal overproduction of Th1 cytokines. J. Immunol. 167, 4574–4584, 2001. 67. Dunn, D. et al. Mother to child transmission of toxoplasmosis: Risk estimates for clinical counselling. Lancet 353, 1829– 1833, 1999. 68. Remington, J.S, McLead, R., and Thulliez, P. Toxoplasmosis. In: Remington, J.S., and Klein, J. (eds), Infectious Diseases of the Fetus and Newborn Infant. Philadelphia Publishing, WB Saunders, pp. 205–346, 2001. 69. Mitchell, C.D. et al. Congenital toxoplasmosis occurring in infants perinatally infected with human immunode�ciency virus 1. Pediatr. Infect. Dis. J. 9, 512–518, 1990. 70. Su, C. et al. Moving towards an integrated approach to molecular detection and identi�cation of Toxoplasma gondii. Parasitology 136, 1–11, 2009. 71. Sabin, A.B., and Feldman, H.A. Dyes as microchemical indicators of a new immunity phenomenon affecting a protozoon parasite (Toxoplasma). Science 108, 660–663, 1948. 72. Fulton, J.D. Studies on the agglutination of Toxoplasma gondii. Trans. R. Soc. Trop. Med. Hyg. 59, 694–704, 1965. 73. Dubey, J.P., Thulliez, P., and Powell, E.C. Toxoplasma gondii in Iowa sows: Comparison of antibody titers to isolation of T. gondii by bioassays in mice and cats. J. Parasitol. 81, 48–53, 1995. 74. Dubey, J.P. et al. Genetic and biologic characteristics of Toxoplasma gondii infections in free-range chickens from Austria. Vet. Parasitol. 133, 299–306, 2005. 75. Hill, D.E. et al. Comparison of detection methods for Toxoplasma gondii in naturally and experimentally infected swine. Vet. Parasitol. 141, 9–17, 2006. 76. Chatterton, J.M.W. et al. Toxoplasma gondii in vitro culture for experimentation. J. Microbiol. Methods 51, 331–335, 2002. 77. Bohne, W., Heesemann, J., and Gross, U. Reduced replication of Toxoplasma gondii is necessary for induction of bradyzoite-speci�c antigens: A possible role for nitric oxide in triggering stage conversion. Infect. Immun. 62, 1761–1767, 1994. 78. Burg, J.L. et al. Direct and sensitive detection of a pathogenic protozoan, Toxoplasma gondii, by polymerase chain reaction. J. Clin. Microbiol. 27, 1787–1792, 1989. 79. Savva, D. et al. Polymerase chain reaction for detection of Toxoplasma gondii. J. Med. Microbiol. 32, 25–31, 1990. 80. Ellis, J.T. Polymerase chain reaction approaches for the detection of Neospora caninum and Toxoplasma gondii. Int. J. Parasitol. 28, 1053–1060, 1998. 81. Su, C., and Dubey, J.P. Toxoplasma. In: Liu, D. (ed.), Molecular Detection of Foodborne Pathogens. CRC Press, Boca Raton, FL, USA, pp. 741–753, 2009. 82. Homan, W.L. et al. Identi�cation of a 200- to 300-fold repetitive 529 bp DNA fragmentin Toxoplasma gondii, and its use for diagnostic and quantitative PCR. Int. J. Parasitol. 30, 69–75, 2000. 83. Reischl, U. et al. Comparison of two DNA targets for the diagnosis of Toxoplasmosis by real-time PCR using �uorescence resonance energy transfer hybridization probes. BMC Infect. Dis. 3, 7, 2003.
84. Garcia, J.L. et al. Toxoplasma gondii: Detection by mouse bioassay, histopathology, and polymerase chain reaction in tissues from experimentally infected pigs. Exp. Parasitol. 113, 267–271, 2006.
85. Smith, G.C. et al. Prevalence of zoonotic important parasites in the red fox (Vulpes vulpes) in Great Britain. Vet. Parasitol. 118, 133–142, 2003.
86. Contini, C. et al. Evaluation of a real-time PCR-based assay using the lightcycler system for detection of Toxoplasma gondii bradyzoite genes in blood specimens from patients with toxoplasmic retinochoroiditis. Int. J. Parasitol. 35, 275–283, 2005.
87. Jauregui, L.H. et al. Development of a real-time PCR assay for detection of Toxoplasma gondii in pig and mouse tissues. J. Clin. Micriobiol. 39, 2065–2071, 2001.
88. Howe, D.K. et al. Determination of genotypes of Toxoplasma gondii strains isolated from patients with toxoplasmosis. J. Clin. Immunol. 35, 1411–1414, 1997.
89. Ajzenberg, D., Dumètre, A., and Dardé, M.-L. Multiplex PCR for typing strains of Toxoplasma gondii. J. Clin. Microbiol. 43, 1940–1943, 2005.
90. Khan, A. et al. Genotyping of Toxoplasma gondii strains from immunocompromised patients reveals high prevalence of Type I strains. J. Clin. Microbiol. 43, 5881–5887, 2005.
91. Su, C., Zhang, X., and Dubey, J.P. Genotyping of Toxoplasma gondii by multilocus PCR-RFLP markers: A high resolution and simple method for identi�cation of parasites. Int. J. Parasitol. 36, 841–848, 2006. 92. Dubey, J.P. et al. Prevalence of Toxoplasma gondii in dogs from Colombia, South America and genetic characterization of T. gondii isolates. Vet. Parasitol., 145, 45–50, 2007. 93. Sundar, N. et al. Genetic diversity among sea otter isolates of Toxoplasma gondii. Vet. Parasitol. 151, 125–132, 2008. 94. Prestrud, K.W. et al. Direct high-resolution genotyping of Toxoplasma gondii in arctic foxes (Vulpes lagopus) in the remote arctic Svalbard archipelago reveals widespread clonal Type II lineage. Vet. Parasitol. 158, 121–128, 2008. 95. Dubey, J.P. Re�nement of pepsin digestion method for isolation of Toxoplasma gondii from infected tissues. Vet. Parasitol. 81, 75–77 (1998c). 96. Kong, J.T. et al. Serotyping of Toxoplasma gondii infections in humans using synthetic peptides. J. Infect. Dis. 187, 1484–1495, 2003. 97. Nowakowska, D. et al. Genotyping of Toxoplasma gondii by multiplex PCR and peptide-based serological testing of samples from infants in Poland diagnosed with congenital toxoplasmosis. J. Clin. Microbiol. 44, 1382–1389, 2006. 98. Peyron, F. et al. Serotyping of Toxoplasma gondii in chronically infected pregnant women: Predominance of type II in Europe and types I and III in Colombia (South America). Microb. Infect. 8, 2333–2340, 2006.
Section III
Platyhelminthes: Cestoda 21 Chapter 21 - Diphyllobothrium
1. Chai, J.Y., Murrell, K.D., and Lymbery, A.J. Fish-borne parasitic zoonoses: Status and issues. Int. J. Parasitol. 35, 1233, 2005.
2. Dick, T.A. Diphyllobothriasis: The Diphyllobothrium latum human infection conundrum and reconciliation with a worldwide zoonosis. In: Murrell, K.D. and Fried, B. (eds), FoodBorne Parasitic Zoonoses: Fish and Plant-Borne Parasites. Springer, New York, pp. 151–84, 2007.
3. Muller, R. Worms and Human Disease, 2nd edn. CABI, Wallingford, UK, pp. 63–105, 2002.
4. WHO. Parasitic Zoonoses. Report of a WHO expert committee with the participation of FAO. WHO Techn. Rep. Ser. 637, 1, 1979.
5. Dupouy-Camet, J., and Peduzzi, R. Current situation of human diphyllobothriasis in Europe. Euro Surveill. 9, 5, 2004.
6. Scholz, T. et al. Update on the human broad tapeworm (genus Diphyllobothrium), including clinical relevance. Clin. Microbiol. Rev. 22, 146, 2009.
7. Semenas, L., Kreiter A., and Urbanski J. New cases of human diphyllobothriosis in Patagonia, Argentine. Rev. Saúde Pública 35, 214, 2001.
8. Sagua, H. et al. Nuevos casos de infección humana por Diphyllobothrium paci�cum (Nybelin, 1931) Margolis, 1956 en Chile y su probable relación con el fenómeno de El Nin´o, 1975–2000. Bol. Chil. Parasitol. 56, 22, 2001.
9. Sampaio, J.L. et al. Diphyllobothriasis, Brazil. Emerg. Infect. Dis. 11, 1598, 2005. 10. Brabec, J., Kuchta R., and Scholz T. Paraphyly of the Pseudophyllidea (Platyhelminthes: Cestoda): Circumscription of monophyletic clades based on phylogenetic analysis of ribosomal RNA. Int. J. Parasitol. 36, 1535, 2006. 11. Ashford, R.W., and Crewe, W. The Parasites of Homo Sapiens: An Annotated Checklist of the Protozoa, Helminths, and Arthropods for which We are Home. CRC Press, London, 2003. 12. Delyamure, S.L., Skryabin A.S., and Serdiukov A.M. Diphyllobothriata-�atworm parasites of man, mammals and birds. In: Principles of Cestodology. Vol. 9. Nauka, Moscow, Russia, 1985. [In Russian]. 13. Kamo, H. Guide to Identi�cation of Diphyllobothriid Cestodes. Tokyo, Japan, 1999. 14. Wicht B. et al. First record of human infection with the tapeworm Diphyllobothrium nihonkaiense in North America. Am. J. Trop. Med. Hyg. 78, 235, 2008. 15. Arizono, N. et al. Diplogonoporiasis in Japan: Genetic analyses of �ve clinical isolates, Parasitol. Int. 57, 212, 2008. 16. von Bonsdorff, B. Diphyllobothriasis in Man. Academic Press, London, 1977. 17. Andersen, K., and Halvorsen, O. Egg size and form as taxonomic criteria in Diphyllobothrium (Cestoda, Pseudophyllidea), Parasitology. 76, 229, 1978. 18. Andersen, K., and Gibson, D. A key to three species of larval Diphyllobothrium Cobbold, 1858 (Cestoda: Pseudophyllidea) occurring in European and North American freshwater �shes, Syst. Parasitol. 13, 3, 1989. 19. Dick, T.A., and Poole, B.C. Identi�cation of Diphyllobothrium dendriticum and Diphyllobothrium latum from freshwater �shes of central Canada. Can. J. Zool. 63, 196, 1985. 20. Garcia, L.S., and Bruckner, D.A. Diagnostic Medical Parasitology. American Society for Microbiology, Washington, DC, 1993. 21. Wicht, B., de Marval, F., and Peduzzi, R. Diphyllobothrium nihonkaiense (Yamane et al., 1986) in Switzerland: First molecular evidence and case reports. Parasitol. Int. 56, 195, 2007. 22. Yamane, Y. et al. Diphyllobothrium nihonkaiense sp. nov. (Cestoda: Diphyllobothriidae): Revised identi�cation of Japanese broad tapeworm. Shimane J. Med. Sci. 10, 29, 1986. 23. Fukumoto, S. et al. Distinction between Diphyllobothrium nihonkaiense and Diphyllobothrium latum by immunoelectrophoresis. Jpn. J. Parasitol. 37, 91, 1988. 24. Matsuura, T., Bylund, G., and Sugane, K. Comparison of restriction fragment length polymorphisms of ribosomal DNA between Diphyllobothrium nihonkaiense and D. latum. J. Helminthol. 66, 261, 1992. 25. De Vos, T., and Dick, T.A. Differentiation between Diphyllobothrium dendriticum and Diphyllobothrium latum using isoenzymes, restriction pro�les and ribosomal gene probes. Sys. Parasitol. 13, 161, 1989. 26. Mariaux, J. A molecular phylogeny of the eucestoda. J. Parasitol. 84, 114, 1998. 27. Olson, P.D., and Caira, J.N. Evolution of the major lineages of tapeworms (Platyhelminthes: Cestoda) inferred from 18S ribosomal DNA and elongation factor-1. J. Parasitol. 85, 1134, 1999. 28. Isobe, A. et al. Molecular phylogeny of Diphyllobothrium nihonkaiense (Yamane et al., 1986) and other diphyllobothiid tapeworms based on mitochondrial cytochrome c oxidase subunit I gene sequence. Parasitol. Int. 47 (suppl 1), 138, 1998.
29. Ando, K. et al. Five cases of Diphyllobothrium nihonkaiense infection with discovery of plerocercoids from an infective source, Oncorhynchus masou ishikawae. J. Parasitol. 87, 96, 2001.
30. Logan, F.J. et al. The phylogeny of diphyllobothriid tapeworms (Cestoda: Pseudophyllidea) based on ITS-2 rDNA sequences. Parasitol. Res. 94, 10, 2004.
31. Yera, H., Nicoulaud, J., and Dupouy-Camet, J. Use of nuclear and mitochondrial DNA PCR and sequencing for molecular identi�cation of Diphyllobothrium isolates potentially infective for humans. Parasite 15, 402, 2008.
32. Wicht, B. et al. Inter- and intra-speci�c characterization of tapeworms of the genus Diphyllobothrium (Cestoda: Diphyllobothriidea) from Switzerland, using nuclear and mitochondrial DNA targets. Parasitol. Int. 59, 35, 2010.
33. Hillis, D.M., and Dixon, M.T. Ribosomal DNA: Molecular evolution and phylogenetic inference. Q. Rev. Biol. 66, 411, 1991.
34. Skerikova, A. et al. Is the human-infecting Diphyllobothrium paci�cum a valid species or just a South American population of the holarctic �sh broad tapeworm, D. latum? Am. J. Trop. Med. Hyg. 75, 307, 2006.
35. Yera, H. et al. Putative Diphyllobothrium nihonkaiense acquired from a Paci�c salmon (Oncorhynchus keta) eaten in France: Genomic identi�cation and case report. Parasitol. Int. 55, 45, 2006.
36. Brown, W.M., George, Jr. M., and Wilson, A.C. Rapid evolution of animal mitochondria DNA. Proc. Natl Acad. Sci. USA 76, 1967, 1979.
37. Luton, K., Walker, D., and Blair, D. Comparisons of ribosomal internal transcribed spacers from two congeneric species of �ukes (Platyhelminthes: Trematoda: Digenea). Mol. Biochem. Parasitol. 56, 323, 1992.
38. Bowles, J., Blair, D., and McManus, D.P. Genetic variants within the genus Echinococcus identi�ed by mitochondrial DNA sequencing. Mol. Biochem. Parasitol. 54, 165, 1992. 39. Hall, T. BioEdit. 5.0. Distributed by the Author. Department of Microbiology, North Carolina State University, Raleigh, NC, 2001. 40. Nicoulaud, J., Yera, H., and Dupouy-Camet, J. Prevalence of Diphyllobothrium latum, L., 1758 infestation in Perca °uviatilis from Lake Leman.Parasite 12, 362, 2005 (in French). 41. Wicht, B. et al. Imported diphyllobothriasis in Switzerland: Molecular evidence of Diphyllobothrium dendriticum (Nitsch, 1824). Parasitol. Res. 102, 201, 2008. 42. Shimizu, H. et al. Diphyllobothriasis nihonkaiense: Possibly acquired in Switzerland from imported Paci�c salmon. Intern. Med. 47, 1359, 2008. 43. Paugam A. et al. Diphyllobothrium nihonkaiense infection: A new risk in relation with the consumption of salmon. Presse. Med. 38, 675, 2009. 44. Yamasaki, H., and Kuramochi, T. A case of Diphyllobothrium nihonkaiense infection possibly linked to salmon consumption in New Zealand. Parasitol. Res. 105, 583, 2009. 45. Torres, P. et al. Introduced and native �shes as infection foci of Diphyllobothrium spp. in humans and dogs from two localities at Lake Panguipulli in Southern Chile. Comp. Parasitol. 71, 111, 2004. 46. Nakao, M. et al. Mitochondrial genomes of the human broad tapeworms Diphyllobothrium latum and D. nihonkaiense (Cestoda: Diphyllobothriidae). Parasitol. Res. 101, 233, 2007. 47. Arizono, N. et al. Mitochondrial DNA divergence in populations of the tapeworm Diphyllobothrium nihonkaiense and its phylogenetic relationship with Diphyllobothrium klebanovskii. Parasitol. Intern. 58, 22, 2008. 48. Ruttenber, A.J. et al. Diphyllobothriasis associated with salmon consumption in Paci�c Coast states. Am. J. Trop. Med. Hyg. 33, 455, 1984. 49. Rausch, R.L., and Hilliard D.K. Studies on the helminth fauna of Alaska. XLIX. The occurrence of Diphyllobothrium latum (Linnaeus, 1758) (Cestoda: Diphyllobothriidae) in Alaska, with notes on other species. Can. J. Zool. 48, 1201, 1970. 22 Chapter 22 - Dipylidium
1. Nea�e, R.C., and Marty, A.M., Unusual infections in humans. Clin. Microbiol. Rev., 6, 34, 1993.
2. Turner, J.A., Human dipylidiasis (dog tapeworm infection) in the United States. J. Pediatr., 61, 763, 1962. 3. Shane, S.M. et al., A case of Dipylidium caninum in Baton Rouge, Louisiana. Int. J. Zoonoses, 13, 59, 1986. 4. Dai, R.S. et al., Severe infection of adult dogs with helminths in Hunan Province, China poses signi�cant public health concerns. Vet. Parasitol., 160, 348, 2009. 5. Currier, R.W. 2nd, Kinzer, G.M., and DeShields, E., Dipylidium caninum infection in a 14-month-old child. South Med. J., 66, 1060, 1973. 6. Reddy, S.B., Infestation of a �ve-month-old infant with Dipylidium caninum. Del. Med. J., 54, 455, 1982. 7. Hamrick, H.J. et al., Two cases of dipylidiasis (dog tapeworm infection) in children: update of an old problem. Pediatrics, 72, 114, 1983. 8. Wijesundera, M.D., and Ranaweera, R.L., Case reports of Dipylidium caninum, a pet associated infection. Ceylon Med. J., 34, 27, 1989. 9. Chappell, C.L., Enos, J.P., and Penn, H.M., Dipylidium caninum, an underrecognized infection in infants and children. Pediatr. Infect. Dis. J., 9, 745, 1990. 10. Raitiere, C.R., Dog tapeworm (Dipylidium caninum) infestation in a 6-month-old infant. J. Family Pract., 34, 101, 1992. 11. Reid, C.J., Perry, F.M., and Evans, N., Dipylidium caninum in an infant. Eur. J. Pediatr., 151, 502, 1992. 12. Ferraris, S. et al., Dipylidium caninum in an infant. Eur. J. Pediatr., 152, 702, 1993. 13. Gadre, D.V., Kumar, A., and Mathur, M., Infection by Dipylidium caninum through pet cats. Indian J. Pediatr., 60, 151, 1993. 14. Stiles, C.W., A case of infection with the double-pored tapeworm (Dipylidium caninum) in an American child. Am. Med., 5, 65, 1993. 15. Jackson, D. et al., Dipylidiasis in a 57-year-old woman. Med. J. Aust., 2, 740, 1977. 16. Samkari, A. et al., Dipylidium caninum mimicking recurrent enterobius vermicularis (pinworm) infection. Clin. Pediatr. (Phila), 47, 397, 2008. 17. Molina, C.P., Ogburn, J., and Adegboyega, P., Infection by Dipylidium caninum in an infant. Arch. Pathol. Lab. Med., 127, e157, 2003. 18. Ramana, K.V. et al., Human dipylidiasis: A case report of Dipylidium caninum infection in teaching hospital at Karimnagar. Online J. Health Allied Sci., 10, 28, 2011. 19. Wijesundera, M.D., The use of praziquantel in human infection with Dipylidium. Trans. R. Soc. Trop. Med. Hyg., 83, 383, 1989. 20. Jones, W.E., Niclosamide as a treatment for Hymenolepis diminuta and Dipylidium caninum infection in man. Am. J. Trop. Med. Hyg., 28, 300, 1979. 21. Lee, S.U., Chun, H.C., and Huh, S., Molecular phylogeny of parasitic Platyhelminthes based on sequences of partial 28S rDNA D1 and mitochondrial cytochrome c oxidase subunit I. Korean J. Parasitol., 45, 181, 2007. 22. Trachsel, D., Deplazes, P., and Mathis, A., Identi�cation of taeniid eggs in the faeces from carnivores based on multiplex PCR using targets in mitochondrial DNA. Parasitology, 134, 911, 2007. 23. Dyachenko, V. et al., Cost-effective method of DNA extraction from taeniid eggs. Parasitol. Res., 102, 811, 2008. 24. Bowles, J., and McManus, D.P., Genetic characterization of the Asian Taenia, a newly described taeniid cestode of humans. Am. J. Trop. Med. Hyg., 50, 33, 1994. 23 Chapter 23 - Echinococcus and Echinococcosis
1. Nelson, G.S., Pester, F.R., and Rickman, R. The signi�cance of wild animals in the transmission of cestodes of medical importance in Kenya. Trans. R. Soc. Trop. Med. Hyg. 59, 507, 1965.
2. Rausch, R.L., and D’Alessandro, A. The epidemiology of echinococcosis caused by Echinococcus oligarthrus and E. vogeli in the neotropics. In: Craig, P.S., and Pawlowski, Z. (eds), Cestode Zoonoses: Echinococcosis an Emergent and Global Problem. IOS Press, Amsterdam, The Netherlands, pp. 107–113, 2002.
3. Eckert, J., and Deplazes, P. Biological, epidemiological, and clinical aspects of echinococcosis, a zoonosis of increasing concern. Clin. Microbiol. Rev. 17, 107, 2004.
4. Eckert, J. et al. WHO/OIE Manual on Echinococcosis in Humans and Animals: A Public Health Problem of Global Concern. World Organisation for Animal Health, Paris, France, 265pp., 2001.
5. Ito, A., Nakao, M., and Sako, Y. Echinococcosis: Serological detection of patients and molecular identi�cation of patients and molecular identi�cation of parasites. Future Microbiol. 2, 439, 2007.
6. Macpherson, C.N.L. An active intermediate host role for man in the life cycle of Echinococcus granulosus in Turkana, Kenya. Am. J. Trop. Med. Hyg. 32, 397, 1983.
7. Thompson, R.C.A., and McManus, D.P. Aetiology: Parasites and life-cycles. In: WHO/OIE Manual on Echinococcosis in Humans and Animals: A Public Health Problem of Global Concern. World Organisation for Animal Health, Paris, France, p. 1, 2001. 8. Nakao, M. et al. A molecular phylogeny of the genus Echinococcus inferred from complete mitochondrial genomes. Parasitology 134, 713, 2007. 9. Thompson, R.C.A., and McManus, D.P. Towards a taxonomic revision of the genus Echinococcus. Trends Parasitol. 18, 452, 2002. 10. Hüttner, M. et al. Genetic characterization and phylogenetic position of Echinococcus felidis Ortlepp, 1937 (Cestoda: Taeniidae) from the African lion. Int. J. Parasitol. 38, 861, 2008. 11. Vogel, H. Üeber den Echinococcus multilocularis Süddeutschlands. I. Das Bandwurmstadium von Stämmen menschlicher und tierischer Herkunft. Z. Tropenmed. Parasitol. 8, 404, 1957. 12. Xiao, N. et al. Echinococcus shiquicus n. sp., a taeniid cestode from Tibetan fox and plateau pika in China. Int. J. Parasitol. 35, 693, 2005. 13. Lühe, M. Cystotänien südamerikanischer Feliden. Zoologische Jahrbücher, Abteitlung für allgemeine Zool. Physiol. Tiere 12 Suppl, 787, 1910. 14. Rausch, R.L., and Bernstein, J.J. Echinococcus vogeli sp. n. (Cestoda: Taeniidae) from the bush dog, Speothos venaticus (Lund). Z. Tropenmed. Parasitol. 23, 25, 1972. 15. Nakao, M. et al. State-of-the-art Echinococcus and Taenia: Phylogenetic taxonomy of human-pathogenic tapeworms and its application to molecular diagnosis. Infect. Genet. Evol. 10, 444, 2010. 16. Xiao, N. et al. Dual infection of animal hosts with different Echinococcus species in the eastern Qinghai-Tibet plateau region of China. Am. J. Trop. Med. Hyg. 75, 292, 2006. 17. Saarma, U. et al. A novel phylogeny for the genus Echinococcus, based on nuclear data, challenges relationships based on mitochondrial evidence. Parasitology 136, 317, 2009. 18. D’Allessandro, A., and Rausch, R.L. New aspects of neotropical polycystic (Echinococcus vogeli) and unicystic (Echinococcus oligarthrus) echinococcosis. Clin. Microbiol. Rev. 21, 380, 2008. 19. Hüttner, M. et al. A survey of Echinococcus species in wild carnivores and livestock in East Africa. Int. J. Parasitol. 39, 1269, 2009. 20. Nakao, M. et al. Geographic pattern of genetic variation in the fox tapeworm Echinococcus multilocularis. Parasitol. Int. 58, 384, 2009. 21. Ito, A. et al. Histopathological, serological and molecular con�rmation of alveolar echinococcosis cases in Mongolia. Am. J. Trop. Med. Hyg. 82, 266, 2010. 22. Rausch, R.L. Life cycle pattern and geographic distribution of Echinococcus species. In: Thompson, R.C.A. and Lymbery, A.J. (eds), Echinococcus and Hydatid Disease. CAB International, Oxon, UK, p. 89, 1995. 23. Romig, T., Dinkel, A., and Mackenstedt, U. The present situation of echinococcosis in Europe. Parasitol. Int. 55, S187, 2006. 24. Verster, A. Review of Echinococcus species in South Africa. Onderstepoort J. Vet. Res. 32, 7, 1965. 25. Nakao, M. et al. Genetic polymorphisms of Echinococcus tapeworms in China as determined by mitochondrial and nuclear DNA sequences. Int. J. Parasitol. 40, 379, 2010. 26. Johnson, K.S. et al. Vaccination against ovine cysticercosis using a de�ned recombinant antigen. Nature 338, 585, 1989. 27. Lightowlers, M.W. et al. Molecular and genetic characterization of the host-protective oncosphere antigens of taeniid cestode parasites. Int. J. Parasitol. 33, 1207, 2003.
28. Lightowlers, M.W. Cestode vaccines: Origins, current status and future prospects. Parasitology 133, S27, 2006.
29. Jabbar, A. et al. The ultrastructure of taeniid cestode oncospheres and localization of host-protective antigens. Parasitology 137, 521, 2010.
30. Thompson, R.C.A. Biology and systematic of Echinococcus. In: Thompson, R.C.A., and Lymbery, A.J. (eds), Echinococcus and Hydatid Disease, Chapter 1. CAB International, Oxon, p. 1, 1995.
31. Nakaya, K. et al. Usefulness of severe combined immunode�ciency (scid) and inbred mice for studies of cysticercosis and echinococcosis. Parasitol. Int. 56, S91, 2006.
32. Rausch, R.L., D’Alessandro, D., and Rausch, V.R. Characteristics of the larval Echinococcus vogeli Rausch and Bernstein, 1972 in the natural intermediate host, the paca, Cuniculus paca L. (Rodentia: Dasyproctidae). Am. J. Trop. Med. Hyg. 30, 1043, 1981.
33. Nakao, M., Sako, Y., and Ito, A. Isolation of polymorphic microsatellite loci from the tapeworm Echinococcus multilocularis. Infect. Genet. Evol. 3, 159, 2003.
34. Badaraco, J.L. et al. Using mitochondrial and nuclear markers to evaluate the degree of genetic cohesion among Echinococcus populations. Exp. Parasitol., 119, 453, 2008.
35. Okamoto, M. et al. Evidence of hybridization between Taenia saginata and Taenia asiatica. Parasitol. Int. 59, 70, 2010.
36. Craig, P.S. et al. A large focus of alveolar echinococcosis in central China. Lancet 340, 826, 1992.
37. Craig, P.S. et al. Human echinococcosis: A neglected disease? Trop. Med. Health 35, 283, 2007.
38. Schantz, P.M. et al. Epidemiology and control of hydatid disease. In: Thompson, R.C.A., and Lymbery, A.J. (eds), Echinococcus and Hydatid Disease. CAB International, Oxon, 233pp., 1995.
39. Gemmell, M.A. et al. Epidemiology. In: WHO/OIE Manual on Echinococcosis in Humans and Animals:A Public Health Problem of Global Concern. World Organisation for Animal Health, Paris, France, 143pp., 2001.
40. Craig, P.S., and McManus, D.P. Echinococcosis: Transmission biology and epidemiology. Parasitology 127, S1, 2003.
41. Gurbadam, A. et al. Mongolian and Japanese joint conference on “Echinococcosis: Diagnosis, treatment and prevention in Mongolia” June 4, 2009. Parasit. Vectors 3, 8, 2010.
42. Li, T. et al. Echinococcosis in Tibetan populations, western Sichuan province, China. Emerg. Infect. Dis. 11, 1866, 2005.
43. Schantz, P.M. et al. Echinococcosis on the Tibetan plateau: Prevalence and risk factors for cystic and alveolar echinococcosis in Tibetan populations in Qinghai province, China. Parasitology 127, S109, 2003.
44. Ito, A. et al. Control of echinococcosis and cysticercosis: A public health challenge to international cooperation in China. Acta Trop., 86, 3, 2003.
45. Tang, C. et al. Alveolar Echinococcus species from Vulpes corsac in Hulunbeier, Inner Mongolia, China, and differential development of metacestodes in experimental rodents. J. Parasitol. 92, 719, 2006.
46. Tang, C. et al. Studies on the alveolar Echinococcus species in northward Daxingan mountains, Inner Mongolia, China. III. Echinococcus russicensis sp. nov. Chin. J. Zoonoses 23, 957, 2007.
47. Bretagne, S. et al. Echinococcus multilocularis: Microsatellite polymorphism in U1 snRNA genes. Exp. Parasitol. 82, 324, 1996.
48. Bart, J.M. et al. Genotyping of human cystic echinococcosis in Xinjiang, PR China. Parasitology 133, 571, 2006. 49. Bart, J.M. et al. EmsB, a tandem repeated multi-loci microsatellite, new tool to investigate the genetic diversity of Echinococcus multilocularis. Infect. Genet. Evol. 6, 390, 2006. 50. Knapp, J. et al. Assessment of use of microsatellite polymorphism analysis for improving spatial distribution tracking of Echinococcus multilocularis. J. Clin. Microbiol. 45, 2943, 2007. 51. Knapp, J. et al. Genetic diversity of Echinococcus multilocularis on a local scale. Infect. Genet. Evol. 8, 367, 2008. 52. Knapp, J. et al. Genetic diversity of the cestode Echinococcus multilocularis in red foxes at a continental scale in Europe. PLoS Negl. Trop. Dis. 3, e452, 2009. 53. Knapp, J. et al. The genomic Echinococcus microsatellite EmsB sequences: From a molecular marker to the epidemiological tool. Parasitology 137, 439, 2010. 54. Casulli, A. et al. Multi-locus microsatellite analysis supports the hypothesis of an autochthonous focus of Echinococcus multilocularis in northern Italy. Int. J. Parasitol. 39, 837, 2009. 55. Knapp, J. et al. Echinococcus vogeli infection in a hunter, French Guiana. Emerg. Infect. Dis. 15, 2029, 2009. 56. Brunetti, E. et al. Expert consensus for the diagnosis and treatment of cystic and alveolar echinococcosis in humans. Acta Trop. 114, 1, 2010. 57. Li, T. et al. Speci�c IgG responses to recombinant antigen B and Em18 in cystic and alveolar echinococcosis in China. Clin. Vaccine Immunol. 17, 470, 2010. 58. Li, T. et al. Widespread co-endemicity of human cystic and alveolar echinococcosis on the eastern Tibetan plateau, northwest Sichuan/southeast Qinghai, China. Acta Trop. 113, 248, 2010. 59. Scharf, G. et al. Radiographic, ultrasonographic, and computed tomographic appearance of alveolar echinococcosis in dogs. Vet. Radiol. Ultrasound 45, 411, 2004. 60. Staebler, S. et al. Serological diagnosis of canine alveolar echinococcosis. Vet. Parasitol. 141, 243, 2006. 61. Lightowlers, M.W., and Gottstein, B. Echinococcosis/ hydatidosis: Antigens, immunological and molecular diagnosis. In: Thompson, R.C.A., and Lymbery, A.J. (eds), Echinococcus and Hydatid Disease. CAB International, Oxon, UK, 355pp., 1995. 62. Ito, A. Serologic and molecular diagnosis of zoonotic larval cestode infections. Parasitol. Int. 51, 221, 2002. 63. Ito, A., and Craig, P.S. Immunodiagnostic and molecular approaches for the detection of taeniid cestode infections. Trends Parasitol. 19, 377, 2003. 64. Ito, A. et al. Differential serodiagnosis for cystic and alveolar echinococcosis using fractions of Echinococcus granulosus cyst �uid (antigen B) and E. multilocularis protosoclex (Em18). Am. J. Trop. Med. Hyg. 60, 188, 1999. 65. Ito, A. et al. Evaluation of an enzyme-linked immunosorbent assay (ELISA) with af�nity-puri�ed Em18 and an ELISA with recombinant Em18 for differential diagnosis of alveolar echinococcosis: Results of a blind test. J. Clin. Microbiol. 40, 4161, 2002. 66. Sako, Y. et al. Alveolar echinococcosis: Characterization of diagnostic antigen Em18 and serological evaluation of recombinant Em18. J. Clin. Microbiol. 40, 2760, 2002. 67. Xiao, N. et al. Evaluation of use of recombinant Em18 and af�nity-puri�ed Em18 for serological differentiation of alveolar echinococcosis from cystic echinococcosis and other parasitic infections. J. Clin. Microbiol. 41, 3351, 2003.
68. Ito, A. et al. Novel antigens for neurocysticercosis: Simple method for preparation and evaluation for serodiagnosis. Am. J. Trop. Med. Hyg. 59, 291, 1998.
69. Sako, Y. et al. Molecular characterization ad diagnostic value of Taenia solium low-molecular-weight antigen genes. J. Clin. Microbiol. 38, 4439, 2000.
70. Graichen, D.A.S. et al. Expression and diversity of Echinococcus multilocularis AgB genes in secondary infected mice: Evaluating the in�uence of T-cell immune selection on antigenic variation. Gene. 392, 98, 2007.
71. Riganò, R. et al. Echinococcus granulosus antigen B impairs human dendritic cell differentiation and polarizes immature dendritic cell maturation towards a Th2 cell response. Infect. Immun. 75, 1667, 2007.
72. Siracusano, A. et al. Immunomodulatory mechanisms during Echinococcus granulosus infection. Exp. Parasitol. 119, 483, 2008.
73. Siracusano, A. et al. Molecular cross-talk in host-parasite relationships: The intriguing immunomodulatory role of Echinococcus antigen B in cystic echinococcosis. Int. J. Parasitol. 38, 1371, 2008.
74. Siles-Lucas, M., and Gottstein, B. The 14-3-3 protein: A key molecule also in parasites. Trends Parasitol. 19, 575, 2003.
75. Brehm, K. et al. The molecular mechanisms of larval cestode development: First steps into an unknown world. Parasitol. Int. 55, S15, 2006.
76. Brehm, K. Echinococcus multilocularis as an experimental model in stem cell research and molecular host-parasite interaction. Parasitology 237, 537, 2010.
77. Gottstein, B. et al. Molecular survival strategies of Echinococcus multilocularis in the murine host. Parasitol. Int. 55, S45, 2006.
78. Haag, K.L. et al. The EG95 antigen of Echinococcus spp. contains positively selected amino acids, which may in�uence host speci�city and vaccine ef�cacy. PLoS One 4, e5362, 2009.
79. Mejri, N, Hemphill, A., and Gottstein, B. Triggering and modulation of the host-parasite interplay by Echinococcus multilocularis: A review. Parasitology 137, 557, 2010.
80. Oriol, R. et al. Puri�cation of lipoprotein antigens of Echinococcus granulosus from sheep hydatid �uid. Am. J. Trop. Med. Hyg. 20, 569, 1971.
81. Lightowlers, M.W. et al. Subunit composition and speci�city of major cyst �uid antigens of Echinococcus granulosus. Mol. Biochem. Parasitol. 37, 171, 1989.
82. Siracusano, A. et al. Detection of antibodies against Echinococcus granulosus major antigens and their subunits by immunoblotting. Trans. R. Soc. Trop. Med. Hyg. 85, 239, 1991.
83. Ioppolo, S. et al. Immunological responses to antigen B from Echinococcus granulosus cyst �uid in hydatid patients. Parasitol. Immunol. 18, 571, 1996.
84. Ito, A. et al. Evaluation of Em18-, Em16-, antigen B-western blots, Em2 plus -ELISA and four other tests for differential serodiagnosis of alveolar and cystic echinococcosis patients in Poland. Parasitol. Int. 47, 95, 1998.
85. Mamuti, W. et al. Usefulness of hydatid cyst �uid of Echinococcus granulosus developed in mice with secondary infection for serodiagnosis of cystic echinococcosis in humans. Clin. Diagn. Lab. Immunol. 9, 573, 2002.
86. Leggatt, G.R., Yang, W., and McManus, D.P. Serological evaluation of the 12 kDa subunit of antigen B in Echinococcus granulosus cyst �uid by immunoblot analysis. Trans. R. Soc. Trop. Med. Hyg. 86, 1, 1992. 87. Frosch, P. et al. Sequence heterogeneity of the echinococcal antigen B. Mol. Biochem. Parasitol. 64, 171, 1994. 88. Fernández, V. et al. Molecular characterization of a novel 8 kDa subunit of Echinococcus granulosus antigen B. Mol. Biochem. Parasitol. 77, 247, 1996. 89. Chamale, G. et al. Echinococcus granulosus antigen B is encoded by a gene family. Mol. Biochem. Parasitol. 116, 233, 2001. 90. Arend, A.C. et al. The Echinococcus granulosus antigen B shows a high degree of genetic variability. Exp. Parasitol. 108, 76, 2004. 91. Mamuti, W. et al. Echinococcus multilocularis: Developmental stage-speci�c expression of antigen B 8-kDa-subunits. Exp. Parasitol. 113, 75, 2006. 92. Tappe, D. et al. Close relationship between clinical regression and speci�c serology in the follow-up of patients with alveolar echinococcosis in different clinical stages. Am. J. Trop. Med. Hyg. 80, 792, 2009. 93. Bresson-Hadni, S. et al. Should possible recurrence forbid liver transplantation in patients with end-stage alveolar echinococcosis? A 20-yr follow up. Liver Transpl. 17, 855, 2011. 94. Tappe, D. et al. Immunoglobulin G subclass responses to recombinant Em18 in the follow-up of patients with alveolar echinococcosis in different clinical stages. Clin. Vaccine Immunol. 17, 944, 2010. 95. Mamuti, W. et al. Molecular cloning, expression, and serological evaluation of an 8-kilodalton subunit of antigen B from Echinococcus multilocularis. J. Clin. Microbiol. 42, 1082, 2004. 96. Müller, N. et al. Improved serodiagnosis of alveolar echinococcosis of humans using an in vitro-produced Echinococcus multilocularis antigen. Parasitology 134, 879, 2007. 97. Fujimoto, N. et al. Usefulness of recombinant Em18-ELISA to evaluate ef�cacy of treatment in patients with alveolar echinococcosis. J. Gastroenterol. 40, 426, 2005. 98. Ishikawa, Y. et al. Serological monitoring of progression of alveolar echinococcosis with multi-organ involvement using recombinant Em18. J. Clin. Microbiol. 47, 3191, 2009. 99. Wen, H. et al. Immunoblot evaluation of IgG and IgG subclass antibody responses for immunodiagnosis of human alveolar hydatidosis. Trans. R. Soc. Trop. Med. Hyg. 89, 485, 1995. 100. Gottstein, B., D’Alessandro, A., and Rausch, R.L. Immunodiagnosis of polycystic hydatid disease/polycystic echinococcosis due to Echinococcus vogeli. Am. J. Trop. Med. Hyg. 53, 558, 1995. 101. Bart, J.M. et al. Comparison of several commercial serologic kits and Em18 serology for detection of human alveolar echinococcosis. Diagn. Microbiol. Infect. Dis. 59, 93, 2007. 102. Brehm, K. et al. Characterization of the genomic locus expressing the ERM-like protein of Echinococcus multilocularis. Mol. Biochem. Parasitol. 100, 147, 1999. 103. Sako, Y. et al. Development of an immunochromatographic test to detect antibodies against recombinant Em18 for diagnosis of alveolar echinococcosis. J. Clin. Microbiol. 47, 252, 2009. 104. Liance, M. et al. Immunodiagnosis of Echinococcus infections: Con�rmatory testing and species differentiation by a new commercial Western blot. J. Clin. Microbiol. 38, 3718, 2000. 105. Ito, A., Schantz, P.M., and Wilson, J.F. Em18, a new serodiagnostic marker for differentiation of active and inactive cases of alveolar hydatid disease. Am. J. Trop. Med. Hyg. 52, 41, 1995.
106. Ito, A. et al. Immunodiagnosis of alveolar echinococcosis by enzyme-linked immunosorbent assay using a partially puri�ed Em18/16 enriched fraction. Clin. Diagn. Lab. Immunol. 4, 57, 1997.
107. Ito, A. et al. Antibody responses against Em18 and Em16 serological markers in alveolar and cystic echinococcosis patients from northwest China. Jpn. J. Med. Sci. Biol. 50, 19, 1997.
108. Ito, A. et al. An imported case of cystic echinococcosis in Japan diagnosed by imaging and serology with con�rmation of Echinococcus granulosus-speci�c DNA sequences. Am. J. Trop. Med. Hyg. 58, 790, 1998.
109. Kern, P. et al. WHO classi�cation of alveolar echinococcosis: Principles and application. Parasitol. Int. 55, S283, 2006.
110. Bresson-Hadni, S. et al. Imaging and non-surgical interventional treatment in human alveolar echinococcosis. Parasitol. Int. 55, S267, 2006.
111. Ishikawa, Y. et al. In vivo evaluation of the viability of human hepatic alveolar echinococcal tissue from patients. Clin. Parasitol. 1, 157, 1990. [In Japanese].
112. Bowles, J., and McManus, D.P. Rapid discrimination of Echinococcus species and strains using a polymerase chain reaction-based RFLP method. Mol. Biochem. Parasitol. 57, 231, 1993.
113. Gasser, R.B. and Chilton, N.B. Characterisation of taeniid cestode species by PCR-RFLP of ITS2 ribosomal DNA. Acta Trop. 59, 31, 1995.
114. Zhang, L. et al. Screening for different genotypes of Echinococcus granulosus within China and Argentina by single-strand conformation polymorphism (SSCP) analysis. Trans. R. Soc. Trop. Med. Hyg. 93, 329, 1999.
115. Bartholomei-Santos, M.L. et al. Isolation and characterization of microsatellites from the tapeworm Echinococcus granulosus. Parasitology 126, 599, 2003.
116. Le, T.H. et al. Complete mitochondrial genomes con�rm the distinctiveness of the horse-dog and sheep-dog strains of Echinococcus granulosus. Parasitology 124, 97, 2002.
117. Nakao, M. et al. The complete mitochondrial DNA sequences of the cestode Echinococcus multilocularis (Cyclophyllidea; Taeniidae). Mitochondrion 1, 497, 2002.
118. Yamasaki, H. et al. Mitochondrial DNA diagnosis for cestode zoonoses: Application to formalin-�xed paraf�n-embedded tissue specimens. Southeast Asian J. Trop. Med. Public Health 38(Suppl. 1), 166, 2007. 119. Yamasaki, H. et al. Genetic analysis of Echinococcus multilocularis originating from a patient with alveolar echinococcosis occurring in Minnesota in 1977. Am. J. Trop. Med. Hyg. 79, 245, 2008. 120. Bowles, J. et al. Genetic variants within the genus Echinococcus identi�ed by mitochondrial DNA sequencing. Mol. Biochem. Parasitol. 54, 165, 1992. 121. Nakao, M. et al. Mitochondrial genetic code in cestodes. Mol. Biochem. Parasitol. 111, 415, 2000. 122. Bowles, J., and McManus, D.P. NADH dehydrogenase 1 gene sequences compared for species and strains of the genus Echinococcus. Int. J. Parasitol. 23, 969, 1993. 123. Moro, P.L. et al. Molecular identi�cation of Echinococcus isolates from Peru. Parasitol. Int. 58, 184, 2009. 124. Bowles, J., Blair, D., and McManus, D.P. Molecular genetic characterization of the cervid strain (“northern form”) of Echinococcus granulosus. Parasitology 109, 215, 1994. 125. Kamenetzky, L. et al. Several strains of Echinococcus granulosus infect livestock and humans in Argentina. Infect. Genet. Evol. 2, 129, 2002. 126. Lavikainen, A. et al. Molecular genetic characterization of Fennoscandian cerrvid strain, a new genotypic group (G10) of Echinococcus granulosus. Parasitology 127, 207, 2003. 127. Moks, E. et al. First report of Echinococcus granulosus G8 in Eurasia and a reappraisal of the phylogenetic relationships of ‘genotypes’ G5-G10. Parasitology 135, 647, 2008. 128. Nkouawa, A. et al. Loop-mediated isothermal ampli�cation method for differentiation and rapid detection of Taenia species. J. Clin. Microbiol. 47, 168, 2009. 129. Nkouawa, A. et al. Evaluation of loop-mediated isothermal ampli�cation method using fecal specimens for differential detection of Taenia species. J. Clin. Microbiol. 48, 3350, 2010. 130.Yanagida, T. et al. Neurocysticercosis: Assessing where the infection was acquired from. J. Travel Med. 17, 206, 2010. 131. Jongwietiwes, U. et al. Isolated intradural-extramedullary spinal cysticercosis: A case report. J. Travel Med. 18, 284, 2011. 132. Lavikainen, A. et al. Molecular characterization of Echinococcus isolates of cervid origin from Finland and Sweden. Parasitology 133, 565, 2006. 133. Lavikainen, A. et al. Mitochondrial DNA data reveal cryptic species within Taenia krabbei. Parasitol. Int. 59, 290, 2010. 24 Chapter 24 - Hymenolepis
1. Coombs, I. Helminthic species recovered from humans. In: Crompton, D.W.T. and Savioli, L. (eds), Handbook of Helminthiasis for Public Health, CRC Press, Taylor & Francis Group, Boca Raton, FL, pp. 12–24, 2006. 2. Wittner, M., and Tanowitz, H.B. Other cestode infections. In: Guerrant, R.L., Walker, D.H., Weller, P.F. (eds.), Tropical Infectious Diseases: Principles, Pathogens, and Practice. 1st edn., Churchill Livingston, New York, pp. 1026–1027, 1999. 3. Ghazal, A.M., and Avery, R.A. Population dynamics of Hymenolepis nana in mice: Fecundity and the crowding effect. Parasitology. 69, 403, 1974. 4. Schantz, P.M. Tapeworms (cestodiasis). Gastroenterol. Clin. North Am. 25, 637, 1996. 5. Ito, A. Basic and applied immunology in cestode infections from Hymenolepis to Taenia and Echinococcus. Int. J. Parasitol. 27, 1203, 1997. 6. Andreasssen, J. Immunity to adult cestodes: Basic knowledge and vaccination problems: A review. Parasitologia 33, 45, 1991. 7. Ito, A., Nakao, M., and Wandra, T. Human teaniasis and cysticercosis in Asia. Lancet 362, 1918, 2003. 8. Mehraj, V. et al. Prevalence and factors associated with intestinal parasitic infection among children in an urban slum of Karachi. PLoS One 3, e3680, 2008. 9. Richards, F.O. Jr., Diphyllobothrium, Dipylidium, and Hymenolepis species. In: Long, S.S., Pickering, L.K., and Prober, C.G. (eds.), Principles and Practice of Pediatric Infectious Diseases, Churchill Livingstone, New York, p. 1351, 2003. 10. Beaver, P.C., Jung, R.C., and Cupp, E.W. (eds), In: Clinical Parasitology, 9th ed., Lea & Febiger, Philadelphia, PA, pp. v365–367, 1984. 11. Bogitsh, B.J., Carter, C.E., and Oeltmann, T.N. (eds), In: Human Parasitology, 3rd ed., Elsevier Academic Press, London, pp. 274–278, 2005. 12. Andreassen, J., and Birklund, D. In vivo effects of immune mouse serum on three species of Hymenolepis tapeworms. Bull. Scand. Soc. Parasitol. 3, 26, 1993. 13. Hopkins, C.A., and Allen, L.M. Hymenolepis diminuta: The role of tail in determining the position of the worm in the intestine of the rat. Parasitology 79, 401, 1979. 14. Voge, M., and Heyneman, D., Effect of high temperature on the larval development of Hymenolepis nana and Hymenolepis diminuta (Cestoda: Cyclophyllidea). J. Parasitol. 44, 249, 1958. 15. Katz, M., Despommier, D.D., and Gwadz, R.W., Tapeworms of minor medical importance. Parasit. Dis. 16, 88, 1989. 16. Chu, G.S., Palmieri, J.R., and Sullivan, J.T. Beettle eating: A Malaysian folk medical practice and its public health implication. Trop. Geog. Med. 29, 422, 1977. 17. Crompton, D.W.T. How much human helminthiasis is there in the world? J. Parasitol. 85, 397, 1999. 18. Conchedda, M. et al. Immune response to the cestode Hymenolepis nana: Cytokine production during infection with eggs or cysts. Int. J. Parasitol. 27, 32, 1997. 19. Mirdha, B.R., and Samantaray, J.C. Hymenolepis nana: A common cause of paediatric diarrhoea in urban slum dwellers in India. J. Trop. Pediatr. 48, 331, 2002. 20. Lucas, S.B. et al. Abnormal development of Hymenolepis nana larvae in immunocompromised mice. J. Helminthol. 54, 75, 1980. 21. Sirivichayakul, C. et al. Hymenolepis nana infection in Thai children. J. Med Assoc. Thai. 83, 1035, 2000. 22. Dwinell, M.B., Bass P., and Oaks, J.A. Hymenolepis diminuta fractions but not previous tapeworm infection stimulate intestinal myoelectric alteration in vivo in the rat. J. Parasitol. 84, 673, 1998.
23. Mohammed, M.A., and Hepazi, M.A. Intestinal permeability in Hymenolepis nana as re�ected by non-invasive lactulose/ mannitol dual permeability test and its impact on nutritional parameters of patients. J. Egypt. Soc. Parasitol. 37, 877, 2007.
24. Lloyd, S. Other cestode infections: Hymenolepiosis, diphyllobothriosis, coenurosis, and other adult and larval cestodes. In: Palmer, S.R., Soulsby, L., Simpson, D.I.H. (eds.), Zoonoses, Oxford Medical Press, Oxford, London, UK, pp. 651–663, 1998.
25. Pawlowski, P.W. Cestodes: Teaniasis, dyphyllobothriasis, hymenolepiasis, and others. In: Warren, K.S., and Mahmoud, A.A.F. (eds.), Tropical and Geographical Medicine, McGrawHill Book Company, New York, pp. 471–486, 1984.
26. Dworkin, M.S., and Wan, P.C. Helminth infection in HIVinfected persons in the United States, 1990–1999. Clin. Infect. Dis. 30, 241, 2000.
27. Weiis, M.E., and Raszaka, W.V. Jr., Cestode infection in children. Adv. Pediatr. Infect. Dis. 12, 109, 1996.
28. Manson, P.R., and Petterson, B.A. Epidemiology of Hymenolepis nana infections in primary school children in urban and rural communities in Zimbabwe. J. Parasitol. 80, 245, 1994.
29. Quihui-Cota, L. et al. Prevalence and intensity of intestinal parasitic infections in relation to nutritional status in Mexican school children. Trans. R. Soc. Trop. Med. Hyg. 98, 653, 2004. 30. Maggi, P. et al. Hymenolepis nana parasites in adopted children. Clin. Infect. Dis. 41, 571, 2005.
31. Black, R.E., Lopez de, R., and Brown, K.H. Incidence and etiology of infantile diarrhea and major route of transmission in Hausecar, Peru. Am. J. Epidemiol. 129, 785, 1989.
32. Gama-Eddin, F.M., Aboul-Atta, A.M., and Hassounah, O.A. Extra-intestinal Hymenolepis nana cysticercoidiasis in asthamatic and �larised Egyptian patients. J. Egypt. Soc. Parasitol. 16, 517, 1986.
33. Vito, D.L., Cinza, R., and Giuseppe, A. Skin eruption associated with Hymenolepis nana infection. Int. J. Dermatol. 43, 357, 2004.
34. Al-Hussani, M.K. et al. Phlyctenular eye disease is association with Hymenolepis nana in Egypt. Br. J. Ophthalmol. 63, 627, 1979.
35. Olds, R.J. The affect of the tapeworm Hymenolepis nana on immunity to tuberculosis in mice. J. Hyg. 67, 233, 1969.
36. Fagbemi, B.O., and Christensen, N.O. Delayed expulsion of Hymenolepis diminuta in Trgpanosome brucei infected mice. Zeit, Parasit. 70, 663, 1984.
37. Lucas, S.B. et al. Aberrant form of Hymenolepis nana: Possible opportunistic infection in immunocompromised patients. Lancet 2, 1372–1373, 1979.
38. Alghali, S.T., Hagan, P., and Robinson, M. Hymenolepis citelli (Cestoda) and Nematospiroides dubins (Nemchi) inter speci�c interaction in mice. Exp. Parasitol. 60, 364, 1985.
39. Ito, A., and Yamamoto, Y. Mechanism of the parasite’s survival in the immunized host as well as the mechanism of immunity to reinfection. Primary and secondary infections with Hymenolepis nana eggs. 1. Changes in haematological values. Jap. J. Parasitol. 31, 203, 1976.
40. Isaak, D.D., Jacobson, R.H., and Reed, N.D. Thymus dependence of tapeworm Hymenolepis diminuta elimination from mice. Infect. Immun. 12, 1478, 1975.
41. Isaak, D.D., Jacobson, R.H., and Reed, N.D. The course of Hymenolepis nana infection in thymus de�cient mice. Int. Arch. Allergy Appl. Immunol. 55, 504, 1977.
42. Alghali, S.T.O., and Grenics, R.K. Immunity to tapeworms: Intra speci�c cross-protective interactions between Hymenolepis citelli, H. diminuta and H. microstoma in mice. Parasitology 92, 665, 1986. 43. Ito, A. et al. Hymenolepis nana: Immunity against oncosphere challenge in mice previously given viable or non-viable oncosphere of H. nana, H. diminuta, H. microstoma and Taenia taeniaforms. Int. J. Parasitol. 21, 241, 1991. 44. Ito, A., Onitake, K., and Andreasen, J. Lumen phase cross immunity between Hymenolepis microstoma and H. nana in mice. Int. J. Parasitol. 18, 1019, 1988. 45. Ito, A. Hymenolepis nana: Survival in the immunized mouse. Exp. Parasitol. 49, 248, 1980. 46. Van der Vorst et al. In�uence of an Hymenolepis diminuta infection on IgI and IgA antibodies to mouse intestinal eosinophus. Int. Arch. Allergy Appl. Immunol. 87, 281, 1988. 47. Ito, A. Basic and applied immunology in cestode infections from Hymenolepis to Taenia and Echinococcus. Int. J. Parasitol. 27, 1203, 1997. 48. Palmas, C., Wakelin, D., and Gabriele, F. Transfer of immunity against Hymenolepis nana in mice with lymphoid cells or serum from infected donors. Parasitology 89, 287, 1984. 49. Asano, K. et al. Hymenolepis nana: Passive transfer of mouse immunity by T-cell subset of phenotype Lyt-1. Exp. Parasitol. 61, 373, 1986. 50. Ajami, A., and Ra�ee, A. Cytokine production in Hymenolepis infection. Iran. J. Immunol. 4, 236, 2007. 51. Niwa, A., and Miyazato, T. Enhance of intestinal eosinophilia during H. nana infection in mice. J. Helminthol. 70, 33, 1996. 52. Ito, A. et al. Analysis of antibody responses to Hymenolepis nana infection in mice by the enzyme-linked immunosorbent assay and immunoprecipitator. Parasite Immunol. 16, 265, 1988. 53. Bortoletti, G., Gabriele, F., and Palmas, C. Mechanisms of protective immunity in Hymenolepis nana/mouse model. Parasitologia 34, 17, 1992. 54. Hopkins, C.A. Immunological memory in mice to adult Hymenolepis diminuta (Cestoda). J. Parasitol. 68, 32, 1982. 55. Craig, P., and Ito, A. Intestinal cestodes. Curr. Opin. Infect. Dis. 20, 524, 2007. 56. Voge, M. Concentrated method for helminthic ova and larvae. In: MacInnis, A.J., and Voge, M. (eds), Experiment and Techniques in Parasitology, W.H. Freeman, San Francisco, pp. 132–134, 1986. 57. Turant, A.L. Comparison of formalin–ethyl ether sedimentation, formalin–ethyl acetate sedimentation, and zinc sulfate �oatation technique for detection of intestinal parasites. J. Clin. Microbiol. 13, 882, 1981. 58. Lussier, G., and Loew, F.M. Natural Hymenolepis nana infection in Mongolian gerbils (Meriones unguiculatus). Can. Vet. J. 11, 5, 1970. 59. Movvsesyan, S.O. et al. Experimental hymenolepiasis of rats: Preliminary data on histological changes of visceral organs. Acta Parasitol. 53, 193, 2008. 60. Shimoda, K., Furakawa, T., and Miyazato, T. Clinicopathological changes of mice following primary and secondary infection with Hymenolepis nana eggs. 1. Changes in haematological values. Jap. J. Parasitol. 31, 203, 1982. 61. Ito, A. et al. Analysis of antibody response to Hymenolepis nana infection in mice by enzyme-linked immunosorbent assay and immunoprecipitation. Parasitol. Immunol. 10, 265, 1988. 62. Gomez-Priego, A., Godinez-Hana, A.L., and GulterrezQiuiraz, M. Detection of serum antibodies in human Hymenolepis –infection by enzyme immunoassay. Trans. R. Soc. Trop. Med. Hyg. 85, 645, 1991. 63. Castillo, R.M. et al. Effect of treatment on serum antibody to Hymenolepis nana detected by enzyme-linked immunosorbent assay. J. Clin. Microbiol. 29, 413, 1991.
64. Montenegro, T. et al. The diagnostic importance of species speci�c and cross reactive components of Taenia solium, Echinococcus granulosus and Hymenolepis nana, Rev. Inst. Med. Trop. Sao Paulo. 36, 327, 1994.
65. Gonenc, B. Analysis of the crude antigen of Hymenolepis nana from mice by SDS-PAGE and the determination of speci�c antigens in protein structure by Western blotting. Turk. J. Vet. Animal Sci. 26, 1067, 2002.
66. Mota, I. and Wong, D. Homologus and heterologus passive cutaneous anaphylaxis activity of mouse antisera during the course of immunization. Life Sci. 8, 813, 1969.
67. Ito, A. IgG and IgE antibodies to Hymenolepis nana detected in infected mouse sera by gel diffusion and passive cutaneous anaphylaxis tests. J. Parasitol. 70, 170, 1984.
68. Krawczuk, S., Rode, W., and Machnicka, B. Puri�cation and immunologic reactivity of Hymenolepis diminuta surface antigens. Parasitol. Res. 76, 707, 1990.
69. Larssh, J.E. Serological studies on the mouse strain of the dwarf tapeworm Hymenolepis nana var. fraterna. Am. J. Hyg. 37, 289, 1943.
70. Coleman, R.M., Carty, J.M., and Graziadei, W.D. Immunogenicity and phylogenetic relationship of tapeworm antigens produced by Hymenolepis nana and Hymenolepis diminuta. Immunology, 15, 297, 1968.
71. Macnish, M.G. et al. A molecular phylogeny of nuclear and mitochondrial sequences in Hymenolepis nana (Cestoda) supports the existence of a cryptic species. Parasitology 125, 567, 2002. 72. Okamoto, M. et al. Phylogenetic relationships of three hymenolepidid species inferred from nuclear ribosomal and mitochondrial DNA sequences. Parasitology 115, 661, 1997. 73. Mohammadzadeh, T. et al. Study on the genomic diversity of Hymenolepis nana between rat and mouse isolates by RAPDPCR. Iran. J. Vet. Res. 18, 16, 2007. 74. Walsh, P.S., Metzger, D.A., and Higuchi, R. Chelex 100 as a medium for simple extraction of DNA for PCR based typing from forensic material. BioTechniques 10, 506, 1991. 75. Paxinos, E. et al. A non-invasive method for distinguishing among canid species: Ampli�cation and enzyme restriction of DNA from dung. Mol. Ecol. 6, 483, 1997. 76. Heukelbach, J. et al. Selective mass treatment with ivermectin to control intestinal helminthiases and parasitic skin diseases in a severely affected population. Bull. W.H.O. 82, 563, 2004. 77. McCrckon, R.O., Lipkomitz, K.B., and Dronen, N.O. Ef�cacy of albendazole and mebendazole against Hymenolepis microstoma and Hymenolepis diminuta. Parasitol. Res. 78, 108, 1992. 78. Gupta, S., and Katiyar, J.C. Comparative activity of anticestode drugs praziquantel, niclosamide and compared 77-6 against Hymenolepis nana. J. Helminthol. 7, 31, 1983. 79. Andrews, P., Thomas, H., and Weber, H. The in-vitro uptake of 14C–Praziquantel by cestodes, trematodes and a nematode. J. Parasitol. 66, 920, 1980. 80. Chero, J.C. et al. Hymenolepis nana infection: Symptoms and response to nitazoxanide in �eld conditions. Trans. R. Soc. Trop. Med. Hyg. 101, 203, 2007. 25 Chapter 25 - Mesocestoides
1. Chandler, A.C. First record of a case of human infection with tapeworms of the genus Mesocestoides. Am. J. Trop. Med. 22, 493, 1942.
2. Gleason, N.N., and Healy, G.R. Report of a case of Mesocestoides (Cestoda) in a child in Missouri. J. Parasitol. 53, 83, 1967.
3. Gleason, N.N., Kornblum, R., and Walzer, P. Mesocestoides (Cestoda) in a child in New Jersey treated with Niclosamide (Yomesan ® ). Am. J. Trop. Med. Hyg. 22, 757, 1973.
4. Gutierrez, Y., Buchino, J.J., and Schubert, W.K. Mesocestoides (Cestoda) infection in children in the United States. J. Pediatr. 93, 245, 1978.
5. Hutchison, W.F., and Martin, J.B. Mesocestoides (Cestoda) in a child in Mississippi treated with paromomycin sulfate (Humatin ® ). Am. J. Trop. Med. Hyg. 29, 478, 1980.
6. Ohtomo, H. et al. Therapeutic effect of paramomycin sulfate on the 13th case of Mesocestoides lineatus infection found in Japan. Jpn. J. Antibiot. 36, 632, 1983.
7. Min, D.Y. Cestodes infections in Korea. Kor. J. Parasitol. 28, 123, 1990.
8. Eom, K.S., Kim, S.H., and Rim, H.J. Second case of human infection with Mesocestoides lineatus in Korea. Kor. J. Parasitol. 30, 147, 1992.
9. Schlutz, L.J. et al. Mesocestoides (Cestoda) infection in a California child. Pediatric Infect. Dis. J. 11, 332, 1992. 10. Fuentes, J.V., Galan-Puchades, M.T., and Malone, J.B. A new case report of human Mesocestoides infection in the United States. Am. J. Trop. Med. Hyg. 68, 566, 2003. 11. Marty, A.M., and Nea�e, R.C. Hymenolepiasis and miscellaneous cyclophyllidiases. In: Meyers, W.M. (ed.), Pathology of Infectious Diseases. Chapter 12. Armed Forces Institute of Pathology, American Registry of Pathology, Washington, 2000. 12. Garin, Y.J.F. et al. Case report: Human brain abscess due to a tetra-acetabulate plerocercoid metacestode (Cyclophyllidea). Am. J. Trop. Med. Hyg. 72, 513, 2005. 13. Ihalamulla, R.L., Gunatilaka, H., and Fernado, S.D. Human sparganosis in Sri Lanka: A case report and review of the cases. J. Parasit. Dis. 32, 30, 2008. 14. Reid, W.A., and Reardon M.J. Mesocestoides in the baboon and its development in laboratory animals. J. Med. Primatol. 5, 345, 1976. 15. Hubbard, G.B. et al. Mesocestoides infection in captive olive baboons (Papio cynocephalus anubis). Lab. Anim. Sci. 43, 625, 1993. 16. Fincham, J.E. et al. Pleural Mesocestoides and cardiac shock in an obese vervet monkey (Cercopithecus aethiops). Vet. Pathol. 32, 330, 1995. 17. Crosbie, P.R. et al. Diagnostic procedures and treatment of eleven dogs with peritoneal infections caused by Mesocestoides spp. J. Vet. Med. Assoc. 213, 1578,1998. 18. Crosbie, P.R., Padgett, K.A., and Boyce, W.M. Mesocestoides spp. tapeworm infections in dogs in California, Calif. Vet. 54, 15, 2000. 19. Bonfanti, U. et al. Clinical, cytological and molecular evidence of Mesocestoides sp. infection in a dog from Italy. J. Vet. Med. 51, 435, 2004. 20. Haziroglu, R. et al. Peritoneal tetrathyridosis in a Siamese cat—A case report. Vet. Archiv. 75, 543, 2005. 21. Venco, L. et al. Ultrasonographic features of peritoneal cestodiasis caused by Mesocestoides sp. in a dog and in a cat. Vet. Radiolol. Ultrasound 46, 417, 2005. 22. Eleni, C. et al. Proliferative peritoneal and pleural cestoiasis in a cat caused by metacestodes of Mesocestoides sp. Anatomohistopathological �ndings and genetic identi�cation. Parasite 14, 71, 2007. 23. Foronda, P. et al. First larval record of Mesocestoides in Carnivora of Tenerife (Canary Islands). J. Parasitol. 93, 138, 2007. 24. Wirtherle, N. et al. First case of canine peritoneal larval cestodosis caused by Mesocestoides lineatus in Germany, Parasitol. Int. 56, 317, 2007. 25. Crosbie, P.R. et al. Molecular systematics of Mesocestoides spp. (Cestoda: Mesocestoididae) from domestic dogs (Canis familiaris) and coyotes (Canis latrans), J. Parasitol. 86, 350, 2000. 26. Wardle, R.A., McLeod, J.A., and Radinovsky, S. Advances in the Zoology of Tapeworms 1950–1970. University of Minnesota Press, Minneapolis, p. 274, 1974. 27. Brooks, D.R., Hoberg, E.P., and Weekes, P.J. Preliminary phylogenetic systematic analysis of the major lineages of the Eucestoda (Platyhelminthes: Cercomeria). Proc. Biol. Soc. Wash. 104, 651, 1991. 28. Mariaux, J., A molecular phylogeny of the Eucestoda. J. Parasitol. 84, 114, 1998. 29. Hoberg, E.P. Jones, A., and Bray, R.A. Phylogenetic analysis among the families of the Cyclophyllidea (Eucestoda) based on comparative morphology, with new hypotheses for co-evolution in vertebrates. System. Parasitol. 42, 51, 1999.
30. Specht, D., and Voge, M. Asexual multiplication of Mesocestoides tetrathyridia in laboratory animals. J. Parasitol. 52, 268, 1965.
31. Eckert, J., Von Brand, T., and Voge, M. Asexual multiplication of Mesocestoides corti (Cestoda) in the intestine of dogs and skunks. J. Parasitol. 55, 241, 1969.
32. Loos-Frank, B. Shedding of gravid proglottids and destrobilation in experimental infections of foxes with Mesocestoides leptothylacus Loos-Frank, 1980 (Cestoda). J. Helminthol. 61, 213, 1987.
33. Rausch, R.L. Family Mesocestoididae Fuhrmann, 1907. In: Khalil, L.F., Jones, A., and Bray, R.A. (eds), Keys to the Cestode Parasites of Vertebrates. CAB International, Wallingford, 1994.
34. Schmidt, G.D., Handbook of Tapeworm Identi�cation. CRC Press, Boca Raton, FL, 675p, 1986.
35. Voge, M. North American cestodes of the genus Mesocestoides. J. Calif. Pub. Zool. 59, 125, 1955.
36. Padgett, K.A. et al. Systematics of Mesocestoides (Cestoda: Mesocestoididae): Evaluation of molecular and morphological variation among isolates. J. Parasitol. 91, 1435, 2005.
37. Etges, F.R. The proliferative tetrathyridium of Mesocestoides vogae sp. N. (Cestoda). J. Helminthol. Soc. Wash. 58, 181, 1991.
38. Gubányi, A., and Eszterbauer, E. Morphological investigation of Mesocestoides (Cestoda, Mesocestoididae) species parasitizing Vulpes vulpes in Hungary. Miscnea Zool. Hung. 12, 11, 1998.
39. Von Nickisch-Rosenegk, M., Lucius, R., and Loos-Frank, B. Contributions to the phylogeny of the Cyclophyllidea (Cestoda) inferred from mitochondrial 12S rDNA. J. Mol. Evol. 48, 586, 1999.
40. Literák, I. et al. Mesocesotides litteratus (Batsch, 1786) (Cestoda: Cyclophyllidea: Mesocestoididae) from the red fox: Morphological and 18S rDNA characterization of European isolates. Helminthologia 43, 191, 2006.
41. Loos-Frank, B. One or two intermediate hosts in the life cycle of Mesocestoides (Cyclophyllidea, Mesocestoididae)? Parasitol. Res. 77, 726, 1991.
42. Padgett, K.A., and Boyce, W.M. Life-history studies on two molecular strains of Mesocestoides (Cestoda: Mesocestoididae): Identi�cation of sylvatic hosts and infectivity of immature life stages. J. Parasitol. 90, 108, 2004.
43. Webster, J.D. Fragmentary studies on the life history of the cestode Mesocestoides latus. J. Parasitol. 35, 83, 1949. 44. Sharpilo, V.P., and Kornyushin, V.V. Locomotory activity of mature cestode proglottids in external environment and egg dissemination pattern they display. Vestnik Zoologii. 4, 24, 1995. 45. Padgett, K.A., and Boyce, W.M. Ants as �rst intermediate hosts of Mesocestoides on San Miguel Island, USA. J. Helminthol. 79, 67, 2005. 46. Soldatowa, A.P. A contribution to the study of the development cycle in the cestode Mesocestoides lineatus (Goeze, 1782) parasitic in carnivorous mammals. Doklady Akademii Nauk SSSR. 45, 310, 1944. 47. McAllister, C.T., and Conn, D.B. Occurrence of tetrathyridia of Mesocestoides sp. (Cestoidea: Cyclophyllidea) in North American anurans (Amphibia). J. Wildl. Dis. 26, 540,1990. 48. Voge, M. New host records for Mesocestoides (Cestoda: Cyclophyllidea) in California. Am. Midland. Nat. 49, 249, 1953. 49. McAllister, C.T. et al. A new host and locality record for Mesocestoides sp. tetrathyridia (Cestoidea: Cyclophyllidea), with a summary of the genus from snakes of the world. J. Parasitol. 77, 329, 1991. 50. Voge, M., and Berntzen, A.K. Asexual multiplication of larval tapeworms as the cause of fatal parasitic ascites in dogs. J. Parasitol. 49, 983, 1963. 51. Williams, J.F., Lindsay, M., and Engelkirk, P. Peritoneal cestodiasis in a dog. JAVMA. 186, 1103, 1985. 52. Roberts, L.R. and Janovy, J. Jr. Foundations of Parasitology, 8th ed., Chapter 20. McGraw Hill, New York, 2005. 53. Conn, D.B. The rarity of asexual reproduction among Mesocestoides tetrathyridia (Cestoda). J. Parasitol. 76, 453, 1990. 54. Widmer, E.A., and Specht, H.D. Isolation of asexually proliferative tetrathyridia (Mesocestoides sp.) from the southern Paci�c rattlesnake (Crotalus viridis helleri) with additional data from two previous isolates from the Great Basin fence lizard (Sceloporus occidentalis longipes). J. Parasitol. 78, 921, 1992. 55. Caruso, K.J. et al. Cytologic diagnosis of peritoneal cestodiasis in dogs caused by Mesocestodies sp. Vet. Clin. Pathol. 32, 50, 2003. 56. Hermánek J. Nonspeci�c immunomodulation in�uences resistance of mice to experimental infection with Mesocestoides corti and Ascaris sum. J. Helminthol. 65, 121,1991. 57. Poulin, R. The evolution of life history strategies in parasitic animals. Adv. Parasitol. 37, 107, 1996. 58. Alvarez, L.I., Mottier, M.L., and Lanusse, C.E. Drug transfer into target helminth parasites. Trends Parasitol. 23, 97, 2007. 26 Chapter 26 - Spirometra
7. Andersen, K.L., Description of musculature differences in spargana of Spirometra (Cestoda; Pseudophyllidea) and Tetrathyridia of Mesocestoides (Cestoda; Cyclophyllidea) and their value in identi�cation. J. Helminthol., 57, 331, 1983.
8. Alarcón de Noya, B., Torres, J.R., and Noya, O., Maintenance of Sparganum proliferum in vitro and in experimental animals. Int. J. Parasitol., 22, 835, 1992.
9. Mueller, J.F., The biology of Spirometra. J. Parasitol., 60, 3, 1974.
10. Fukushima, T., and Yamane, Y., How does the sparganosis occur? Parasitol. Today, 15, 124, 1999.
11. Gray, M.L. et al., Sparganosis in feral hogs (Sus scrofa) from Florida. J. Am. Vet. Med. Assoc., 215, 204, 1999.
12. Ooi, H.K. et al., Survey of Spirometra erinaceieuropaei in frogs in Taiwan and its experimental infection in cats. J. Helminthol., 74, 173, 2000.
13. Nithiuthai, S. et al., Waterborne zoonotic helminthiases. Vet. Parasitol., 126, 167, 2004.
14. Zhou, P. et al., Food-borne parasitic zoonoses in China: Perspective for control. Trends Parasitol., 24, 190, 2008.
15. Berger, L. et al., Severe sparganosis in Australian tree frogs. J. Wildl. Dis., 45, 921, 2009.
16. Dai, R.S. et al., Severe infection of adult dogs with helminths in Hunan Province, China poses signi�cant public health concerns. Vet. Parasitol., 160, 348, 2009.
17. Li, M.W. et al., Sparganosis in mainland China. Int. J. Infect. Dis., 15, e154, 2011.
18. Anantaphruti, M.T., Nawa, Y., and Vanvanitchaim, Y., Human sparganosis in Thailand: An overview. Acta Trop., 118, 171, 2011.
19. Norman, S.H., and Kreutner, A., Jr., Sparganosis: Clinical and pathologic observations in ten cases. South Med. J., 73, 297, 1980.
20. Kina, Y. et al., A glycoprotein from Spirometra erinaceieuropaei plerocercoids suppresses osteoclastogenesis and proin�ammatory cytokine gene expression. Int. J. Parasitol., 35, 1399, 2005.
21. Cho, J.H. et al., Subcutaneous and musculoskeletal sparganosis: Imaging characteristics and pathologic correlation. Skeletal Radiol., 29, 402, 2000.
22. Botterel, F., and Bourée, P., Ocular sparganosis: A case report. J. Travel Med., 10, 245, 2003.
23. Pampiglione, S., Fioravanti, M.L., and Rivasi, F., Human sparganosis in Italy. Case report and review of the European cases. APMIS, 111, 349, 2003.
24. Koo, J.H. et al., A case of sparganosis mimicking a varicose vein. Korean J. Parasitol., 44, 91, 2006.
25. Koo, M. et al., Cases and literature review of breast sparganosis. World J. Surg., 35, 573, 2011.
26. Wiwanitkit, V., A review of human sparganosis in Thailand. Int. J. Infect. Dis., 9, 312, 2005.
27. Grif�n, M.P., Tompkins, K.J., and Ryan, M.T., Cutaneous sparganosis. Am. J. Dermatopathol., 18, 70, 1996.
28. Uh, H.S. et al., Sparganosis infesting in the penis; a case report with review of Korean literatures. Korean J. Urol., 38, 873, 1997.
29. Sakamoto, T. et al., Testicular sparganosis in a child from Uruguay. Acta Trop., 88, 83, 2003.
30. Lim, D., Kim, C.S., and Kim, S.I., Sparganosis presenting as spermatic cord hydrocele in six-year-old boy. Urology, 70, 1223.e1, 2007.
31. Sarukawa, S. et al., Case of subcutaneous sparganosis: Use of imaging in de�nitive preoperative diagnosis. J. Dermatol., 34, 654, 2007. 32. Yun, S.J. et al., A case of vesical and scrotal sparganosis presenting as a scrotal mass. Korean J. Parasitol., 48, 57, 2010. 33. Lee, K.J., Myung, N.H., and Park, H.W., A case of sparganosis in the leg. Korean J. Parasitol., 48, 309, 2010. 34. Koh, T.W. et al., Empyema thoracis associated with sparganosis: A case report. Korean J. Thorac. Cardiovasc. Surg., 21, 761, 1988. 35. Shimada, M. et al., Two cases of sparganosis having onsets of pleural effusion. Jpn. J. Parasitol., 38, 145, 1989. 36. Oh, S.J., Chi, J.G., and Lee, S.E., Eosinophilic cystitis caused by vesical sparganosis: A case report. J. Urol., 149, 581, 1993. 37. Tanaka, S. et al., A case report of pleural sparganosis. Parasitol. Int., 46, 73, 1997. 38. Ishii, H. et al., A rare case of eosinophilic pleuritis due to sparganosis. Intern. Med., 40, 783, 2001. 39. Kim, J.R., and Lee, J.M., A case of intramuscular sparganosis in the sartorius muscle. J. Korean Med. Sci., 16, 378, 2001. 40. Sim, S. et al., A case of breast sparganosis. Korean J. Parasitol., 40, 187, 2002. 41. Kim, H.J. et al., A case of sparganosis in the submandibular triangle. J. Clin. Otolaryngol. Head Neck Surg., 16, 330, 2005. 42. Kim, D.H. et al., Pleural sparganosis: A case report. Korean J. Thorac. Cardiovasc. Surg., 39, 502, 2006. 43. Lee, J.H. et al., Abdominal sparganosis presenting as an abscess with �stulous communication to the bowel. Am. J. Roentgenol., 185, 1084, 2005. 44. Yang, K.M. et al., Sparganosis combined with inguinal hernia. Korean J. Urol., 46, 1366, 2005. 45. Iwatani, K. et al., Sparganum mansoni parasitic infection in the lung showing a nodule. Pathol. Int., 56, 674, 2006. 46. Park, J.H. et al., A surgically con�rmed case of breast sparganosis showing characteristic mammography and ultrasonography �ndings. Korean J. Parasitol., 44, 151, 2006. 47. Hong, S.J. et al., Breast and scrotal sparganosis: Sonographic �ndings and pathologic correlation. J. Ultrasound Med., 29, 1627, 2010. 48. Lee, J.H. et al., A case of sparganosis that presented as a recurrent pericardial effusion. Korean Circ. J., 41, 38, 2011. 49. Sen, D.K. et al., Cestode larva (Sparganum) in the anterior chamber of the eye. Trop. Geogr. Med., 41, 270, 50. Yoon, K.C. et al., Eyelid sparganosis. Am. J. Ophthalmol., 138, 873, 2004. 51. Subudhi, B.N. et al., Ocular sparganosis. J. Indian Med. Assoc., 104, 529, 2006. 52. Kubota, T., and Itoh, M., Sparganosis associated with orbital myositis. Jpn. J. Ophthalmol., 51, 311, 2007. 53. Yang, J.W., Lee, J.H., and Kang, M.S., A case of ocular sparganosis in Korea. Korean J. Ophthalmol., 21, 48, 2007. 54. Otranto, D., and Eberhard, M.L., Zoonotic helminths affecting the human eye. Parasit. Vectors, 4, 41, 2011. 55. Mentz, M.B. et al., Human ocular sparganosis in southern Brazil. Rev. Inst. Med. Trop. Sao Paulo, 53, 51, 2011. 56. Holodniy, M. et al., Cerebral sparganosis: Case report and review. Rev. Infect. Dis., 13, 155, 1991. 57. Jeong, S.C. et al., Cerebral sparganosis with intracerebral hemorrhage: A case report. Neurology, 50, 503, 1998. 58. Cummings, T.J. et al., Parasitic lesion of the insula suggesting cerebral sparganosis: Case report. Neuroradiology, 42, 206, 2000. 59. Kwon, J.H., and Kim, J.S., Sparganosis presenting as a conus medullaris lesion; case report and literature review of the spinal sparganosis. Arch. Neurol., 61, 1126, 2004.
60. Rengarajan, S., Nanjegowda, N., and Bhat, D., Cerebral sparganosis: A diagnostic challenge. Br. J. Neurosurg., 22, 784, 2008.
61. Ou, Q., Li, S.J., and Cheng, X.J., Cerebral sparganosis: A case report. Biosci. Trends, 4, 145, 2010.
62. Moulinier, R. et al., Human proliferative sparganosis in Venezuela: Report of a case. Am. J. Trop. Med. Hyg., 31, 358, 1982.
63. Nakamura, T. et al., Human proliferative sparganosis. Am. J. Clin. Pathol., 94, 224, 1990.
64. Meric, R. et al., Disseminated infection caused by Sparganum proliferum in an AIDS patient. Histopathology, 56, 824, 2010.
65. Kong, Y. et al., Cleavage of immunoglobulin G by excretorysecretory cathepsin S-like protease of Spirometra mansoni plerocercoid. Parasitology, 109, 611, 1994.
66. Kong, Y. et al., Differential expression of the 27 kDa cathepsin L-like cysteine protease in developmental stages of Spirometra erinacei. Korean J. Parasitol., 38, 195, 2000.
67. Song, T., Wang, W.S., and Mai, W.W., CT and MR characteristics of cerebral sparganosis. Am. J. Neuroradiol., 28, 1700, 2007.
68. Morakote, N., and Kong, Y., Antigen speci�city of 36 and 31 kDa proteins of Spirometra erinacei plerocercoid in tissue invading nematodiasis. Korean J. Parasitol., 31, 169, 1993.
69. Yeo, I.S., Yong, T.S., and Im, K., Serodiagnosis of human sparganosis by a monoclonal antibody-based competition ELISA. Yonsei Med. J., 35, 43, 1994.
70. Chung, Y.B. et al., IgG antibody responses in early experimental sparganosis and IgG subclass responses in human sparganosis. Korean J. Parasitol., 38, 145, 2000.
71. Rahman, M., Lee, E.G. and Bae, Y.A., Two-dimensional immunoblot analysis of antigenic proteins of Spirometra plerocercoid recognized by human patient sera. Parasitol. Int., 60, 139, 2011. 72. Liu, D.-W., Kato, H., and Sugane, K., The nucleotide sequence and predicted secondary structure of small subunit (18S) ribosomal RNA from Spirometra erinaceieuropaei. Gene, 184, 221, 1997. 73. Brabec, J., Kuchta, R., and Scholz, T., Paraphyly of the Pseudophyllidea (Platyhelminthes: Cestoda): Circumscription of monophyletic clades basedon phylogenetic analysis of ribosomal RNA. Int. J. Parasitol., 36, 1535, 2006. 74. Okamoto, M. et al., Intraspeci�c variation of Spirometra erinaceieuropaei and phylogenetic relationship between Spirometra and Diphyllobothrium inferred from mitochondrial CO1 gene sequences. Parasitol. Int., 56, 235, 2007. 75. Lee, S.U., Huh, S., and Phares, C.K., Genetic comparison between Spirometra erinacei and S. mansonoides using PCRRFLP analysis. Korean J. Parasitol., 35, 277, 1997. 76. Liu, W. et al., Survey of Spirometra erinaceieuropaei spargana infection in the frog Rana nigromaculata of the Hunan Province of China. Vet. Parasitol., 173, 152, 2010. 77. Zhu, X.Q. et al., Single-strand conformation polymorphismbased analysis reveals genetic variation within Spirometra erinacei (Cestoda: Pseudophyllidea) from Australia. Mol. Cell. Probes, 16, 159, 2002. 78. Kim, D.G. et al., Cerebral sparganosis: Clinical manifestations, treatment, and outcome. J. Neurosurg., 85, 1066, 1996. 79. Noya, O. et al., Sparganum proliferum: An overview of its structure and ultrastructure. Int. J. Parasitol., 22, 631, 1992. 80. Cui, J. et al., Serodiagnosis of experimental sparganum infections of mice and human sparganosis by ELISA using ES antigens of Spirometra mansoni spargana. Parasitol. Res., 108, 1551, 2011. 27 Chapter 27 - Taenia
1. Verster, A. A taxonomic revision of the genus Taenia Linnaeus, 1758s. str. Onderstepoort J. Vet. Res. 36, 3–58, 1969.
2. Loos-Frank, B. An up-date of Verster’s, 1969.“Taxonomic revision of the genus Taenia Linnaeus” (Cestoda) in table format. Syst. Parasitol. 45, 155–183, 2000.
3. Eom, K.S., and Rim, H.J. Morphologic descriptions of Taenia asiatica sp. Korean J. Parasitol. 31, 1–6, 1993.
4. Rausch, R.L. Taenia pencei sp. from the Ringtail, Bassariscus astutus (Carnivora: Procyonidae), in Texas, U.S.A. Comp. Parasitol. 70, 1–10, 2003.
5. Hoberg, E.P., Phylogeny of Taenia: Species de�nitions and origins of human parasites. Parasitol. Int. 55(Suppl.), S23–S30, 2006.
6. Abuladze, K.I. Taeniata of animals and man, and diseases caused by them. Akademiya Nauk USSR, Moscow, pp 59–67, 1964. (translated by Israel’s Program for Scienti�c Translation, Jerusalem, 1970.)
7. Soulsby, E.J.L. Helminths, Arthropods and Protozoa of Domesticated Animals. Lea & Febiger, Philadelphia, 1982.
8. Eom, K.S. What is Asian Taenia? Parasitol. Int. 55 (Suppl.), S137–S141, 2006.
9. Ito, A., Nakao, M., and Wandra, T. Human taeniasis and cysticercosis in Asia. Lancet 362, 1918–1920, 2003.
10. Ito, A. et al. Dogs as alternative intermediate hosts of Taenia solium in Papua (Irian Jaya), Indonesia con�rmed by highly speci�c ELISA and immunoblot using native and recombinant antigens and mitochondrial DNA analysis. J. Helminthol. 76, 311–314, 2002.
11. Ito, A. et al. Neurocysticercosis: Clinical manifestation, neuroimaging, serology and molecular con�rmation of histopathologic specimens. Southeast Asian J. Trop. Med. Public Health 37(Suppl 3), 74–81, 2006.
12. Wandra, T. et al. Current situation of taeniasis and cysticercosis in Indonesia. Trop. Med. Health 35, 323–328, 2007. 13. Schantz, P.M. et al. Neurocysticercosis in an orthodox Jewish community in New York City, N. Engl. J. Med. 327, 692–695, 1992.
14. Al Shahrani, D. et al. First case of neurocysticercosis in Saudi Arabia. J. Trop. Pediatr. 49, 58–60, 2003.
15. Hira, P.R. et al. Cysticercosis: Imported and autochthonous infections in Kuwait. Trans. R. Soc. Trop. Med. Hyg. 98, 233–239, 2004.
16. Ito, A. et al. Chapter 62. Taenia, In: Liu, D. (ed), Molecular Detection of Foodborne Pathogens, Taylor & Francis Group, London, pp. 827–838, 2009.
17. Okamoto, M. et al. Asian Taenia: Species or subspecies? Southeast Asian J. Trop. Med. Public Health, 38 (Suppl 1), 125–130, 2007.
18. Fan, P.C. Taiwan Taenia and taeniasis, Parasitol. Today. 4, 86–88, 1988.
19. Okamoto, M. et al. Molecular variation of Taenia solium in the world, Southeast Asian J. Trop. Med. Public Health, 32(Suppl 2), 90–93, 2001.
20. Nakao, M. et al. A phylogenetic hypothesis for the distribution of two genotypes of the pig tapeworm Taenia solium worldwide. Parasitology 124, 657–662, 2002. 21. Ito, A. et al. Chapter 5. Mitochondrial DNA of Taenia solium: From basic to applied science. In: Singh, G., and Prabhakar, S. (eds), Taenia solium Cysticercosis, CAB International, 47–55, 2002. 22. Chung, J.Y. et al. Molecular determination of the origin of acephalic cysticercus. Parasitology 130, 239–246, 2005. 23. Hinojosa-Juarez, A.C. et al. Genetic similarity between cysticerci of Taenia solium isolated from human brain and from pigs. Infect. Genet. Evol. 8, 653–656, 2008. 24. Anantaphruti, M.T. Molecular and serological survey on taeniasis and cysticercosis in Kanchanaburi Province, Thailand. Parasitol. Int.. 59, 326, 2010. 25. Dryden, M.W., Payne, P.A., and Smith, V. Accurate diagnosis of Giardia spp and proper fecal examination procedures. Vet. Therapeutics 7, 4–14, 2006. 26. Okamoto, M. et al. Evidence of hybridization between Taenia saginata and Taenia asiatica. Parasitol. Int. 59, 70–74, 2010. 27. Yamasaki, H. et al. Mitochondrial DNA diagnosis for taeniasis and cysticercosis. Parasitol. Int. 55, S81, 2006. 28. Okamoto, M., and Ito, A. Puri�cation of nucleic acids from helminths. In: Liu, D. (ed), Handbook of Nucleic Acid Puri�cation, Chapter 13, Taylor & Francis Group, London, pp. 271–291, 2009. 29. Azuma, H. et al. Intraspeci�c variation of Taenia taeniaeformis as determined by various criteria. Parasitol. Res. 81, 103, 1995. 30. Yap, K.W., and Thompson, R.C.A. CTAB precipitation of Cestode DNA. Parasitol. Today. 3, 220, 1987. 31. Yap, K.W. et al. Taenia hydatigena: Isolation of mitochondrial DNA, molecular cloning, and physical mitochondrial genome mapping. Exp. Parasitol. 63, 288, 1987. 32. Okamoto, M. et al. Phylogenetic relationships of three hymenolepidid species inferred from nuclear ribosomal and mitochondrial DNA sequences, Parasitology 115, 661, 1997. 33. Copeland, C.S. et al. Identi�cation of the Boudicca and Sinbad retrotransposons in the genome of the human blood �uke Schistosoma haematobium. Mem. Inst. Oswaldo Cruz 101, Online. 2006. 34. Matoba, Y. et al. Detection of a taeniid species Taenia taeniaeformis from a feral raccoon Procyon lotor and its epidemiological signi�cance. Mamm. Study. 28, 157, 2003. 35. Yamasaki, H. et al. DNA differential diagnosis of taeniasis and cysticercosis by multiplex PCR. J. Clin. Microbiol. 42, 548, 2004. 36. Gobert, G.N. et al. Copro-PCR based detection of Schistosoma eggs using mitochondrial DNA markers. Mol. Cell. Probes 19, 250, 2005. 37. Li, T. et al. Taeniasis/cysticercosis in a Tibetan population in Sichuan Province, China. Acta Trop. 100, 223, 2006. 38. Harmon, A.F. et al. Improved methods for isolating DNA from Ostertagia ostertagi eggs in cattle feces. Vet. Parasitol. 135, 297, 2006. 39. Nunes, C.M. et al. Fecal specimens preparation methods for PCR diagnosis of human taeniosis. Rev. Inst. Med. Trop., Sao Paulo 48, 45, 2006. 40. Rishi, A.K., and McManus, D.P. DNA probes which unambiguously distinguish Taenia solium from T. saginata, Lancet 2(8570), 1275–1276, 1987. 41. Flisser, A. et al. Specific detection of Taenia saginata eggs by DNA hybridization. Lancet 2(8625), 1429–1430, 1988. 42. Harrison, L.J., Delgado, J., and Parkhouse, R.M. Differential diagnosis of Taenia saginata and Taenia solium with DNA probes, Parasitology 100, 459–461, 1990.
43. Chapman, A. et al. Isolation and characterization of speciesspeci�c DNA probes from Taenia solium and Taenia saginata and their use in an egg detection assay. J. Clin. Microbiol. 33, 1283–1288, 1995.
44. González, L.M. et al. PCR tools for the differential diagnosis of Taenia saginata and Taenia solium taeniasis/cysticercosis from different geographical locations. Diagn. Microbiol. Infect. Dis. 42, 243–249, 2002.
45. Nunes, C.M. et al. Taenia saginata: Differential diagnosis of human taeniasis by polymerase chain reaction-restriction fragment length polymorphism assay. Exp. Parasitol. 110, 412–415, 2005.
46. Gasser, R.B., Zhu, X., and Woods, W. Genotyping Taenia tapeworms by single-strand conformation polymorphism of mitochondrial DNA. Electrophoresis 20, 2834–2837, 1999.
47. Vega, R. et al. Population genetic structure of Taenia solium from Madagascar and Mexico: Implications for clinical pro�le diversity and immunological technology. Int. J. Parasitol. 33, 1479–1485, 2003.
48. Dias, A.K. et al. Taenia solium and Taenia saginata: Identi�cation of sequence characterized ampli�ed region (SCAR) markers. Exp. Parasitol. 117, 9–12, 2007.
49. Yamasaki, H. et al. DNA differential diagnosis of human taeniid cestodes by base excision sequence scanning thymine-base reader analysis with mitochondrial genes. J. Clin. Microbiol. 40, 3818–3821, 2002.
50. González, L.M. et al. Differential diagnosis of Taenia saginata and Taenia solium infection by PCR. J. Clin. Microbiol. 38, 737–744, 2000.
51. Jeon, H.K. et al. Differential diagnosis of Taenia asiatica using multiplex PCR. Exp. Parasitol. 121, 151–156, 2009.
52. Bowles, J., and McManus, D.P. Genetic characterization of the Asian Taenia, a newly described taeniid cestodes of humans. Am. J. Trop. Med. Hyg. 50, 33–44, 1994. 53. von Nickisch-Rosenegk, M., Silva-Gonzalez, R., and Lucius, R. Modi�cation of universal 12S rDNA primers for speci�c ampli�cation of contaminated Taenia spp. (Cestoda) gDNA enabling phylogenetic studies. Parasitol. Res. 85, 819–825, 1999. 54. Okamoto, M. et al. Phylogenetic relationships within Taenia taeniaeformis variants and other taeniid cestodes inferred from the nucleotide sequence of the cytochrome c oxidase subunit I gene. Parasitol. Res. 81, 451–458, 1995. 55. Nakao, M. et al. Mitochondrial genetic code in cestodes. Mol. Biochem. Parasitol. 111, 415–424, 2000. 56. Nakao, M. et al. A phylogenetic hypothesis for the distribution of two genotypes of the pig tapeworm Taenia solium worldwide. Parasitology 124, 657–662, 2002. 57. Nkouawa, A. et al. Loop-mediated isothermal ampli�cation method for differentiation and rapid detection of Taenia species. J. Clin. Microbiol. 47, 168–174, 2009. 58. Nakao, M., Sako, Y., and Ito, A., Isolation of polymorphic microsatellite loci from the tapeworm Echinococcus multilocularis. Infect. Genet. Evol. 3, 159–163, 2003. 59. Bartholomei-Santos, M.L. et al. Isolation and characterization of microsatellites from the tapeworm Echinococcus granulosus. Parasitology 126, 599–605, 2003. 60. Bart, J.M. et al. EmsB, a tandem repeated multiloci microsatellite, new tool to investigate the genetic diversity of Echinococcus multilocularis. Infect. Genet. Evol. 6, 390–400, 2006. 61. Knapp, J. et al. Genetic diversity of the cestode Echinococcus multilocularis in red foxes at a continental scale in Europe. PLoS Negl. Trop. Dis. 3, e452, 2009. 62. Campbell, G. et al. Genetic variation in Taenia solium. Parasitol. Int. 55, S121–S126, 2006. 63. Anantaphruti, M.T. et al. Sympatric occurrence of Taenia solium, T. saginata, and T. asiatica, Thailand. Emerg. Infect. Dis. 13, 1413–1416, 2007.
Section IV
Platyhelminthes: Trematoda 28 Chapter 28 - Clonorchis
4. McConnell, J.F.P. Anatomy and pathological relations of a new species of liver �uke. Lancet 2, 271, 1875.
5. Loos, A. Clonorchis (1907). Cited in: Yamaguti, S. (ed.), Synopsis of Digenetic Trematodes of Vertebrates. Keigaku, Tokyo, 1971.
6. Ijiima, I. Notes on Distoma endemicum Baelz. J. Coll. Sci. Imperial Univ. Jpn. 1, 47, 1887.
7. Kobayashi, H. 1914. Fish as intermediate hosts in clonorchiasis. Cited in: Despommier, D.D., Gwadz, R., and Hotez, P.J., Parasitic Diseases. Springer Verlag, Heidelberg, 1995.
8. Muto, S. On the primary intermediate host of Clonorchis sinensis. Chuo Igakakai Zasshi 25, 49, 1918.
9. Song, I.C., Lee, J.S., and Rim, H.J. Epidemiological studies on the distribution of Clonorchis sinensis infection in Korea (in Korean). Korean Univ. Med. J. 20, 165, 1983.
10. Haswell-Elkins, M.R. Lung and liver �ukes. In: Cox, F.E.G. et al. (eds), Parasitology, Topley and Wilson’s Microbiology and Microbial Infections, 10th edn., Hoddes Arnold, ASM Press, Washington, 2005.
11. Wykoff, D.E. Studies on Clonorchis sinensis IV: Production of eggs in experimentally infected rabbits. J. Parasitol. 45, 91, 1959.
12. Haas, W., Granzer, M., and Brockelman, C.R. Opisthorchis viverrini: Finding and recognition of the �sh host by the cercariae. Exp. Parasitol. 71, 422, 1990.
13. Sithithaworn, P. et al. Relationship between faecal egg count and worm burden of Opisthorchis viverrini in human autopsy cases. Parasitology 102, 277, 1991.
14. Piekarski, G. Medizinische Parasitologie in Tafeln. Springer Verlag, Heidelberg, 1987.
15. Chai, J.Y., Murrel, K.D., and Lymbery, A.J. Fish-borne parasitic zoonoses: Status and issues. Int. J. Parasitol. 35, 1233, 2005.
16. Kim, Y.H. Carcinoma of the gall bladder associated with clonorchiasis: Clinicopathologic and CT evaluation. Abdom. Imaging 28, 83, 2003.
17. Leung, J.W. et al. Hepatic clonorchiasis—A study by endoscopic retrograde cholangiopancreatography. Gastrointest. Endosc. 35, 226, 1989.
18. Leung, J.W., and Yu, A.S. Hepatolithiasis and biliary parasites. Bull. Clin. Gastroenterol. 11, 681, 1997.
19. Chen, M.G., Lu, Y., and Hua, X.J. Progress in assessment of morbidity due to Clonorchus sinensis infection: A review of recent literature. Trop. Dis. Bull. 91, 7, 1994.
20. Bunnag, D., and Harinasuta, T. Opisthorchiasis, clonorchiasis and paragonimiasis. In: McGraw, R.P., and Mclvor, D. (eds), Tropical and Geographic Medicine. McGraw-Hill, New York, 1984.
21. MacFadzean, A.J.S., and Yeung, R.T.T. Acute pancreatitis due to Clonorchis sinensis. Trans. R. Soc. Trop. Med. Hyg. 60, 466, 1966.
22. Hou, P.C. The relationship between primary carcinoma of the liver and infestation with Clonorchis sinensis. J. Pathol. Bacteriol. 72, 239, 1965. 23. Komiya, Y. Clonorchis and clonorchiasis. Adv. Parasitol. 4, 53, 1966. 24. Mehlhorn, H. et al. Ultrastructural investigations on the effects of praziquantel on human trematodes from Asia (Clonorchis sinensis, Metagonimus yokogawai, Opisthorchis viverrini, Paragonimus westermani, Schistosoma japonicum). Drug Res. 33, 91, 1983. 25. Rim, H.J. et al. Clinical evaluation of the therapeutic ef�cacy of praziquantel (Embay 8440) against Clonorchis sinensis infection in man. Ann. Trop. Med. Parasitol. 75, 27, 1981. 26. Rim, H.J. The current pathology and chemotherapy of clonorchiasis. Korean J. Parasitol. 24 (Suppl. 3), 27, 1986. 27. Hänel, H., and Raether, W. Chemotherapy. In: Mehlhorn, H. (ed), Encyclopedia of Parasitology, Vols. 1 and 2. Springer, Heidelberg, New York, 2008. 28. Feldheim, W., and Knobloch, J. Serodiagnosis of Opisthorchis viverrini infestation by an enzyme immuno-assay. Tropenmed. Parasitol. 33, 8, 1982. 29. Müller, B., Schmidt, J., and Mehlhorn, H. Sensitive and species—Speci�c detection of Clonorchis sinensis by PCR in infected snails and �shes. Parasitol. Res. 100, 911, 2007. 30. Le, T.H. et al. Clonorchis sinensis and Opisthorchis viverrini: Development of a mitochondrial-based, multiplex PCR for their identi�cation and discrimination. Exp. Parasitol. 112, 109, 2006. 31. Müller, B., Schmidt, J., and Mehlhorn, H. PCR diagnosis of infections with different species of Opisthorchiidae using a rapid clean-up procedure for stool samples and speci�c primers. Parasitol. Res. 100, 905, 2007. 32. WHO, Manual of Basic Techniques. Geneva, Switzerland, 1995. 33. Schwartz, D.A. Review: Helminths in the induction of cancer: Opisthorchis viverrini, Clonorchis sinensis and cholangiocarcinoma. Trop. Geogr. Med. 32, 95, 1980. 34. Schuster, R., and Leberegelbefall, In: Aspöck H., Amoeben, Bandwürmer etc. und parasitäre Erkrankungen des Menschen in Mitteleuropa. Denisia 6, 291, 2002. 35. Wilde, J., Eiden, J., and Yoken, R. Removal of inhibitory substances from human fecal specimens for detection of group a rotaviruses by transcriptase and polymerase chain reactions. J. Clin. Microbiol. 28, 1300, 1990. 36. Wonggratanacheewin, S. et al. Detection of Opisthorchis viverrini in human stool specimens by PCR. J. Clin. Microbiol. 40, 3879, 2002. 37. Ando, K. et al. Nucleotide sequence of mitochondrial CO I and ribosomal ITS II genes of Opisthorchis viverrini in northeast Thailand. Southeast Asian J. Trop. Med. Public Health 32 (Suppl 2), 17, 2001. 38. Harnnoi, T. et al., Speci�c gravity of Opisthorichis viverrini egg. J. Helminthol. 72, 359, 1998. 39. Rahman, S.M. et al. Early detection and estimation of infection burden by real-time PCR in rats experimentally infected with Clonorchis sinensis. Parasitol. Res. 110, 72–78, 2011. 29 Chapter 29 - Dicrocoelium
TABLE 29.1
List of PCR and Sequencing Primers for
Dicrocoelium spp.
Site Direction Name Sequence (5’–3’) Reference
18S Forward Dd18SF1 GATAACGGGTAACGGGGAAT [10]
18S Reverse Dd18SR1 AACCTCTGACTTTCGCTCCA [10]
18S Forward Dd18SF2 TGGAGCGAAAGTCAGAGGTT [10]
18S Reverse Dd18SR2 GTACAAAGGGCAGGGACGTA [10]
5.8S Forward Dd58SF1 ATATTGCGGCCATGGGTTAG [10]
28S Reverse Dd28SR1 ACAAACAACCCGACTCCAAG [10] (Trematoda, Digenea) eggs in the faeces of lambs and ewes in porma basin (Leòn, Nw Spain). Ann. Parasitol. Hum. Comp. 66, 57, 1991.
21. Malek, A.E. Occurrence of Dicrocoelium hospes in Mali and Senegal, West Africa. J. Helminthol. 54, 45, 1980.
22. Asanji, M.F., and Williams, M.O. The effect of sex on seasonal variation on single and double infection of cattle in Sierra Leone by Dicrocoelium hospes and Fasciola gigantica. Vet. Parasitol. 15, 247, 1984.
23. Wenker, C. et al. Dikrozoliose bei Neuweltkameliden. Tieraztl. Prax. 26, 355, 1998.
24. Gunsser, I., Hanichen, T., and Maierl, J. Leberegelbefall bei Neuweltkameliden. Liver �uke in South American camelids. Tierarztl. Prax. 27, 187, 1999.
25. Somvanshi, R., and Kaul, G.L. Dicrocoeliasis in Pashima goats. Ind. Vet. Med. J. 13, 42, 1989.
26. Jithendran, K.P., and Bhat, T.K. Prevalence of dicrocoeliosis in sheep and goats in Himachal Pradesh, India. Vet. Parasitol. 61, 265, 1996.
27. Tang, C. et al. Studies on the biology and epizootiology of Dicrocoelium chinensis in the eastern Inner Mongol Autonomous region. Acta Zool. Sinica. 29, 340 (1983) (in Chinese).
28. Tang, C. et al. Investigations on the biology of a few Dicrocoelium species of sheep on Qinghai Plateau. Acta Zool. Sinica. 31, 254 (1985) (in Chinese).
29. Chartier, C., and Reche, B. Gastrointestinal helminths and lungworms of French dairy goats: Prevalence and geographical distribution in poitou-charentes. Vet. Res. Comm. 16, 327, 1992.
30. Boray, J.C. Flukes of domestic animals In: Gafaar, S.M., Howard, W.E., and Marsh, R.E. (eds), Parasites, Pests and Predators. Elsevier, Amsterdam, pp. 179–218, 1985.
31. Alunda, J.M., and Rojo-Vàzquez’s, F.A. Survival and infectivity of Dicrocoelium dendriticum eggs under �eld conditions in NW Spain. Vet. Parasitol. 13, 245, 1983.
32. Manga-González, M.Y. Some aspects of the biology and helminthofauna of Helicella (Helicella) itala (Linnaeus, 1758) (Mollusca) Natural infection by Dicrocoeliidae (Trematoda). Rev. Ibér Parasitol. Extraordinario, 131, 1987.
33. Fasanella, A., Lia, R., and Giangaspero, A. Cernuella virgata (Mollusca: Gasteropoda: Pulmonata) hôte intermédaire de Dicrocoelium dendriticum (Rudholphi, 1819) dans la region de Puglia, Italie. Parasite. 2, 331, 1995.
34. Manga-Gonzàlez, M.Y. et al. Studies on experimental infection by Dicrocoelium dendriticum (Trematoda) of Cernuella (Xeromagna) cespitum arigonis (Mollusca) specimens in a natural environment. In: Guerra, A., Rolan, E., and Rocha, F. (eds), Abst. 12th Int. Malacol. Congr. Vigo, 1995.
35. Gonzàlez-Lanza, C. et al. Larval development of Dicrocoelium dendriticum in Cernuella (Xeromagna) cespitum arigonis under controlled laboratory conditions. J. Helminthol. 71, 311, 1997.
36. Schuster, R. Infection patterns in the �rst intermediate host of Dicrocoelium dendriticum. Vet. Parasitol. 47, 235, 1993.
37. Alunda, J.M., and Rojo-Vàzquez, F.A. Survival and infectivity of Dicrocoelium dendriticum eggs under �eld conditions in NW Spain. Vet. Parasitol. 13, 245, 1983.
38. Schuster, R. Zur beein�ussung von Helicella obvia durch Dicrocoelium- Parthenitae. Angew. Parasitol. 33, 61, 1992.
39. Gomez, B.J. et al. Alteraciones histopatologicas producidas por Dicrocoelium dendriticum (Trematoda) en dos especies de Cernuella (Mollusca) sacri�cadas a los tres meses postinfestacion Iberus. 14, 189, 1996. 40. Schuster, R., Factors in�uencing the metacercarial intensity in ants and the size of Dicrocoelium dendriticum metacercarial cysts. J. Helminthol. 65, 275, 1991. 41. Wikler, W. Evolution-oriented ethology, kin selection, and altruistic parasites. Z. Tierpsyschol. 42, 206, 1976. 42. Manga-González, M.Y., and González-Lanza, C. Field and experimental studies on Dicrocoelium dendriticum and dicrocoeliasis in northern Spain. J. Helminthol. 79, 291, 2005. 43. González-Lanza, C. et al. Coprological study of the Dicrocoelium dendriticum (Digenea) egg elimination by cattle in highland areas in León Province, Northwest Spain. Parasitol. Res. 79, 488, 1993. 44. Sotiraki, S.T. et al. The effect of transportation and con�nement stress on egg production by Dicrocoelium dendriticum in sheep. J. Helminthol. 73, 337, 1999. 45. Eckert, J., and Hertzberg, H. Parasite control in transhumant situations. Vet. Parasitol. 54, 103, 1994. 46. Sanchez-Campos, S. et al. Morphologic and biochemical changes caused by experimentally induced dicroceliosis in hamsters (Mesocricetus auratus). Comp. Med. 50, 147, 2000. 47. Sanchez-Campos, S., Tunon Gonzalez, P., and GonzalezGallego, J. Oxidative stress and changes in liver antioxidant enzymes induced by experimental dicrocoeliosis in hamsters. Parasitol. Res. 85, 468, 1999. 48. Holmes, P.H. et al. The anaemia in fasciolosis studies with G-labelled red cells. J. Comp. Pathol. 78, 415, 1968. 49. Dargie, J.D., and Allonby, E.W. Pathophysiology of single and challenge infections of Haemonchus contortus in Merino sheep: Study on the red cell kinetics and the “self-cure” phenomenon. Int. J. Parasitol. 5, 147, 1975. 50. Wolff, K., Hauser, B., and Wild, P. Dicrocoeliose des schafes: Untersuschungen zur pathogenese und zur regeneration der laber nach therapie. Berl. Münch Tierärztl. Wschr. 97, 378, 1984. 51. Camara, L., P�ster, K., and Aeschlimann, A. Analyse Histopathologique de foie de bovin infestè par Dicrocoelium dendriticum. Vet. Res. 27, 87, 1996. 52. Theodoridis, Y. et al. Pathophysiological studies on Dicrocoelium dendriticum infection in sheep. Vet. Parasitol. 39, 61, 1991. 53. Wedrychowicz, H. et al. Local antibody responses in the bile and faeces of sheep infected with Fasciola hepatica. Res. Vet. Sci. 37, 44, 1984. 54. Wedrychowicz, H. et al. Somatic antigens of adult Dicrocoelium dendriticum recognised by bile antibodies of naturally infected cattle. Vet. Parasitol. 56, 47, 1995. 55. Haridy, F.M. et al. A preliminary study on dicrocoeliasis in Egypt, with a general review. J. Egypt Soc. Parasitol. 33, 85, 2003. 56. Stahel, E. Spurious infections by the small liver �uke (Dicrocoelium dendriticum) in Switzerland 1976–1980 Stahel E. Schweiz Med Wochenschr. 111, 1159, 1981. 57. el-Shiekh Mohamed, A.R., and Mummery, V. Human dicrocoeliasis. Report on 208 cases from Saudi Arabia. Trop. Geogr. Med. 42, 1, 1990. 58. Helmy, M.M., and Al-Mathal, E.M. Human infection with Dicrocoelium dendriticum in Riyadh district (Saudi Arabia). J. Egypt Soc. Parasitol. 33, 139, 2003. 59. Ondriska, F. et al. A rare case of human autochthonous dicrocoeliasis in Czechoslovakia. Bratisl. Lek Listy 90, 467, 1989. 60. Jelinek, T., Rack, J., and Adusu, E. Human infection with Dicrocoelium dendriticum, Deut. Med. Wochenschr. 129, 2538, 2004.
61. Karadag, B. et al. An unusual case of biliary obstruction caused by Dicrocoelium dentriticum. Scand J Infect Dis. 37, 385, 2005.
62. Sing, A., Tybus, K., and Fackler, I. Acute urticaria associated with Dicrocoelium dendriticum infestation. Indian J. Med. Microbiol. 26, 97, 2008.
63. Rehbein, S., Kokott, S., and Lindner, T. Evaluation of techniques for the enumeration of Dicrocoelium eggs in sheep faeces, Zentralbl. Veterinarmed A. 46, 133, 1999.
64. Campo, R., Manga-Gonzàlez, M.Y., and Gonzàlez-Lanza, C. Relationship between egg output and parasitic burden in lambs experimentally infected with different doses of Dicrocoelium dendriticum (Digenea). Vet. Parasitol., 87, 139, 2000.
65. Schröder, L., and Geyer, E. Experimentelle Dikrozöliose: Bildung humoraler Antikörper beim Goldhamster (Mesocricetus auratus). Behring Inst. Mitt. 60, 24, 1976.
66. Calamel, M. Application de la technique d’immuno�uorescence indirecte a l’étude épidemiologique de la dicrocoeliose. Rec. Mèd. Vèt. 153, 343, 1977.
67. Piergili Fioretti, D. et al. Il metodo ELISA per la ricerca di anticorpi in ovini parassitati da Dicrocoelium dendriticum. Rivista di Parassitologia. 3,289, 1980. 68. Savitskii, I.G., and Ponomareva, A.M. Diagnosis of ovine dicrocoeliasis by the ELISA. Veterinariya. 1, 43 (1984) (in Russian).
69. Gonzàlez-Lanza, C. et al. IgG antibody response to ES or somatic antigens of Dicrocoelium dendriticum (Trematoda) in experimentally infected sheep. Parasitol. Res. 86, 472, 2000.
70. Baldelli, B. et al. Dicroceliosi ovina: Studio degli anticorpi verso Dicrocoelium dendriticum in agnelli nati da pecore spontaneamente infette. Rivista di Parassitologia. 1, 23, 1981.
71. Ambrosi, M. et al. Dicroceliosi ovina: Insorgenza e decorso della infezione da Dicrocoelium dendriticum studiati con metodi parassitologici e sierologici (ELISA) in quattro gruppi di ovini di traccia. Rivista di Parassitologia. 3, 299, 1980. 72. Revilla-Nuín, B. et al. Partial characterization and isolation of 130 kDa antigenic protein of Dicrocoelium dendriticum adults. Vet. Parasitol. 134, 229, 2005. 73. Broglia, A. et al. Experimental ELISA for diagnosis of ovine dicrocoeliosis and application in a �eld survey. Parasitol. Res. 104, 949, 2009. 74. Campo, R. et al. Characterization of adult Dicrocoelium dendriticum by isoelectric focusing. J. Helminthol. 72, 109, 1998. 75. Sandoval, H. et al. Preliminary study on genetic variability of Dicrocoelium dendriticum determined by random ampli�ed polymorphic DNA. Parasitol. Int. 48, 21, 1999. 76. Morgan, J.A., and Blair, D. Nuclear rDNA ITS sequence variation in the trematode genus Echinostoma: An aid to establishing relationships within the 37-collar-spine group. Parasitology. 111, 609, 1995. 77. Jousson, O. et al. Use of the ITS rDNA for elucidation of some life-cycles of Mesometridae (Trematoda, Digenea). Int. J. Parasitol. 28, 1403, 1998. 78. Casey, S.P. et al. Use of ITS rDNA for discrimination of European green- and brown-banded sporocysts within the genus Leucochloridium Carus, 1835 (Digenea: Leucochloriidae). Syst. Parasitol. 56, 163, 2003. 79. Heussler, V. et al. A DNA probe for the detection of Dicrocoelium dendriticum in ants of Formica spp. and Lasius spp. Parasitol. Res. 84, 505, 1998. 80. Maurelli, M.P. et al. Characterization of the 28S and the second internal transcribed spacer of ribosomal DNA of Dicrocoelium dendriticum and Dicrocoelium hospes. Parasitol. Res. 101, 1251, 2007. 81. Otranto, D., and Traversa, D. Dicrocoeliosis of ruminants: A little known �uke disease. Trends Parasitol. 19, 12–15, 2003. 30 Chapter 30 - Echinostomes
6. Toledo, R., Muñoz-Antolí, C., and Esteban, J.G. The life-cycle of Echinostoma friedi n. sp. (Trematoda: Echinostomatidae) in Spain and a discussion on the relationships within the “revolutum” group based on cercarial chaetotaxy. Syst. Parasitol. 45, 199, 2000.
7. Kostadinova, A., Cercarial chaetotaxy of Echinostoma miyagawai Ishii, 1932 (Digenea: Echinostomatidae), with a review of the sensory patterns in the “revolutum” group. Syst. Parasitol. 44, 201, 1999.
8. Morgan, J.A.T., and Blair, D. Nuclear rDNA ITS sequence variation in the trematode genus Echinostoma: An aid to establishing relationships within the 37-collar-spine group. Parasitology 111, 609, 1995.
9. Morgan, J.A.T., and Blair, D. Mitochondrial ND1 gene sequence used to identify echinostome isolates from Australia and New Zealand. Int. J. Parasitol. 28, 493, 1998.
10. Morgan, J.A.T., and Blair, D. Relative merits of nuclear ribosomal internal transcribed spacers and mitochondrial CO1 and ND1 genes for distinguishing among Echinostoma species (Trematoda). Parasitology. 116, 289, 1998.
11. Morgan, J.A.T., and Blair, D. Molecular biology of echinostomes. In: Fried, B., and Gracyzk, T.K., (eds), Echinostomes as Experimental Models for Biological Research. Chapter 13. Kluwer Academic Publishers, Dordrecht, 2000.
12. Sorensen, R.E., Curtis, J., and Minchella, D.J. Intraspeci�c variation in the rDNA ITS loci of 37-collar-spined echinostomes from North America: Implications for sequencebased diagnoses and phylogenetics. J. Parasitol. 84, 992, 1998.
13. Maldonado, A. Jr. et al. Description of the adult worms of a new Brazilian isolate of Echinostoma paraensei (Platyhelminthes: Digenea) from its natural vertebrate host Nectomys squamipes by light and scanning electron microscopy and molecular analysis. Parasitol. Res. 87, 840, 2001.
14. Kostadinova, A. et al. Phylogenetic relationships of Echinostoma Rudolphi, 1809 (Digenea: Echinostomatidae) and related genera re-assessed via DNA and morphological analyses. Syst. Parasitol. 54, 159, 2003.
15. Grabda-Kazubska, B. et al. A phylogenetic analysis of trematodes of the genus Echinoparyphium and related genera based on sequencing of internal transcribed spacer region of rDNA. Acta Parasitol. 43, 116, 1998.
16. Huffman, J.E., and Fried, B. Echinostoma and echinostomiasis. Adv. Parasitol. 29, 215, 1990.
17. Haas, W. Snail-host �nding by miracidia and cercariae: Chemical host cues. Parasitol. Today 11, 468, 1995.
18. Haberl, B. Host-�nding in Echinostoma caproni: Miracidia and cercariae use different signals to identify the same snail species. Parasitology 120, 470, 2000.
19. Kanev, I. Life-cycle, delimitation and redescription of Echinostoma revolutum (Froelich, 1982) (Trematoda: Echinostomatidae). Syst. Parasitol. 28, 125, 1994.
20. Kanev, I. Redescription of Echinostoma trivolvis (Cort, 1914) (Trematoda: Echinostomatidae) with a discussion on its identity. Syst. Parasitol. 32, 61, 1995.
21. Muñoz-Antolí, C. et al. Infectivity of Echinostoma friedi miracidia to different snail species under experimental conditions. J. Helminthol. 80, 323, 2006.
22. Ataev, G.L. et al. Migration and development of mother sporocysts of Echinostoma caproni (Digenea: Echinostomatidae). J. Parasitol. 83, 444, 1997.
23. Haas, W. Physiological analyses of host-�nding behaviour in trematode cercariae: Adaptations for transmission success. Parasitology. 109, S15, 1994. 24. Haas, W. et al. Finding and recognition of the snail intermediate hosts by 3 species of echinostome cercariae. Parasitology 110,133, 1995. 25. Körner, M., and Haas, W. Chemo-orientation of echinostome cercariae towards their snail-hosts: Amino acids signal a low host-speci�city. Int. J. Parasitol. 28, 511, 1998. 26. Körner, M., and Haas, W. Chemo-orientation of echinostome cercariae towards their snail-hosts: The stimulating structure of amino acids and other attractans. Int. J. Parasitol. 28, 517, 1998. 27. Muñoz-Antolí, C. et al. Interactions related to non-host snails in the host-�nding process of Euparyphium albuferensis and Echinostoma friedi (Trematoda: Echinostomatidae) miracidia. Parasitol. Res. 91, 353, 2003. 28. Toledo, R., Muñoz-Antolí, C., and Fried, B. The use of echinostomes to study host–parasite relationships between larval trematodes and invertebrate and cold-blooded vertebrate hosts. Parasitol. Res. 100, 1177, 2007. 29. Fried, B., and Huffman, J.E. The biology of the intestinal trematode Echinostoma caproni. Adv. Parasitol. 38, 311, 1996. 30. Graczyk, T.K., and Fried, B. Echinostomiasis: A common but forgotten food-borne disease. Am. J. Trop. Med. Hyg. 58, 501, 1998. 31. Fried, B., Graczyk, T.K., and Tamang, L. Food-borne intestinal trematodiases in humans. Parasitol. Res. 93, 159, 2004. 32. Hosier, D.W., and Fried, B. Infectivity, growth, and distribution of Echinostoma revolutum (Trematoda) in Swiss webster and ICR mouse. Proc. Helminthol. Soc. Washington 53, 173, 1986. 33. Odaibo, A.B., Christensen, N.Ø., and Ukoli, F.M.A. Establishment, survival and fecundity in Echinostoma caproni (Trematoda) infections in NMRI mice. Proc. Helminthol. Soc. Washington 55, 265, 1988. 34. Odaibo, A.B., Christensen, N.Ø., and Ukoli, F.M.A. Further studies on the population regulation in Echinostoma caproni infections in NMRI mice. Proc. Helminthol. Soc. Washington 56, 192, 1989. 35. Christensen, N.Ø., Simonsen, P.E., Odaibo, A.B., and Mahler, H. Establishment, survival and fecundity of Echinostoma caproni (Trematoda) infections in hamsters and birds. J. Helminthol. Soc. Washington 57, 104, 1990. 36. Toledo, R. et al. The comparative development of Echinostoma caproni (Trematoda: Echinostomatidae) adults in experimentally infected hamsters and rats. Parasitol. Res. 93, 439, 2004. 37. Toledo, R. et al. An experimental study of the reproductive success of Echinostoma friedi (Trematoda: Echinostomatidae) in the golden hamster. Parasitology 126, 433, 2003. 38. Toledo, R. et al. A quantitative approach to the experimental transmission success of Echinostoma friedi (Trematoda: Echinostomatidae) in rats. J. Parasitol. 92, 16, 2006. 39. Carney, W.P. Echinostomiasis —A snail-borne intestinal disease. Southeast Asian J. Trop. Med. Public Health 22, 206, 1991. 40. Xiao, X. et al. In vivo and in vitro encystment of Echinochasmus liliputanus cercariae and biological activity of the metacercariae. J. Parasitol. 91, 492, 2005. 41. Keiser, J., and Utzinger, J. Chemotherapy for major foodborne trematodes: A review. Expert Opin. Pharmacother. 5, 1711, 2004. 42. Haaseb, M.A., and Eveland, L.K., Human echinostomiasis: Mechanisms of pathogenesis and host resistance. In: Fried, B., and Gracyzk, T.K. (eds), Echinostomes as Experimental Models for Biological Research. Chapter 4. Kluwer Academic Publishers, Dordrecht, 2000. 43. Chai, J.Y. Echinostomes in humans. In: Fried, B., and Toledo, R. (eds), The Biology of Echinostomes: From the Molecule to the Community. Chapter 7. Springer, New York, 2009.
44. Chai, J.Y., and Lee, S.H. Food-borne intestinal trematode infections in the Republic of Korea. Parasitol. Int. 51, 129, 2002.
45. Toledo, R., Esteban, J.G., and Fried, B. Immunology and pathology of intestinal trematodes in their de�nitive hosts. Adv. Parasitol. 63, 285, 2006.
46. Chattopadyay, U.K. et al. A case of echinostomiasis in tribal community in Bengal. Ann. Trop. Med. Parasitol. 84, 193, 1990.
47. Chai, J.Y. et al. A case of echinostomiasis with ulcerative lesions in the duodenum. Korean J. Parasitol. 32, 201, 1994.
48. Cho, C.M. et al. A human case of Echinostoma hortense (Trematoda: Echinostomatidae) infection diagnosed by gastroduodenal endoscopy in Korea. Korean J. Parasitol. 41, 117, 2003.
49. Lee, S.H. et al. Chronological observation of intestinal lesions of rats experimentally infected with Echinostoma hortense. Korean J. Parasitol. 28, 45, 1990. (In Korean).
50. Lee, S.K. et al. An epidemiological survey of Echinostoma hortense infection in Chongsong-gun, Kyongbuk Province. Korean J. Parasitol. 26, 199, 1988.
51. Lee, S.J., and Hong, S.J. Gastric echinostomiasis diagnosed by endoscopy. Gastrointest. Endosc. 55, 440, 2002.
52. Chang, Y.D. et al. A human infection of Echinostoma hortense in duodenal bulb diagnosed by endoscopy. Korean J. Parasitol. 43, 57, 2005.
53. Park, C.J., and Kim, J. A human case of Echinostoma hortense infection diagnosed by endoscopy in area of southwestern Korea. Korean J. Med. 71, 229, 2006. (In Korean).
54. Poland, G.A., Navin, T.R., and Sarosi, G.A. Outbreak of parasitic gastroenteritis among travelers returning from Africa. Arch. Intern. Med. 145, 2220, 1985.
55. Bindseil, E., and Christensen, N.Ø. Thymus-independent crypt hyperplasia and villous atrophy in the small intestine of mice infected with the trematode Echinostoma revolutum. Parasitology 88, 431, 1984.
56. Kim, S., and Fried, B. Pathological effects of Echinostoma caproni (Trematoda) in the domestic chick. J. Helminthol. 63, 227, 1989.
57. Toledo, R. et al. Echinostoma caproni: Intestinal pathology in the golden hamster, a highly compatible host, and the Wistar rat, a less compatible host. Exp. Parasitol. 112, 164, 2006.
58. Fujino, T., and Fried, B. Expulsion of Echinostoma trivolvis (Cort, 1914) Kanev, 1985 and retention of E. caproni Richard, 1964 (Trematoda, Echinostomatidae) in C3H mice: Pathological, ultrastructural, and cytochemical effects on the host intestine. Parasitol. Res. 79, 286, 1993.
59. Huffman, J.E., Iglesias, D., and Fried, B. Echinostoma revolutum: Pathology of extraintestinal infection in the golden hamster. Int. J. Parasitol. 18, 873, 1988.
60. Mabus, J., Huffman, J.E., and Fried, B. Humoral and cellular response to infection with Echinostoma revolutum in the golden hamster, Mesocricetus auratus. J. Helminthol.. 62, 127, 1988.
61. Isaacson, A.C., Huffman, J.E., and Fried, B. Infectivity, growth, development and pathology of Echinostoma caproni (Trematoda) in the golden hamster. Int. J. Parasitol. 19, 943, 1989.
62. Fried, B., Huffman, J.E., and Weiss, P.M. Single and multiple worm infections of Echinostoma caproni (Trematoda) in the golden hamster. J. Helminthol. 64, 75, 1990. 63. Fujino, T., and Fried, B. The expulsion of Echinostoma trivolvis from C3H mice: Differences in glycoconjugates in mouse versus hamster small intestinal mucosa during infection. J. Helminthol. 70, 115, 1996. 64. Weinstein, M.S., and Fried, B. The expulsion of Echinostoma trivolvis and retention of Echinostoma caproni in the ICR mouse: Pathological effects. Int. J. Parasitol. 21, 255, 1991. 65. Hamamoto, T. et al. A case of Echinostoma hortense infection diagnosed by the upper gastrointestinal endoscopy. Japan J. Gastroenterol. Soc. 94, 487, 1997. (in Japanese). 66. Simonsen, P.E., Estambale, B.B., and Agger, M. Antibodies in the serum of golden hamsters experimentally infected with the intestinal trematode Echinostoma caproni. J. Helminthol. 65, 239, 1991. 67. Agger, M.K., Simonsen, P.E., and Vennervald, B.J. The antibody response in serum, intestinal wall and intestinal lumen of NMRI mice infected with Echinostoma caproni. J. Helminthol. 67, 169, 1993. 68. Graczyk, T.K., and Fried, B. ELISA method for detecting anti-Echinostoma caproni (Trematoda: Echinostomatidae) immunoglobulins in experimentally infected ICR mice. J. Parasitol. 80, 544, 1994. 69. Graczyk, T.K., and Fried, B. An enzyme-linked immunosorbent assay for detecting anti-Echinostoma trivolvis (Trematoda) IgG in experimentally infected ICR mice. Crossreactivity with E. caproni. Parasitol. Res. 81, 710, 1995. 70. Toledo, R. et al. Immunological characterization of somatic and excretory/secretory antigens of Echinostoma caproni (Trematoda: Echinostomatidae) in experimentally infected rats. Comp. Parasitol. 71, 42, 2004. 71. Toledo, R. et al. Kinetics of antibodies and antigens in serum of mice experimentally infected with Echinostoma caproni (Trematoda: Echinostomatidae). J. Parasitol. 91, 978, 2005. 72 Cho, Y.K. et al. Differential immune pro�les following experimental Echinostoma hortense infection in BALB/c and C3H/ HeN mice. Parasitol. Res. 100, 1053, 2007. 73. Sotillo, J. et al. Echinostoma caproni: Kinetics of IgM, IgA and IgG subclasses in the serum and intestine of experimentally infected rats and mice. Exp. Parasitol. 116, 390, 2007. 74. Toledo, R. et al., Development of an antibody-based capture enzyme-linked immunosorbent assay for detecting Echinostoma caproni (Trematoda) in experimentally infected rats: Kinetics of coproantigen excretion. J. Parasitol. 89, 1227, 2003. 75. Toledo, R. et al. Kinetics of Echinostoma caproni (Trematoda: Echinostomatidae) antigens in feces and serum of experimentally infected hamsters and rats. J. Parasitol. 90, 752, 2004. 76. Saric, J. et al. Metabolic pro�ling of an Echinostoma caproni infection in the mouse for biomarker discovery. PLoS Negl. Trop. Dis. 2, e254, 2008. 77. Vasta, J.D., Fried, B., and Sherma, J. High performance thin layer chromatographic analysis of neutral lipids in the urine of BALB/c mice infected with Echinostoma caproni. Parasitol. Res. 102, 625, 2008. 78. Vasta, J.D., Fried, B., and Sherma, J. Determination and quanti�cation of amino acids in the urine of BALB/c mice infected with Echinostoma caproni by high-performance thin-layer chromatography-densitometry. J. Liq. Chrom. Relat. Tech. 32, 1210, 2009. 79. Voltz, A. et al. Isoenzyme analysis of Echinostoma liei: Comparison and hybridization with other African species. Exp. Parasitol. 66, 13, 1988. 80. Vasilev, I. et al. Comparative electrophoretic studies of certain species of the genus Echinostoma with 37-collar spines. Khelminthologia 6, 31, 1978.
81. Ross, G.C., Fried, B., and Southgate, V.R. Echinostoma revolutum and E. liei: Observations on enzymes and pigments. J. Nat. Hist. 23, 981, 1989.
82. Saijuntha, W., Sithithaworn, P., and Andrews, R.H. Genetic differentiation of Echinostoma revolutum and Hypodereaum conoideum from domestic ducks in Thailand by multilocus enzyme electrophoresis. J. Helminthol. 28, 1, 2009.
83. Fujino, T., Takahashi, Y., and Fried, B. A comparison of Echinostoma trivolvis and E. caproni using random ampli�ed polymorphic DNA analysis. J. Helminthol. 69, 263, 1995. 84. Fujino, T. et al. Speci�c primers for the detection of genomic DNA of Echinostoma trivolvis and E. caproni (Trematoda: Echinostomatidae). Mol. Cell. Probes 11, 77, 1997. 85. Petrie, J.L., Burg, E.F., 3rd, and Cain, G.D. Molecular characterization of Echinostoma caproni and E. paraensei by random ampli�cation of polymorphic DNA (RAPD) analysis. J. Parasitol. 82, 360, 1996. 86. Hertel, L.A., Stricker, S.A., and Loker, E.S. Calcium dynamics of hemocytes of the gastropod Biomphalaria glabrata: Effects of digenetic trematodes and selected bioactive compounds. Invert. Biol. 119, 27, 2000. 31 Chapter 31 - Fasciola
1. Mas-Coma, S., and Bargues, M.D. Human liver �ukes: A review. Res. Rev. Parasitol. 57, 145, 1997.
2. Mas-Coma, S., Esteban, J.G., and Bargues, M.D. Epidemiology of human fascioliasis: A review and proposed new classi�cation. Bull. WHO 77, 340, 1999.
3. Mas-Coma, S., Bargues, M.D., and Valero, M.A. Fascioliasis and other plant-borne trematode zoonoses. Int. J. Parasitol. 35, 1255, 2005.
4. Ali, H. et al. Genetic characterisation of Fasciola samples from different host species and geographical localities revealed the existence of F. hepatica and F. gigantica in Niger. Parasitol. Res. 102, 1021, 2008.
5. Ashra�, K. et al. Phenotypic analysis of adults of Fasciola hepatica, Fasciola gigantica and intermediate forms from the endemic region of Gilan, Iran. Parasitol. Int. 55, 249, 2006.
6. Lin, R.Q. et al. Sequence analysis of the �rst internal transcribed spacer of rDNA supports the existence of the intermediate Fasciola between F. hepatica and F. gigantica in mainland China. Parasitol. Res. 101, 813, 2007.
7. Le, T.H. et al. Human fascioliasis and the presence of hybrid/ introgressed forms of Fasciola hepatica and Fasciola gigantica in Vietnam. Int. J. Parasitol. 38, 725, 2008.
8. Hashimoto, K.T. et al. Mitochondrial DNA and nuclear DNA indicate that the Japanese Fasciola species is F. gigantica. Parasitol. Res. 83, 220, 1997.
9. Itagaki, T. et al. Taxonomic status of the Japanese triploid forms of Fasciola: Comparison of mitochondrial ND1 and COI sequences with F. hepatica and F. gigantica. J. Parasitol. 84, 445–448, 1998.
10. Itagaki, T. et al. Genetic characterization of parthenogenic Fasciola sp. in Japan on the basis of the sequences of ribosomal and mitochondrial DNA. Parasitology 131, 679, 2005.
11. Itagaki, T. et al. Molecular characterization of parthenogenic Fasciola sp. in Korea on the basis of DNA sequences of ribosomal ITS1 and mitochondrial NDI gene. J. Vet. Med. Sci. 67, 1115, 2005.
12. Prociv, P., Walker, J.C., and Whitby, M. Human ectopic fascioliasis in Australia: First case report. Med. J. Aust. 156, 349, 1992.
13. Walker, S.M. et al. The distribution of Fasciola hepatica and Fasciola gigantica within southern Tanzania—Constraints associated with the intermediate host. Parasitology 135, 495, 2008.
14. Esteban, J.G., Bargues, M.D., and Mas-Coma, S. Geographical distribution, diagnosis and treatment of human fascioliasis: A review. Res. Rev. Parasitol. 58, 13, 1998. 15. Haseeb, A.N. et al. A review on fascioliasis in Egypt. J. Egypt. Soc. Parasitol. 32, 317, 2002. 16. Nithiuthai, S. et al. Waterborne zoonotic helminthiases. Vet. Parasitol. 126, 67, 2004. 17. Zhou, P. et al. Food-borne parasitic zoonoses in China: Perspective for control. Trends Parasitol. 24, 190, 2008. 18. Keiser, J., and Utzinger, J. Food-borne trematodiases. Clin. Microbiol. Rev. 22, 466, 2009. 19. Gutiérrez, A. et al. Fasciola hepatica: Identi�cation of molecular markers for resistant and susceptible Pseudosuccinea columella snail hosts. Exp. Parasitol. 105, 211, 2003. 20. Bargues, M.D., and Mas-Coma, S. Reviewing lymnaeid vectors of fasciolosis by ribosomal DNA sequence analyses. J. Helminthol. 79, 257, 2005. 21. Kozak, M., and Wedrychowicz, H. The performance of a PCR assay for �eld studies on the prevalence of Fasciola hepatica infection in Galba truncatula intermediate host snails. Vet. Parasitol. 168, 25, 2010. 22. Goral, V. et al. A case of biliary fascioliasis by Fasciola gigantica in Turkey. Korean J. Parasitol. 49, 65, 2011. 23. Goral, V. et al. Biliary Fasciola gigantica case report from Turkey. Southeast Asian J. Trop. Med. Public Health 42, 509, 2011. 24. Arjona, R. et al. Fascioliasis in developed countries: A review of classic and aberrant forms of the diseases. Medicine (Baltimore). 74, 13, 1995. 25. Almendras-Jaramillo, M. et al. Hepatic fascioliasis in children: Uncommon clinical manifestations. Arch. Gastroenterol. 34, 241, 1997. 26. Chen, M.G., and Mott, K.E. Progress in assessment of morbidity due to Fasciola hepatica infection: A review of recent literature. Trop. Dis. Bull. 87, R1, 1990. 27. Dalimi, A., and Jabarvand, M. Fasciola hepatica in the human eye. Trans. R. Soc. Trop. Med. Hyg. 99, 798, 2005. 28. Zhou, L. et al. Multiple brain hemorrhages and hematomas associated with ectopic fascioliasis in brain and eye. Surg. Neurol. 69, 516, 2008. 29. Sezgın, O. et al. Fasciola hepatica-induced acute pancreatitis: Report of two cases and review of the literature. Turk. J. Gastroenterol. 21, 183, 2010. 30. Aslan, F. et al. A rare cause of cholangitis: Fasciola hepatica. Turk. J. Gastroenterol. 21, 329, 2010. 31. Acosta-Ferreira, W., Vercelli-Retta, J., and Falconi, L.M. Fasciola hepatica human infection. Histopathological study of sixteen cases. Virchows. Arch. A. Pathol. Anat. Histopathol. 383, 319, 1979. 32. Ishii, Y., Nakamura-Uchiyama, F., and Nawa, Y. A praziquantel- ineffective fascioliasis case successfully treated with triclabendazole. Parasitol. Int. 51, 205, 2002. 33. Carrada-Bravo, T. Fascioliasis: Diagnosis, epidemiology and treatment. Rev. Gastroenterol. Mex. 68, 135, 2003. 34. Kumar, P., Singh, V.K., and Singh, D.K. Combination of molluscicides with attractant carbohydrates and amino acids in bait formulation against the snail Lymnaea acuminata. Eur. Rev. Med. Pharmacol. Sci. 15, 550, 2011. 35. Kabaalioglu, A. et al. US-guided gallbladder aspiration: A new diagnostic method for biliary fascioliasis. Eur. Radiol. 9, 880, 1999. 36. Moustafa, N.E., Hegab, M.H., and Hassan, M.M. Role of ELISA in early detection of Fasciola copro-antigens in experimentally infected animals. J. Egypt. Soc. Parasitol. 28, 379, 1998. 37. O’Neill, S.M. et al. Immunodiagnosis of Fasciola hepatica (Fascioliasis) in a human population in the Bolivian Altiplano using puri�ed cathepsin-l-cysteine proteinase. Am. J. Trop. Med. Hyg. 58, 417, 1998.
38. Maleewong, W. et al. Fasciola gigantica-speci�c antigens: Puri�cation by a continuous-elution method and its evaluation for the diagnosis of human fascioliasis. Am. J. Trop. Med. Hyg. 61, 648, 1999.
39. Intapan, P.M. Development of rapid agglutination test using Fasciola gigantica speci�c antigen for serodiagnosis of human fascioliasis. Southeast Asian J. Trop. Med. Public Health 35, S313, 2004.
40. Tantrawatpan, C. et al. Serodiagnosis of human fascioliasis by a cystatin capture enzyme-linked immunosorbent assay with recombinant Fasciola gigantica cathepsin L antigen. Am. J. Trop. Med. Hyg. 72, 82, 2005.
41. Wongkham, C. et al. Evaluation of immunoglobulin G subclass antibodies against recombinant Fasciola gigantica cathepsin L1 in an enzyme-linked immunosorbent assay for serodiagnosis of human fasciolosis. Clin. Diagn. Lab. Immunol. 12, 1152, 2005.
42. Demerdash, Z.A. et al. Diagnostic ef�cacy of monoclonal antibody based sandwich enzyme linked immunosorbent assay (ELISA) for detection of Fasciola gigantica excretory/secretory antigens in both serum and stool. Parasit. Vectors 4, 176, 2011.
43. Rahimi, M.T. et al. Evaluation of Fast-ELISA versus standard-ELISA to diagnose human fasciolosis. Arch. Iran. Med. 14, 18, 2011.
44. Luton, K., Walker, D., and Blair, D. Comparisons of ribosomal internal transcribed spacers from two congeneric species of �ukes (Platyhelminthes: Trematoda: Digenea). Mol. Biochem. Parasitol. 56, 323, 1992.
45. Shubkin, C.D. et al. A nucleic acid-based test for the detection of Fasciola hepatica. J. Parasitol. 78, 817, 1992.
46. Heussler, V. et al. DNA probes for the detection of Fasciola hepatica in snails. Mol. Cell. Probes 7, 261, 1993.
47. Rognlie, M.C., Dimke, K.L., and Knapp, S.E. Detection of Fasciola hepatica in infected intermediate hosts using RT-PCR. J. Parasitol. 80, 748, 1994.
48. Rognlie, M.C. et al. Seasonal transmission of Fasciola hepatica in Montana. USA, with detection of infected intermediate hosts using a DNA-based assay. Vet. Parasitol. 65, 297, 1996.
49. Kaplan, R.M. et al. A repetitive DNA probe for the sensitive detection of Fasciola hepatica infected snails. Int. J. Parasitol. 25, 601, 1995.
50. Kaplan, R.M. et al. The prevalence of Fasciola hepatica in its snail intermediate host determined by DNA probe assay. Int. J. Parasitol. 27, 1585, 1997.
51. Agatsuma, T. et al. Molecular evidence of natural hybridization between Fasciola hepatica and F. gigantica. Parasitol. Int. 49, 231, 2000.
52. Ramadan, N.I., and Saber, L.M. Detection of genetic variability in nonhuman isolates of Fasciola hepatica and Fasciola gigantica by the RAPD-PCR technique. J. Egypt. Soc. Parasitol. 34, 679, 2004.
53. Abdel Tawab, A.H., el-Garhy, M.F., and Heckmann, R.A. Molecular polymorphisms associated with Egyptian, European, Chinese and North American Fasciola hepatica. J. Egypt. Soc. Parasitol. 36, 389, 2006.
54. Le, T.H. et al., Molecular con�rmation that Fasciola gigantica can undertake aberrant migrations in human hosts. J. Clin. Microbiol. 45, 648, 2007.
55. McGarry, J.W. et al. PCR-based differentiation of Fasciola species (Trematoda: Fasciolidae), using primers based on RAPD-derived sequences. Ann. Trop. Med. Parasitol. 101, 415, 2007. 56. Li, Q.Y. et al. Sequence-related ampli�ed polymorphism, an effective molecular approach for studying genetic variation in Fasciola spp. of human and animal health signi�cance. Electrophoresis 30, 403, 2009. 57. Ai, L. et al. Speci�c PCR-based assays for the identi�cation of Fasciola species: Their development, evaluation and potential usefulness in prevalence surveys. Ann. Trop. Med. Parasitol. 104, 65, 2010. 58. Ai, L. et al. Rapid identi�cation and differentiation of Fasciola hepatica and Fasciola gigantica by a loop-mediated isothermal ampli�cation (LAMP) assay. Vet. Parasitol. 174, 228, 2010. 59. Dar, Y. et al. Microsatellite analysis of Fasciola spp. in Egypt. Parasitol. Res. 109, 1741, 2011. 60. Prasad, P.K. et al. PCR-based molecular characterization and in silico analysis of food-borne trematode parasites Paragonimus westermani, Fasciolopsis buski and Fasciola gigantica from Northeast India using ITS2 rDNA. Bioinformation 6, 64, 2011. 61. Adlard, R.D. et al. Comparison of the second internal transcribed spacer (ribosomal DNA) from populations and species of Fasciolidae (Digenea). Int. J. Parasitol. 23, 423, 1993. 62. Mostafa, O.M., Taha, H.A., and Ramadan, G. Diagnosis of Fasciola gigantica in snail using the polymerase chain reaction (PCR) assay. J. Egypt. Soc. Parasitol. 33, 733, 2003. 63. Huang, W.Y. et al. Characterisation of Fasciola species from mainland China by ITS-2 ribosomal DNA sequence. Vet. Parasitol. 120, 75, 2004. 64. Alasaad, S. et al. Characterization of Fasciola samples from different host species and geographical localities in Spain by sequences of internal transcribed spacers of rDNA. Parasitol. Res. 101, 1245, 2007. 65. Capuano, F. et al., A comparison of �ve methods for DNA isolation from liver and rumen �ukes to perform ITS-2+ ampli�cation. Parasitologia 49, 27, 2007. 66. Ai, L. et al. Genetic diversity and relatedness of Fasciola spp. isolates from different hosts and geographic regions revealed by analysis of mitochondrial DNA sequences. Vet. Parasitol. 181, 329, 2011. 67. Amor, N. et al. Molecular characterization of Fasciola spp. from the endemic area of northern Iran based on nuclear ribosomal DNA sequences. Exp. Parasitol. 128, 196, 2011. 68. Amor, N. et al. Molecular characterization of Fasciola gigantica from Mauritania based on mitochondrial and nuclear ribosomal DNA sequences. Exp. Parasitol. 129, 127, 2011. 69. Ichikawa, M., and Itagaki, T. Discrimination of the ITS1 types of Fasciola spp. based on a PCR-RFLP method. Parasitol. Res. 106, 757, 2010. 70. Ichikawa, M. et al. Characterization of Fasciola spp. in Myanmar on the basis of spermatogenesis status and nuclear and mitochondrial DNA markers. Parasitol. Int. 60, 474, 2011. 71. Choe, S.E. et al. Genetic analysis of Fasciola isolates from cattle in Korea based on second internal transcribed spacer (ITS-2) sequence of nuclear ribosomal DNA. Parasitol. Res. 109, 833, 2011. 72. Marcilla, A., Bargues, M.D., and Mas-Coma, S. A PCR-RFLP assay for the distinction between Fasciola hepatica and F. gigantica. Mol. Cell. Probes 16, 327, 2002. 73. Rokni, M.B. et al. Identi�cation and differentiation of Fasciola hepatica and Fasciola gigantica using a simple PCR-restriction enzyme method. Exp. Parasitol. 124, 209, 2010.
74. Alasaad, S. et al. A TaqMan real-time PCR-based assay for the identi�cation of Fasciola spp. Vet. Parasitol. 179, 266, 2011.
75. Itagaki, T. et al. Characterization of genetic divergence among species within the genus Fasciola by PCR-SSCP. Jpn. J. Parasitol. 44, 244, 1995.
76. Alasaad, S. et al. A �uorescence-based polymerase chain reaction-linked single-strand conformation polymorphism (F-PCR-SSCP) assay for the identi�cation of Fasciola spp. Parasitol. Res. 108, 1513, 2011.
77. Magalhães, K.G., Passos, L.K., and Carvalho Odos, S. Detection of Lymnaea columella infection by Fasciola hepatica through multiplex-PCR. Mem. Inst. Oswaldo Cruz 99, 421, 2004.
78. Magalhães, K.G. et al. Isolation and detection of Fasciola hepatica DNA in Lymnaea viatrix from formalin-�xed and paraf�n-embedded tissues through multiplex-PCR. Vet. Parasitol. 152, 333, 2008.
79 El-Gozamy, B.R., and Shoukry, N.M. Identi�cation of Egyptian Fasciola species by PCR and restriction endonucleases digestion of the nuclear small subunit ribosomal RNA gene. J. Egypt. Soc. Parasitol. 39, 429, 2009. 80. Caron, Y., Lasri, S. and Losson, B. Fasciola hepatica: An assessment on the vectorial capacity of Radix labiata and R. balthica commonly found in Belgium. Vet. Parasitol. 149, 95, 2007.
81. Caron, Y., Rondelaud, D., and Losson, B. The detection and quanti�cation of a digenean infection in the snail host with special emphasis on Fasciola sp. Parasitol. Res. 103, 735, 2008. 82. Caron, Y. et al. An optimized DNA extraction and multiplex PCR for the detection of Fasciola sp. in lymnaeid snails. Vet. Parasitol. 178, 93, 2011. 83. Krämer, F., and Schnieder, T. Sequence heterogeneity in a repetitive DNA element of Fasciola. Int. J. Parasitol. 28, 1923, 1998. 84. Velusamy, R., Singh, B.P., and Raina, O.K. Detection of Fasciola gigantica infection in snails by polymerase chain reaction. Vet. Parasitol. 120, 85, 2004. 85. Cucher, M.A. et al. PCR diagnosis of Fasciola hepatica in �eld-collected Lymnaea columella and Lymnaea viatrix snails. Vet. Parasitol. 137, 74, 2006. 86. Schweizer, G. et al. Prevalence of Fasciola hepatica in the intermediate host Lymnaea truncatula detected by real time TaqMan PCR in populations from 70 Swiss farms with cattle husbandry. Vet. Parasitol. 150, 164, 2007. 87. Itagaki, T., and Tsutsumi, K. Triploid form of Fasciola in Japan: Genetic relationships between Fasciola hepatica and Fasciola gigantica determined by ITS-2 sequence of nuclear rDNA. Int. J. Parasitol. 28, 777, 1998. 88. Králová-Hromadová, I. et al. Sequence analysis of ribosomal and mitochondrial genes of the giant liver �uke Fascioloides magna (Trematoda: Fasciolidae): Intraspeci�c variation and differentiation from Fasciola hepatica. J. Parasitol. 94, 58, 2008. 32 Chapter 32 - Fasciolopsis
FIGURE 32.5 RNA secondary structures.
7. Jones, A. Family Fasciolidae Railliet, 1895. In: Jones, A., Bray, R.A., and Gibson, D. I. (eds), Keys to the Trematoda, Chapter 6. CABI Publishing and the Natural History Museum, Cambridge, pp. 79–85, 2005.
8. Kumar, V. The digenetic trematodes, Fasciolopsis buski, Gastrodiscoides hominis and Artyfechinostomum malayanum, as zoonotic infections in South Asian countries. Ann. Soc. Belg. Med. Trop. 60, 331, 1980.
9. Roy, B., and Tandon, V. Morphological and microtopographical strain variations among Fasciolopsis buski originating from different geographical areas. Acta Parasitol. 38, 72, 1993.
10. Gilman, R.H. et al. Endemic focus of Fasciolopsis buski in Bangladesh. Am. J. Trop. Med. Hyg. 31, 796, 1982.
11. Smyth, J.D. Animal Parasitology, Cambridge Low Price Editions, Cambridge University Press, Great Britain, p. 212, 1996.
12. Hsieh, H.C. Studies on the epidemiology of Fasciolopsis buski in South Taiwan. Formos. Sci. 14, 95, 1960.
13. Hsu, P.J. A survey of fasciolopsiasis of pigs in Kwantung province. Acta Vet. Zootech. Sin. 7, 143, 1964.
14. Wang, P. et al. Studies on the life-history of Fasciolopsis buski (Lankester, 1857) with consideration of its seasonal infection in pigs. Acta Zool. Sinica. 23, 88, 1977.
15. Nguen Tkhi Le. A study of the biology of the trematode Fasciolopsis buski (Lankester, 1857) in Vietnam. Med. Parazitol. (Moskva) 47, 81, 1978.
16. Tripathi, J.C., Srivastava, H.D., and Dutt, S.C. A note on experimental infection of Helicorbis coenosus and pig with Fasciolopsis buski. Indian J. Anim. Sci. 43, 647, 1973.
17. Buckley, J.J.C. Observations on Gastrodiscoides hominis and Fasciolopsis buski in Assam. J. Helminthol. 17, 1, 1939. 18. Malviya, H.C., and Varma, T.K. On the life history of Fasciolopsis buski. Ninth National Congress of Parasitology, Ujjain, January 29–31, p. 16, 1989.
19. Preet, S., and Prakash, S. Cercarial emergence in Fasciolopsis buski (Lankester). J. Parasit. Dis. 25, 108, 2001.
20. Manning, G.S., and Ratanarat, C. Fasciolopsis buski (Lankester, 1857) in Thailand. Am. J. Trop. Med. Hyg. 19, 613, 1970.
21. Ditrich, O. et al. Larval stages of medically important �ukes (Trematoda) from Vientiane province, Laos, Part II. Cercariae. Ann. Parasitol. Hum. Comp. 67, 75, 1992.
22. Graczyk, T.K. et al. Development of Fasciolopsis buski (Trematoda: Fasciolidae) in Hippeutis umbilicalis and Segmentina trochoideus (Gastropoda: Pulmonata) Parasitol. Res. 86, 324, 2000.
23. Barlow, C. The life cycle of human intestinal �uke Fasciolopsis buski (Lankester). Am. J. Hyg. Monoger. Ser. 4, 1, 1925.
24. Lo, C.T. Life history of the snail, Segmentina hemisphaerula (Benson), and its experimental infection with Fasciolopsis buski (Lankester). J. Parasitol. 53, 735, 1967.
25. Nguyen Van Tho. Evolution cycle of Fasciolopsis buski in the mollusc-intermediate host and in nature. Khoa Hoc Ky Thuat Thu Y (Vet. Sci. Tech.). 9, 43, 2002.
26. Weng, Y.L. et al. Studies on ecology of Fasciolopsis buski and control strategy of fasciolopsiasis. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi. 7, 108, 1989.
27. Nakagawa, K. On the life cycle of Fasciolopsis buski (Lankester). Kitasato Arch. Exp. Med. 4, 159, 1921.
28. Nakagawa, K. The development of Fasciolopsis buski Lankester. J. Parasitol. 8, 161, 1922.
29. Arambulo, P.V., and Moran, N., Food-transmitted parasitic zoonoses—socio-cultural and technological determinants. Int. J. Zoon. 7, 135, 1980. 30. Chandra, S.S. Epidemiology of Fasciolopsis buski in Uttar Pradesh. Indian J. Med. Res. 78, 55, 1984. 31. Jones, M.E.H., Hubbert, W.T., and Hagstad, H.V. Zoonoses— Recognition, Control and Prevention, 1st edn. Iowa State University Press, Ames, p. 245, 1995. 32. D’Souza, P.E., Jagannath, M.S., and Prasanna, K.M. A note on the occurrence of Fasciolopsis buski in pigs, Cheiron 30, 178, 2001. 33. Rim, H.J. Fasciolopsiasis. In: Hillyer, G.V., and Hopla, C.E. (eds), Handbook Series in Zoonoses, Vol. 3. CRC Press, Boca Raton, pp. 89–97, 1982. 34. Malviya, H.C. The susceptibility of mammals to Fasciolopsis buski. J. Helminthol. 59, 19, 1985. 35. Kuntz, R.E., and Lo, C.T. Preliminary studies on Fasciolopsis buski (Lankester, 1857) (giant Asian intestinal �uke) in the United States. Trans. Am. Microsc. Soc. 86, 163, 1967. 36. Hartman, H.A. The intestinal �uke (Fasciolopsis buski) in a monkey. Am. J. Vet. Res. 22, 1123, 1961. 37. Cross, J.H. Changing patterns of some trematode infections in Asia. Arzneimittelforschung 34, 1224, 1984. 38. Sadun, E.H., and Maiphoom, C. Studies on the epidemiology of the human intestinal �uke, Fasciolopsis buski (Lankester) in Central Thailand. Am. J. Trop. Med. Hyg. 2, 1070, 1953. 39. Muttalib, M.A., and Islam, N., Fasciolopsis buski in Bangladesh—A pilot study. J. Trop. Med. Hyg. 78, 135, 1975. 40. Yu, S. et al. Report of the �rst nationwide survey of the distribution of human parasites in China. 1. Regional distribution of parasitic species. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi. 12, 241, 1994. 41. World Health Organization, Foodborne trematode infections. Bull. World Health Organ., 73, 397, 1995. 42. World Health Organization, Control of foodborne trematode infections. World Health Organ. Tech. Rep. Ser. 849, 1, 1995. 43. Jaroonvesama, N. et al. Prevalence of Fasciolopsis buski and other parasitic infections in residents of three villages in Sena district, Ayudhaya Province, Thailand. J. Med. Assoc. Thai. 63, 493, 1980. 44. Shyu, L.Y., Lee, H.H., and Chen, E.R. A preliminary study on epidemiology of fasciolopsiasis in Tainan Hsien, south Taiwan. Chin. J. Microbiol. Immunol. 17, 118, 1984. 45. Thi, L.N., and Sey, O. Infectivity of metacercariae of Fasciolopsis buski (Lankester, 1857) and their development in de�nitive host. Parasitol. Hung. 18, 15, 1985. 46. Gai, L. et al. Relationships between the rate of parasitic infection and the knowledge of prevention. Chin. J. Parasitol. Parasit. Dis. 13, 269, 1995. 47. Lo, C.T., and Lee, K.M. Intestinal parasites among Southeast Asian labours in Taiwan during 1993–1994. Zhonghua Yi Xue Za Zhi (Taipei) 57, 401, 1996. 48. Xu., L. et al. Characteristics and recent trends in endemicity of human parasitic diseases in China. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi. 13, 214, 1995. 49. Li, G. et al. Relationship between the rate of parasitic infection and the knowledge of prevention. Chin. J. Parasitol. Parasit. Dis. 13, 269, 1995. 50. Longqi, X. et al. Characteristics and recent trends in endemicity of human parasitic diseases in China. Chin. J. Parasitol. Parasit. Dis. 13, 214, 1995. 51. Hotez, P.J. et al. Emerging and reemerging helminthiasis and the public health of China. Emerg. Infect. Dis. 3, 303, 1997. 52. Lee, H.H. Studies on epidemiology and treatment of fasciolopsiasis in southern Taiwan. Kaohsiung J. Med. Sci. 2, 21, 1986.
53. Lee, H.H., Shyu, L.Y., and Chen, E.R. Experimental control study of fasciolopsiasis in Taiwan. Kaohsiung J. Med. Sci. 5, 335, 1989.
54. Lee, H.H. Fasciolopsis buski infection among children of Liuying Primary School in Tainan Hsien south Taiwan. Chin. J. Microbiol. 5, 110, 1972.
55. Misra, S.C. et al. Incidence of gastrointestinal parasites in pigs with evaluation of an effective technique for the recovery of their ova or cysts in the faeces. Ind. Vet. J. 49, 140, 1972.
56. Malviya, H.C., and Varma, T.K. Incidence of Fasciolopsis buski infection in pigs and snails. Ninth National Congress of Parasitology, Ujjain, January 29–31, p. 15, 1989.
57. Roy, B., and Tandon, V. Trematodiasis in North East India: A study on the spectrum of digenetic trematodes among pigs, buffaloes, cattle, goats and sheep. Indian J. Anim. Health 31, 5, 1992.
58. Mohan, A., Mirdha, B.R., and Vishaka, M. Fascioliasis in a pregnant woman. Indian J. Gastroenterol. 15, 155, 1996.
59. Mahanta, J. Trematode infection of public health importance in Assam. Second Global Meet on Parasitic Diseases, Indian Society for Parasitology, Hyderabad, August 18–22, p. 72, 1997.
60. Shah, A., Gadgil, R.K., and Manohar, K.D. Fasciolopsiasis in Bombay. A preliminary communication. Indian J. Med. Sci. 20, 805, 1966.
61. Manjarumkar, P.V., and Shah, P.M. Epidemiological study of Fasciolopsis buski in Palghar Taluk. Indian J. Public Health. 16, 3, 1972. 62. Harinasuta, T., Bunnag, D., and Radomyos, P. Ef�cacy of praziquantel on fasciolopsiasis. Arzneimittelforschung 34, 1214, 1984.
63. Yoshihara, S. et al. Helminths and helminthiosis of pigs in the Mekong Delta, Vietnam with special reference to ascariosis and Fasciolopsis buski infection. Jpn. Agric. Res. Q. 33, 193, 1999.
64. Manning, G.S., Brockelman, W.Y., and Yiyant, V., An analysis of the prevalence of Fasciolopsis buski in central Thailand using catalytic models. Am. J. Epidemiol. 93, 354, 1971.
65. Bunnag, D., Radomyos, P., and Harinasuta, T. Field trial on the treatment of fasciolopsiasis with praziquantel. Southeast Asian J. Trop. Med. Public Health 14, 216, 1983.
66. Wiwanitkit, V., Suwansaksri, J., and Chaiyakhun, Y. High prevalence of Fasciolopsis buski in an endemic area of liver �uke infection in Thailand. Med. Gen. Med. 4, 6, 2002.
67. Waikagul, J. Intestinal �uke infections in Southeast Asia. Southeast Asian J. Trop. Med. Public Health 22, 158, 1991.
68. Giboda, M. et al. Current status of food-borne parasitic zoonoses in Laos. Southeast Asian J. Trop. Med. Public Health 22, 56, 1991.
69. Hadidjaja, P. et al. First autochthonous case of F. buski infection in Indonesia. Am. J. Trop. Med. Hyg. 31, 1065, 1982.
70. Margono, S.S. Important human helminthiasis in Indonesia. In: Crompton, D.W.T., Montresor, A., Nesheim, M.C., and Savioli, L. (eds), Controlling Disease due to Helminth Infections. World Health Organization, Geneva, p. 3, 2003.
71. Handoyo, I. et al. A survey of fasciolopsiasis in Sei Papuyu village of Babirik subdistrict, Hulu Sungei Utara Regency, South Kalimantan Province. Trop. Biomed. 3, 113, 1986.
72. Shekhar, K.C. Epidemiological assessment of parasitic zooonoses in Malaysia. Southeast Asian J. Trop. Med. Public Health 22, 337, 1991.
73. Basnuevo, J.G., and Seuc-Chiu, A. Fasciolopsis buski in Cuba; two cases treated with chloroquine, gentian violet, hexylresorcinol and tetrachloroethylene. Rev. Cubana. Med. Trop. Parasitol. 6, 91, 1950.
74. Greenberg, Z. et al. Prevalence of intestinal parasites among Thais in Israel. Harefuah 126, 507, 1994. 75. Schubert, S., and Granz, W. Problems in the differential diagnosis of intestinal diseases in patients who have traveled in the tropics. Z. Gesamte. Med. 36, 230, 1981. 76. Shah, P.M. et al. Fasciolopsis buski infestation in children. Indian Pediatr. 10, 721, 1973. 77. Rahman, K.M., Idris, M., and Khan, A.K.A. A study of fasciolopsiasis in Bangladesh. J. Trop. Med. Hyg. 84, 81, 1981. 78. Chandra, S.S. A �eld study on the clinical aspects of Fasciolopsis buski infections in Uttar Pradesh. Med. J. Armed Forces India 32, 181, 1976. 79. Jaroonvesama, N., Charoenlarp, K., and Areekul, S., Intestinal absorption studies in Fasciolopsis buski infection. Southeast Asian J. Trop. Med. Public Health 17, 582, 1986. 80. Viranuvatti, V., Stitinimankarn, T., and Tansurat, P. A fatal case of infection with Fasciolopsis buski in Thailand. Ann. Trop. Med. Parasitol. 47, 132, 1953. 81. Yu, S.H., and Mott, K.E., Epidemiology and morbidity of food-borne intestinal trematode infections. Trop. Dis. Bull. 91, R125, 1994. 82. Plaut, A.G. Kampanart-Sanyakorn, C., and Mannig, G.S. A clinical study of Fasciolopsis buski infection in Thailand. Trans. R. Soc. Trop. Med. Hyg. 63, 470, 1969. 83. Le, T.H. et al. Case report: Unusual presentation of Fasciolopsis buski in a Vietnamese child. Trans. R. Soc. Trop. Med. Hyg. 98, 193, 2004. 84. Blair, D. et al. Geographical genetic structure within the human lung �uke, Paragonimus westermani, detected from DNA sequences. Parasitology 115, 411, 1997. 85. Hillis, D.M., and Dixon, M.T. Ribosomal DNA: Molecular evolution and phylogenetic inference. Q. Rev. Biol. 66, 411, 1991. 86. Morgan, J.A.T., and Blair, D. Nuclear rDNA ITS sequence variation in the trematode genus Echinostoma: An aid to establishing relationships within the 37-collar spine group. Parasitology 111, 609, 1995. 87. Leon-Regagnon, V., Brooks, D.R., and Perez-Ponce, de Leon, G., Differentiation of Mexican species of Haematoloechus Looss, 1899 (Digenea: Plagiorchiformes): Molecular and morphological evidence. J. Parasitol. 85, 935, 1999. 88. Tkach, V.V., Pawlowski, J., and Sharpilo, V.P., Molecular and morphological differentiation between species of the Plagiorchis vespertilionis group (Digenea, Plagiorchiidae) occurring in European bats, with a re-description of P. vespertilionis group (Muller, 1780). Syst. Parasitol. 47, 9, 2000. 89. Kostadinova, A. et al. Phylogenetic relationships of Echinostoma rudolphi, 1809 (Digenea: Echinostomatidae) and related genera re-assessed via DNA and morphological analyses. Syst. Parasitol. 54, 159, 2003. 90. Scholz, T. et al. A taxonomic study of species of Bothriocephalus Rudolphi, 1808 (Cestoda: Pseudophyllidea) from eels in Japan: Morphological and molecular evidence for the occurrence of B. claviceps (Goeze, 1782) and con�rmation of the validity of B. japonicus Yamaguti, 1934. Syst. Parasitol. 57, 87, 2004. 91. Nolan, M.J., and Cribb, T.H. The use and implications of ribosomal DNA sequencing for the discrimination of digenean species. Adv. Parasitol. 60, 102, 2005. 92. Luton, K., Walker, D., and Blair, D. Comparisons of ribosomal internal transcribed spacers from two congeneric species of �ukes (Platyhelminths: Trematoda: Digenea). Mol. Biochem. Parasitol. 56, 323, 1992. 93. Adlard, R.D. et al. Comparison of the second internal transcribed spacer (ribosomal DNA) from populations and species of Fasciolidae (Digenea). Int. J. Parasitol. 23, 423, 1993.
94. Le, T.H. et al. Phylogenies inferred from mitochondrial gene orders—A cautionary tale from the parasitic �atworms. Mol. Biol. Evol. 17, 1123, 2000.
95. Prasad, P.K. et al. PCR-based determination of internal transcribed spacer (ITS) regions of ribosomal DNA of giant intestinal �uke. Fasciolopsis buski (Lankester, 1857) Looss, 1899. Parasitol. Res. 101, 1581, 2007.
96. Prasad, P.K. et al. Molecular identi�cation of Indian liver �uke, Fasciola (Trematoda: Fasciolidae) based on the ribosomal internal transcribed spacer regions. Parasitol. Res. 103, 1247, 2008.
97. Sambrooke, J., Fritsch, E.F., and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 1989.
98. Bowles, J., Blair, D., and McManus, D.P. A molecular phylogeny of the human schistosomes. Mol. Phylogenet. Evol. 4, 103, 1995.
99. White, B.A. PCR Protocols, Current Methods and Applications, Vol. 15. Humana, Totowa, NJ, 1993.
100. Blair, D., and Barker, S.C., Af�nities of the Gylianchenidae: Utility of the 18S rRNA gene for phylogenetic inference in the Digenea (platyhelminthes). Int. J. Parsitol. 23, 527, 1993.
101. Tamura, K. et al. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596, 2007.
102. Hillis, D.M., and Bull, J.J. An empirical test of bootstrapping as a method for assessing con�dence in phylogenetic analysis. Syst. Biol. 42, 182, 1993. 103. Huelsenbeck, J.P., and Ronquist, F. Bayesian inference of phylogenetic trees. Bioinformatics. 17, 754, 2001. 104. Bailey, T.L., and Elkan, C. Fitting a mixture model by expectation maximization to discover motifs in biopolymers. In: Proceedings of the Second International Conference on Intelligent Systems for Molecular Biology. AAAI Press, Menlo Park, CA, p. 28, 1994. 105. Thompson, R.C.A. et al. Advances in the diagnosis and systematics of parasites of veterinary importance: New and exciting prospects. In: Molecular systematics and diagnosis. Gasser, R.B. and Zarlenga, D.S. (eds), Vet. Parasitol. 125, 69,, 2004. 106. Blair, D. et al. A molecular perspective on the genera Paragonimus Braun, Euparagonimus Chen and Pagumogonimus Chen. J. Helminthol. 74, 295, 1999. 107. Iwagami, M. et al. Molecular phylogeographic studies on Paragonimus westermani in Asia. J. Helminthol. 74, 315, 2000. 108. Bhatti, H.S. et al. Fasciolopsiasis, a re-emerging infection in Azamgarh (Uttar Pradesh). Indian J. Pathol. Microbiol. 43, 73, 2000. 109. Muralidhar, S. et al. Fasciolopsiasis, a persisting problem in eastern U.P.: A case report. Indian J. Pathol. Microbiol. 43, 69, 2000. 110. Mott, K.E. et al. New geographical approaches to control of some parasitic zoonoses. Bull. World Health Org. 73, 247, 1995. 33 Chapter 33 - Gastrodiscoides
1. Jones, A., A comparative morphological study of the subfamily Gastrodiscinae Monticelli, 1892 (Paramphistomidae) with comments on some members of related subfamilies. J. Nat. Hist., 20, 863, 1986.
2. Brennan, G.P., Hanna, R.E., and Nizami, W.A., Ultrastructural and histochemical observations on the tegument of Gastrodiscoides hominis (Paramphistoma: Digenea). Int. J. Parasitol., 21, 897, 1991.
3. Tandon, V., and Maitra, S.C., Surface morphology of Gastrodiscoides hominis (Lewis & McConnell, 1876) Leiper, 1913 (Trematoda: Digenea) as revealed by scanning electron microscopy. J. Helminthol., 57, 339, 1983.
4. Dutt, S.C., and Srivastava, H.D., The life history of Gastrodiscoides hominis (Lewis and McConnel, 1876) Leiper, 1913, the amphistome parasite of man and pig. J. Helminthol., 46, 35, 1972.
5. Dutt, S.C., and Srivastava, H.D., The intermediate host and the cercaria of Gastrodiscoides hominis (Trematoda: Gastrodiscidae) preliminary report. J. Helminthol., 40, 45, 1966. 6. Shrivastav, H.O., and Shah, H.L., On Gastrodiscoides hominis (Lewis and McConnell, 1876) Leiper, 1913 from pigs (Sus scrofa domestica) in Madhya Pradesh, its pathology and public health importance. Indian J. Pathol. Bacteriol., 13, 68, 1970. 7. Chaisiri, K. et al., Human-dominated habitats and helminth parasitism in Southeast Asian murids. Parasitol. Res., 107, 931, 2010. 8. Kumar, V., The digenetic trematodes, Fasciolopsis buski, Gastrodiscoides hominis and Artyfechinostomum malayanum, as zoonotic infections in South Asian countries. Ann. Soc. Belg. Med. Trop., 60, 331, 1980. 9. Murty, C.V., and Reddy, C.R., A case report of Gastrodiscoides hominis infestation. Indian J. Pathol. Microbiol., 23, 303, 1980. 10. Dada-Adegbola, H.O. et al., Gastrodiscoides hominis infection in a Nigerian-case report. West Afr. J. Med., 23, 185, 2004. 11. Barker, S.C. et al., Utility of the D1 domain of nuclear 28S rRNA for phylogenetic inference in the Digenea. Syst. Parasitol., 26, 181, 1993. 12. Capuano, F. et al., A comparison of � ve methods for DNA isolation from liver and rumen �ukes to perform ITS-2+ ampli�cation. Parassitologia, 49, 27, 2007. 13. Lotfy, W.M. et al., A molecular approach for identi�cation of paramphistomes from Africa and Asia. Vet. Parasitol., 174, 234, 2010. 14. Goswami, L.M. et al., Molecular characterization of Gastrodiscoides hominis (Platyhelminthes: Trematoda: Digenea) inferred from ITS rDNA sequence analysis. Parasitol. Res., 104, 1485, 2009. 34 Chapter 34 - Haplorchis
TABLE 34.1
Summary of Molecular Detection Techniques for Heterophyid and Opisthorchiid Flukes
Detection Methods Species Studied PCR Target Source(s) of DNA Template Specificity/ Sensitivity Utility
Conventional PCR and
PCR-RFLP [42] Ht, Hp, Hy, Ov, and Cs Partial COI mtDNA Egg, MC, and AD of HIFs and OLFs High/medium Discriminating mixed infection
Conventional PCR [44, 46] Ht, Hp, Ov, and Cs ITS regions of rDNA Eggs, MC, AD, and fecal samples High/medium Revealing the utility of ITS1 and ITS2 for different purposes
Conventional PCR and
PCR-RFLP [47] Ht, Ov, and Cs ITS2 region of rDNA AD and fecal samples High/medium Detecting and discriminating the species in low mixed infections in fecal samples
Nested-PCR [48] Ht and Ov Partial COI mtDNA Fecal samples from purged individuals High/high Sensitive diagnostic tool for low-intensity Ht and Ov infections
HAT-RAPD, and conventional PCR [49] Ht and 13 other parasites The Ht speci�c HAT-RAPD fragment AD High/high Primer developed may be useful for detecting low Ht infections in endemic areas
Note: MC and AD refer to metacercaria and adult worm, respectively.
11. Sukontason, K. et al. Evidence of Haplorchis taichui infection as pathogenic parasite: Three case reports. Scand. J. Infect. Dis. 37, 388, 2005.
12. Giboda, M. et al. Current status of food-borne parasitic zoonoses in Laos. Southeast Asian J. Trop. Med. Public Health 22, S56, 1991.
13. Nithikathkul, C., and Wongsawad, C. Prevalence of Haplorchis taichui and Haplorchoides sp. metacercariae in freshwater �sh from water reservoirs, Chiang Mai, Thailand. Korean J. Parasitol. 46, 109, 2008.
14. Chai, J.Y. et al. High prevalence of liver and intestinal �uke infections among residents of Savannaket province in Laos. Korean J. Parasitol. 45, 213, 2007.
15. Chai, J.Y. et al. Mixed infections with Opisthorchis viverrini and intestinal �ukes in residents of Vientiane Municipality and Saravane Province in Laos. J. Helminthol. 79, 283, 2005.
16. Touch, S. et al. Discovery of Opisthorchis viverrini metacercariae in freshwater �sh in southern Cambodia. Acta Trop. 111, 108, 2009.
17. Ooi H.K. et al. Metacercariae in �shes of Sun Moon lake which is an endemic area for Clonorchis sinensis in Taiwan. Southeast Asian J. Trop. Med. Public Health 28, 222, 1997.
18. Wang, J.J. et al. Haplorchis infections in intermediate hosts from a clonorchiasis endemic area in Meinung, Taiwan, Republic of China. J. Helminthol. 76, 185, 2002.
19. Elsheikha H.M., and Elshazly A.M. Host-dependent variations in the seasonal prevalence and intensity of heterophyid encysted metacercariae (Digenea: Heterophyidae) in brackish water �sh in Egypt. Vet. Parasitol. 153, 65, 2008.
20. Rim, H.J. Opisthorchiasis. In: Steele J.H. (ed.), CRC Handbook Series in Zoonoses, Section C: Parasitic Zoonoses, Vol. III (Trematode Zoonoses). CRC Press, Boca Raton, FL, USA, p. 109, 1982.
21. Sukontason, K. et al. Prevalence of trematode metacercariae in cyprinoid �sh of Ban Pao district, Chiang Mai Province, northern Thailand. Southeast Asian J. Trop. Med. Public Health 30, 365, 1999.
22. Sukontason, K.L. et al. Prevalence of Opisthorchis viverrini infection among villagers harboring Opisthorchis-like eggs. Southeast Asian J. Trop. Med. Public Health 32, S23, 2001.
23. Kumchoo, K. et al. High prevalence of Haplorchis taichui metacercariae in cyprinoid �sh from Chiang Mai Province, Thailand. Southeast Asian J. Trop. Med. Public Health 36, 451, 2005.
24. Radomyos, P., Radomyos, B., and Tungtrongchitr, A. Multiinfection with helminthes in adults from northeast Thailand as determined by post-treatment fecal examination of adult worms. Trop. Med. Parasitol. 45, 133, 1994.
25. Waikagul, J. Opisthorchis viverrini metacercaria in Thai freshwater �sh. Southeast Asian J. Trop. Med. Public Health 29, 324, 1998.
26. Rim, H.J. et al. Fish-borne trematode metacercariae detected in freshwater �sh from Vientiane Municipality and Savannakhet Province, Lao PDR. Korean J. Parasitol. 46, 253, 2008.
27. De, N.V. et al. The food-borne trematode zoonoses of Vietnam. Southeast Asian J. Trop. Med. Public Health 34, 12, 2003.
28. Thu, N.D. et al. Survey for Zoonotic liver and intestinal trematode metacercariae in cultured and wild �sh in an Giang Province, Vietnam. Korean J. Parasitol. 45, 45, 2007.
29. Chi, T.T. et al. Prevalence of Zoonotic trematodes in �sh from a Vietnamese �sh-farming community. J. Parasitol. 94, 423, 2008.
30. Elshazly, A.M. et al. Natural infection of fresh and brackish water �sh with heterophyid encysted metacercariae in Dakahlia Governorate, Egypt. J. Egypt Soc. Parasitol. 37, 999, 2007. 31. Keiser, J., and Utznger, J. Food-borne trematodiases. Clin. Microbiol. Rev. 22, 466, 2009. 32. Chi, J.K. et al. Intestinal pathology in human metagonimiasis with ultrastructural observations of parasites, J. Korean Med. Sci. 3, 171, 1988. 33. Tantachumrun, T., and Kilks, M. Heterophyid infection in human ileum: Report of three cases. Southeast Asian J. Trop. Med. Public Health 9, 128, 1978. 34. Africa, C.M., de Leon, W.E.D., and Garcia, E.Y. Intestinal heterophyidiasis with cardiac involvement: A contribution to etiology of heart failure. J. Philippine Island Med. Assoc. 15, 358, 1935. 35. Africa, C.M., de Leon, W.E.D., and Garcia, E.Y. Visceral complications in intestinal heterophyidiasis of human. Acta Med. Philippines Monogr. Ser. 1, 1, 1940. 36. Fried, B., Graczyk, T.K., and Tamang, L. Food-borne intestinal trematodiasis in human. Parasiol. Res. 93, 159, 2004. 37. Chai, Y.Y., and Lee, S.H. Intestinal trematodes of humans in Korea: Metagonimus, heterophyids and echinostomes. Korean J. Parasitol. 28, 103, 1990. 38. Hotez, P.J. et al. Helminth infections: The great neglected tropical diseases. J. Clin. Investig. 118, 1311, 2008. 39. Ditrich, O. et al. Comparative morphology of eggs of the Haplorchiinae (Trematoda: Heterophyidae) and some other medically important heterophyid and opisthorchiid �ukes. Folia Parasitol. 39, 123, 1992. 40. Radomyos, B. et al. Opisthorchiasis and intestinal �uke infections in northern Thailand. Southeast Asian J. Trop. Med. Public Health 29, 123, 1998. 41. Tesana, S. et al. Eggshell morphology of the small eggs of human trematodes in Thailand. Southeast Asian J. Trop. Med. Public Health 22, 631, 1991. 42. Ditrich, O. et al. Serological differentiation of human small �uke infections using Opisthorchis viverrini and Haplorchis taichui antigens. Southeast Asian J. Trop. Med. Public Health 22 (Suppl), 174, 1991. 43. Thaenkham, U. et al. Discrimination of Opisthorchis viverrini from Haplorchis taichui using COI sequence marker. Acta Trop. 103, 26, 2007. 44. Sato, M. et al. Discrimination of O. viverrini, C. sinensis, H. pumilio and H. taichui using nuclear DNA-based PCR targeting ribosomal DNA ITS regions. Acta Trop. 109, 81, 2009. 45. Sato, M. et al. Egg laying capacity of H. taichui (Digenea: Heterophyidae) in humans. Korean J. Parasitol. 47, 3, 2009. 46. Sato, M. et al. Copro-DNA diagnosis of Opisthorchis viverrini and Haplorchis taichui infections in an endemic area of Laos. Southeast Asian J. Trop. Med. Public Health 41, 28, 2010. 47. Traub, R.J. et al. A new PCR-based approach indicates the range of Clonorchis sinensis now extend to central Thailand. PLos. Negl. Trop. Dis. 3, e367, 2009. 48. Lovis, L. et al. PCR diagnosis of Opisthorchis viverrini and Haplorchis taichui infections in a Lao community in an area of endemicity and comparison of diagnostic methods for parasitological �eld surveys. J. Clin. Microbiol. 47, 1517, 2009. 49. Wongsawad, C. et al. Haplorchis taichui, Witenberg, 1930: Development of a HAT-RAPD marker for the detection of minute intestinal �uke infection. Exp. Parasitol. 123 158, 2009. 50. Maniatis, T. et al. Molecular Cloning, 1st edn. Cold Spring Harbor Laboratory, New York, 458pp., 1982. 51. Thompson, J.D. et al. The clustal X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 25, 4876, 1997. 52. Hall, T.A. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser. 41, 95, 1999. 35 Chapter 35 - Heterophyidae
13. Park, J.H. et al. A new endemic focus of Heterophyes nocens and other heterophyid infections in a coastal area of Gangjingun, Jeollanam-do. Korean J. Parasitol. 45, 33–38, 2007.
14. Abou-Basha, L.M., Abdel-Fattah, M., Orecchia, P., Di Cave, D., and Zaki, A. Epidemiological study of heterophyiasis among humans in an area of Egypt. East Mediterr. Health J. 6, 932–938, 2000.
15. El Shazly, A.M. et al. Intestinal parasites in Dakahlia governorate, with different techniques in diagnosing protozoa. J. Egypt Soc. Parasitol. 36, 1023–1034, 2006.
16. Africa, C.M., de Leon, W., and Garcia, E.Y. Visceral complications in intestinal heterophyidiasis of man. Acta Medica Philippina, Monographic Series 1, 1–132, 1940.
17. Elsheikha, H.M. Heterophyosis: Risk of ectopic infection. Vet. Parasitol. 147, 341–342, 2007.
18. Sukontason, K., Piangjai, S., and Chaithong, U. Potassium permanganate staining for differentiation the surface morphology of Opisthorchis viverrini, Haplorchis taichui and Phaneropsolus bonnei eggs. Southeast Asian J. Trop. Med. Public Health 30, 371–374, 1999.
19. Elshazly, A.M., Elsheikha, H.M., Rahbar, M.H., and Awad, S.I. Comparison of three immunodiagnostic tests for experimental Heterophyes heterophyes infection in dogs. Vet. Parasitol. 151, 196–202, 2008.
20. Ryang, Y.S., Lee, C.Y., Lee, K.J., Lee, S.H., and Chai, J.Y. An incidental case of human Heterophyes nocens infection diagnosed by sectional morphology in a biopsy specimen of the small intestine. Korean J. Parasitol. 37, 189–194, 1999.
21. Dzikowski, R., Levy, M.G., Poore, M.F., Flowers, J.R., and Paperna, I. Use of rDNA polymorphism for identi�cation of Heterophyidae infecting freshwater �shes. Dis. Aquat. Organ. 59, 35–41, 2004.
22. Levy, M.G. et al. Morphologic, pathologic, and genetic investigations of Bolbophorus species affecting cultured channel cat�sh in the Mississippi delta. J. Aquat. Animal Health 14, 235–246, 2002. 23. Clark, T.K. Molecular approaches and techniques. In: Woo, P.T.K. (ed), Fish Diseases and Disorders, Vol. 1 Protozoan and Metazoan Infections, CAB International, Cambridge, MA, pp. 725–752, 2006.
24. Cunningham, C.O., and Mo, T.A. Random ampli�ed polymorphic DNA (RAPD) analysis of three Norwegian Gyrodactylus salaris populations (Monogenea: Gyrodactylidae). J. Parasitol. 83, 311–314, 1997.
25. Martin-Sanchez, J., Diaz, M., Artacho, M.E., and Valero, A. Molecular arguments for considering Hysterothylacium fabri (Nematoda: Anisakidae) a complex of sibling species. Parasitol. Res. 89, 214–220, 2003.
26. Dzikowski, R., Levy, M.G., Poore, M.F., Flowers, J.R., and Paperna, I. Clinostomum complanatum and Clinostomum marginatum (Rudolphi, 1819) (Digenea: Clinostomidae) are separate species based on differences in ribosomal DNA. J. Parasitol. 90, 413–414, 2004.
27. Dzikowski, R., Levy, M.G., Poore, M.F., Flowers, J.R., and Paperna, I. Genetic and morphologic differentiation of Bolbophorus confusus and B-levantinus (Digenea: Diplostomatidae), based on rDNA SSU polymorphism and SEM. Diseases of Aquatic Organisms 57, 231–235, 2003.
28. Cribb, T.H.B., Bray, R.A., Littlewood, D.T.J., Pichelin, S.P., Herniou, E.A. The Digenea. In: Littlewood, D.T.J., and Bray, R.A. (eds), Interrelationship of the Platyhelminthes. Taylor & Francis Inc., London, pp. 168–185, 2001. 29. Cribb, T.H., Anderson, G.R., Adlard, R.D., and Bray, R.A. A DNA-based demonstration of a three-host life-cycle for the Bivesiculidae (Platyhelminthes: Digenea). Int. J. Parasitol. 28, 1791–1795, 1998. 30. Anderson, G.R. Identi�cation and maturation of the metacercaria of Indodidymozoon pearsoni. J. Helminthol. 73, 21–26, 1999. 31. Jousson, O., Bartoli, P., Zaninetti, L., and Pawlowski, J. Use of the ITS rDNA for elucidation of some life-cycles of Mesometridae (Trematoda, Digenea). Int. J. Parasitol. 28, 1403–1411, 1998. 32. Bartoli, P., Jousson, O., and Russell-Pinto, F. The life cycle of Monorchis parvus (Digenea: Monorchiidae) demonstrated by developmental and molecular data. J. Parasitol. 86, 479–489, 2000. 33. Jousson, O., Bartoli, P., and Pawlowski, J. Molecular identi�cation of developmental stages in Opecoelidae (Digenea). Int. J. Parasitol. 29, 1853–1858, 1999. 34. Jousson, O., and Bartoli, P. The life cycle of Opecoeloides columbellae (Pagenstecher, 1863) n. comb. (Digenea, opecoelidae): Evidence from molecules and morphology. Int. J. Parasitol. 30, 747–760, 2000. 35. Jousson, O., and Bartoli, P. Molecules, morphology and morphometrics of Cainocreadium labracis and Cainocreadium dentecis n. sp. (Digenea: Opecoelidae) parasitic in marine �shes. Int. J. Parasitol. 31, 706–714, 2001. 36. Sorensen, R.E., Curtis, J., and Minchella, D.J. Intraspeci�c variation in the rDNA its loci of 37-collar-spined echinostomes from North America: Implications for sequence-based diagnoses and phylogenetics. J. Parasitol. 84, 992–997, 1998. 37. van Herwerden, L., Blair, D., and Agatsuma, T. Intra- and interindividual variation in ITS1 of paragonimus westermani (Trematoda: Digenea) and related species: Implications for phylogenetic studies. Mol. Phylogenet. Evol. 12, 67–73, 1999. 38. Miura, O. et al. Molecular-genetic analyses reveal cryptic species of trematodes in the intertidal gastropod, Batillaria cumingi (Crosse). Int. J. Parasitol. 35, 793–801, 2005. 39. Miura, O., Torchin, M.E., Kuris, A.M., Hechinger, R.F., and Chiba, S. Introduced cryptic species of parasites exhibit different invasion pathways. Proc. Natl. Acad. Sci. USA 103, 19,818–19,823, 2006. 40. Traub, R.J. et al. A new PCR-based approach indicates the range of Clonorchis sinensis now extends to central Thailand. PLoS Negl. Trop. Dis. 3, e367, 2009. 41. Sato, M., Thaenkham, U., Dekumyoy, P., and Waikagul, J. Discrimination of O. viverrini, C. sinensis, H. pumilio and H. taichui using nuclear DNA-based PCR targeting ribosomal DNA ITS regions. Acta Tropica. 109, 81–83, 2009. 42. Wongsawad, C., Wongsawad, P., Chai, J.Y., and Anuntalabhochai, S. Haplorchis taichui, Witenberg, 1930: Development of a HAT-RAPD marker for the detection of minute intestinal �uke infection. Exp. Parasitol. 123, 158, 2009. 43. Favis, R., and Barany, F. Mutation detection in K-ras, BRCA1, BRCA2, and p53 using PCR/LDR and a universal DNA microarray. Ann. N Y Acad. Sci. 906, 39–43, 2000. 44. Liew, M. et al. Genotyping of single-nucleotide polymorphisms by high-resolution melting of small amplicons. Clin. Chem. 50, 1156–1164, 2004. 36 Chapter 36 - Metagonimus
9. Saito, S., and Shimizu, T. A new trematode, Metagonimus otsurui sp. nov. from the fresh-water �shes (Trematoda: Heterophyidae). Jpn. J. Parasitol. 17, 167, 1968.
10. Chai, J.Y., Murrell, K.D., and Lymbery, A.J. Fish-borne parasitic zoonoses: Status and issues. Int. J. Parasitol. 35, 1233, 2005.
11. Yu, S.H., and Mott, K.E. Epidemiology and morbidity of food-borne intestinal trematode infections. Trop. Dis. Bull. 91, R125, 1994.
12. Chai, J.Y. et al. An epidemiological study of metagonimiasis along the upper reaches of the Namhan River. Korean J. Parasitol. 31, 99, 1993.
13. Chai, J.Y. et al. An egg-negative patient of acute metagonimiasis diagnosed serologically by ELISA. Seoul. J. Med. 30, 139, 1989.
14. Yang, H.J. et al. Molecular differentiation of three species of Metagonimus by simple sequence repeat anchored polymerase chain reaction (SSR-PCR) ampli�cation. J. Parasitol. 86, 1170, 2000.
15. Yu, J.R., and Chai, J.Y. Metagonimus. In: Liu, D. (ed), Molecular Detection of Foodborne Pathogens. CRC Press (Taylor & Francis Group), Boca Raton, FL, 2010.
16. Katsurada, F. On a new trematode. Metagonimus yokogawai, Tokyo Iji Shinshi 1796, 3483, 1912. (in Japanese).
17. Kobayashi, H. On a new genus of the trematode (a preliminary report). Saikingaku Zasshi 204, 780, 1912. (in Japanese).
18. Chai, J.Y. et al. Chronological observation on intestinal histopathology and intraepithelial lymphocytes in the intestine of rats infected with Metagonimus yokogawai. Korean J. Parasitol. 32, 215, 1994. (in Korean).
19. Cho, S.Y., Kang, S.Y., and Lee, J.B. Metagonimiasis in Korea. Arzneimittelforschung 34(9B), 1211, 1984.
20. Ahn, Y.K. Lateolaborax japonicus, a role of the second intermediate host of Metagonimus yokogawai. New Med. J. 26, 135, 1983. (in Korean).
22. Cho, S.Y., Kang, S.Y., and Ryang, Y.S. Helminthes infections in the small intestine of stray dogs in Ejungbu City, Kyunggi Do, Korea. Korean J. Parasitol. 19, 55, 1981.
23. Seo, B.S. et al. Studies on parasitic helminths of Korea 5. Survey on intestinal trematodes of house rats. Korean J. Parasitol. 19, 131, 1981.
24. Huh, S., Sohn, W.M., and Chai, J.Y. Intestinal parasites of cats purchased in Seoul. Korean J. Parasitol, 31, 371, 1993.
25. Miyamoto, K. Studies on zoonoses in Hokkaido 7. Survey of natural de�nitive hosts of Metagonimus yokogawai. Jpn. J. Parasitol. 34, 371, 1985. (in Japanese).
26. Ahn, Y.K. Epidemiological studies on Metagonimus yokogawai infection in Samcheok-gun, Kangwon-do, Korea. Korean J. Parasitol. 22, 161, 1984. (in Korean).
27. Chai, J.Y. et al. High endemicity of Metagonimus yokogawai infection along residents of Samcheok-shi, Kangwon-do. Korean J. Parasitol. 38, 33, 2000.
28. Lee, J.J. et al. Decrease of Metagonimus yokogawai endemicity along the Tamjin River basin. Korean J. Parasitol. 46, 289, 2008.
29. Ito, J. et al. On the prevalence of Metagonimus sp. among the inhabitants at Hamamatsu basin in Shizuoka prefecture, Japan. Jpn. J. Parasitol. 40, 274, 1991.
30. Takahashi, S. On the life history of Metagonimus yokogawai, a new species of Metagonimus, and Exorchis major, Okayama Igakkai Zasshi 41, 2687, 1929. (in Japanese). 31. Asada, S. On Metagonimus yokogawai and its related species. Rinsho Igaku (Clinical Medicine) 22, 43, 1934. (in Japanese). 32. Saito, S. On the differences between Metagonimus yokogawai and Metagonimus takahashii II. The morphological comparisons. Jpn. J. Parasitol. 21, 449, 1972. (in Japanese). 33. Saito, S. On the differences between Metagonimus yokogawai and Metagonimus takahashii II. The experimental infections to the second intermediate hosts. Jpn. J. Parasitol. 22, 39, 1973. (in Japanese). 34. Koga, G. Studies on the genus Metagonimus. Igaku Kenkyu 12, 3471, 1938. (in Japanese). 35. Chai, J.Y. et al. Three morphological types of the genus Metagonimus encysted in the dace, Tribolodon taczanowskii, caught from the Sumjin River. Korean J. Parasitol. 29, 217, 1991. 36. Yamaguti, S. Digenetic trematodes of vertebrates. Systema Helminthum 1, 716, 1958. 37. Kamiya et al. Helminths of stray dogs in Sapporo. II. Jpn. J. Parasitol. 24, 41, 1975. (in Japanese). 38. Ahn, Y.K. Intestinal �ukes of genus Metagonimus and their second intermediate hosts in Kangwon-do, Korean J. Parasitol. 31, 331, 1993. (in Korean). 39. Guk, S.M. et al. Susceptibility of inbred mouse strains to infection with three species of Metagonimus prevalent in the Republic of Korea. J. Parasitol. 91, 12, 2005. 40. Chun, S.K. A study on the metacercaria of Metagonimus takahashii and Exorchis oviformis from Carassius carassius. Bull. Pusan Fish. Coll. 3, 31, 1960. (in Korean). 41. Ahn, Y.K., and Ryang, Y.S. Epidemiological studies on Metagonimus infection along the Hongcheon river, Kangwon Province. Korean J. Parasitol. 26, 207, 1988. (in Korean). 42. Kim, K.H. et al. A numerical taxonomic study on heterophyid trematodes. Korean J. Parasitol. 29, 55, 1991. (in Korean). 43. Yu, J.R. et al. PCR-RFLP pattern of three kinds of Metagonimus in Korea. Korean J. Parasitol. 35, 271, 1997. 44. Lee, S.U. et al. A cytogenetic study on human intestinal trematodes of the genus Metagonimus (Digenea: Heterophyidae) in Korea. Korean J. Parasitol. 37, 237, 1999. 45. Kim, C.H. et al. Studies on the Metagonimus �uke in the Daecheong reservoir and the upper stream of Geum River, Korea. Korean J. Parasitol. 25, 69, 1987. (in Korean). 46. Shimazu, T. Life cycle and morphology of Metagonimus miyatai (Digenea: Heterohpyidae) from Nagano, Japan. Parasitol. Int. 51, 271, 2002. 47. Chai, J.Y., and Lee, S.H. Food-borne intestinal trematode infections in the Republic of Korea. Parasitol. Int. 51, 129, 2002. 48. Kim, C.H. Study on the Metagonimus sp. in Gum River basin, Chungchung-nam Do, Korea. Korean J. Parasitol. 18, 215, 1980. (in Korean). 49. Ahn, Y.K., and Ryang, Y.S. Growth, egg-laying and life span of Miyata type of genus Metagonimus (Trematoda: Heterophyidae) in the �nal hosts. J. Wonju Med. Coll. 8, 1, 1995. (in Korean). 50. Park, M.S. et al. Intestinal parasite infections in the inhabitants along the Hantan River, Chorwon-gun. Korean J. Parasitol. 31, 375, 1993. 51. Yu, J.R., Kwon, S.O., and Lee, S.H. Clonorchiasis and metagonimiasis in the inhabitants along Talchongang (River), Chungwon-gun. Korean J. Parasitol. 32, 267, 1994. 52. Kurokawa, T. Studies on the genus Metagonimus, especially on the life history of M. katsuradai Izumi, 1935 and determination of its �rst intermediate host. Tokyo Iji Shinshi 3161, 2877, 1939. (in Japanese). 53. Besprozvannykh, V.V. et al. Occurrence of Metagonimus katsuradai Izumi, 1935 (Trematoda: Heterophyidae) in the southern Primor’e region. In: Yu, L. (ed), Gel’minty I vyzyvaemye imi zabollevaniya, Mamaev, Vladivostock, Russia, 1987.
54. Chai, J.Y. Study on Metagonimus yokogawai (Katsurada, 1912) in Korea V. Intestinal pathology in experimentally infected albino rats. Seoul J. Med. 20, 104, 1979.
55. Lee, J.B. et al. Study on the pathology of metagonimiasis in experimentally infected cat intestine. Korean J. Parasitol. 19, 109, 1981.
56. Kang, S.Y. et al. A study on intestinal lesions of experimentally reinfected dogs with Metagonimus yokogawai. Korean J. Parasitol. 21, 58, 1983.
57. Chai, J.Y., Kim, J., and Lee, S.H. Invasion of Metagonimus yokogawai into the submucosal layer of the small intestine of immunosuppressed mice. Korean J. Parasitol. 33, 313, 1995.
58. Chai, J.Y., Seo, B.S., and Lee, S.H. Study on Metagonimus yokogawai (Katsurada, 1912) in Korea VII. Susceptibility of various strains of mice to Metagonimus infection and effect of prednisolone. Korean J. Parasitol. 22, 153, 1984.
59. Yu, J.R., Myong, N., and Chai, J.Y. Expression patterns of proliferating cell nuclear antigen in the small intes tine of mice infected with Metagonimus yokogawai and Metagonimus Miyata type. Korean J. Parasitol. 35, 239, 1997. 60. Yamada et al. A case of metagonimiasis complicated with multiple intracerebral hemorrhages and diabetes mellitus. J. Nippon Med. Sch. 75, 32–35, 2008. 61. Rim, H.J. et al. Anthelmintic effects of various drugs against metagonimiasis. Korean J. Parasitol. 16, 117, 1978. (in Korean). 62. Chai, J.Y. et al. Effects of gamma-irradiation on the survival and development of Metagonimus yokogawai metacercariae in rats. Korean J. Parasitol. 33, 297, 1995. 63. Lee, S.H. et al. Comparative morphology of eggs of heterophyids and Clonorchis sinensis causing human infections in Korea. Korean J. Parasitol. 22, 171, 1984. 64. Yu, J.R., Chung, J.S., and Chai, J.Y. Different RAPD patterns between Metagonimus yokogawai and Metagonimus Miyata type. Korean J. Parasitol. 35, 295, 1997. 65. Lee, S.U. et al. Sequence comparisons of 28S ribosomal DNA and mitochondrial cytochrome c oxidase subunit I of Metagonimus yokogawai, M. takahashii and M. miyatai. Korean J. Parasitol. 42, 129, 2004. 37 Chapter 37 - Nanophyetus
11. Dumler, J.S. et al., Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: Uni�cation of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and ‘HE agent’ as subjective synonyms of Ehrlichia phagocytophila. Int. J. Syst. Evol. Microbiol., 51, 2145, 2001.
12. Eastburn, R.L. et al., Praziquantel for treatment of human Nanophyetus salmincola (Troglotrema salmincola) infection. J. Infect. Dis., 160, 896, 1989.
13. Fritsche, T.R. et al., Praziquantel for treatment of human Nanophyetus salmincola (Troglotrema salmincola) infection. J. Infect. Dis., 160, 896, 1989. 14. Foreyt, W.J., and Gorham, J.R., Evaluation of praziquantel against induced Nanophyetus salmincola infections in coyotes and dogs. Am. J. Vet. Res., 49, 563, 1988. 15. Johansen, M.V. et al., Towards improved diagnosis of zoonotic trematode infections in Southeast Asia. Adv. Parasitol., 73, 171, 2010. 16. Amor, N. et al., Molecular characterization of Fasciola gigantica from Mauritania based on mitochondrial and nuclear ribosomal DNA sequences. Exp. Parasitol., 129, 127, 2011. 17. Criscione, C.D., and Blouin, M.S., Life cycles shape parasite evolution: Comparative population genetics of salmon trematodes. Evolution, 58, 198, 2004. 38 Chapter 38 - Opisthorchis
1. Hotez, P.J. et al. Rescuing the bottom billion through control of neglected tropical diseases. Lancet 373, 1570, 2009.
2. Chai, J.Y. et al. Fish-borne parasitic zoonoses: Status and issues. Int. J. Parasitol. 35, 1233, 2005.
3. Liu, W.Q. et al. Comparison of ancient and modern Clonorchis sinensis based on ITS1 and ITS2 sequences. Acta Trop. 101, 91, 2007.
4. Kaewkes, S. Taxonomy and biology of liver �ukes. Acta Trop. 88, 177, 2003.
5. Beaver, P.C., Jung, R.C., and Cupp, E.W. Clinical Parasitology, 9th edn. Lea & Febiger, Philadelphia, 406pp., 1984.
6. Sithithaworn, P. et al. Relationship between faecal egg count and worm burden of Opisthorchis viverrini in human autopsy cases. Parasitology 102(Part 2), 277, 1991.
7. Laoprom, N. et al. Microsatellite loci in the carcinogenic liver �uke, Opisthorchis viverrini and their application as population genetic markers. Infect. Genet. Evol. 10, 146, 2010.
8. Saijuntha, W. et al. Evidence of a species complex within the food-borne trematode Opisthorchis viverrini and possible coevolution with their �rst intermediate hosts. Int. J. Parasitol. 37, 695, 2007.
9. Kaewkes, S. et al. Comparative studies on the morphology of the eggs of Opisthorchis viverrini and lecithodendriid trematodes. Southeast Asian J. Trop. Med. Public Health 22, 623, 1991.
10. Ditrich, O., Giboda, M., Scholz, T., and Beer, S.A. Comparative morphology of eggs of the Haplorchiinae (Trematoda: Heterophyidae) and some other medically important heterophyid and opisthorchid �ukes. Folia Parasitol. 39, 123, 1992.
11. Elkins, D.B. et al. Opisthorchis viverrini: Relationships between egg counts, worms recovered and antibody levels within an endemic community in Northeast Thailand. Parasitology 102(Part 2), 283, 1991. 12. Phongsasakulchoti, P. et al. Emergence of Opisthorchis viverrini cercariae from naturally infected Bithynia (Digoniostoma) siamensis goniomphalos. Southeast Asian J. Trop. Med. Public Health 36(Suppl 4), 189, 2005.
13. Attwood, H.D., and Chou, S.T. The longevity of Clonorchis sinensis. Pathology 10, 153, 1978.
14. Sithithaworn, P. et al. Genetic variation in Opisthorchis viverrini (Trematoda: Opisthorchiidae) from Northeast Thailand and Laos PDR based on random ampli�ed polymorphic DNA analyses. Parasitol. Res. 100, 613, 2007. 15. Laoprom, N. et al. Biological variation within Opisthorchis viverrini sensu lato in Thailand and Lao PDR. J. Parasitol. 95, 1307, 2009. 16. He, Y.X. et al. Strain complex of Schistosoma japonicum in the mainland of China. Southeast Asian J. Trop. Med. Public Health 25, 232, 1994. 17. He, Y.X. et al. Characteristics of different isolates of Schistosoma japonicum from China in the �nal hosts. Southeast Asian J. Trop. Med. Public Health 22, 240, 1991. 18. Park, G.M. Genetic comparison of liver �ukes, Clonorchis sinensis and Opisthorchis viverrini, based on rDNA and mtDNA gene sequences. Parasitol. Res. 100, 351, 2007. 19. Lee, S.U. and Huh, S. Variation of nuclear and mitochondrial DNAs in Korean and Chinese isolates of Clonorchis sinensis. Korean J. Parasitol. 42, 145, 2004. 20. Shekhovtsov, S.V. et al. A novel nuclear marker, Pm-int9, for phylogenetic studies of Opisthorchis felineus, Opisthorchis viverrini, and Clonorchis sinensis (Opisthorchiidae, Trematoda). Parasitol. Res., 106, 293, 2009. 21. Brockelman, W.Y. et al. Field studies on the transmission of the human liver �uke, Opisthorchis viverrini, in northeast Thailand: Population changes of the snail intermediate host. Int. J. Parasitol. 16, 545, 1986. 22. Vichasri, S. et al. Opisthorchis viverrini: Intensity and rates of infection in cyprinoid �sh from an endemic focus in Northeast Thailand. Southeast Asian J. Trop. Med. Public Health 13, 138, 1982. 23. Haas, W. et al. Opisthorchis viverrini: Finding and recognition of the �sh host by the cercariae. Exp. Parasitol. 71, 422, 1990. 24. Sithithaworn, P. et al. Seasonal variation of Opisthorchis viverrini infection in cyprinoid �sh in north-east Thailand: Implications for parasite control and food safety. Bull. World Health Organ 75, 125, 1997. 25. Hinz, E. et al. Opisthorchiasis control in Northeast Thailand: Proposal for a new approach. Appl. Parasitol. 35, 118, 1994. 26. Sithithaworn, P., and Haswell-Elkins, M. Epidemiology of Opisthorchis viverrini. Acta Trop. 88, 187, 2003. 27. Srisawangwong, T. et al. Metacercariae isolated from cyprinoid �shes in Khon Kaen District by digestion technic. Southeast Asian J. Trop. Med. Public Health 28(Suppl 1), 224, 1997. 28. Waikagul, J. Opisthorchis viverrini metacercaria in Thai freshwater �sh. Southeast Asian J. Trop. Med. Public Health 29, 324, 1998. 29. Hinz, E. Intestinal helminths in Bangkok stray dogs and their role in public health (author’s transl). Zentralbl Bakteriol. Mikrobiol. Hyg. [B] 171, 79, 1980. 30. Scholz, T. et al. Helminth parasites of cats from the Vientiane province, Laos, as indicators of the occurrence of causative agents of human parasitoses. Parasite 10, 343, 2003. 31. Enes, J.E., Wages, A.J., Malone, J.B., and Teasana, S. Prevalence of Opisthorchis viverrini infection in the canine and feline in three villages, Khon kaen Province, Northeastern Thailand. Southeast Asian J. Trop. Med. Public Health 41, 36, 2010. 32. WHO. Report of Joint WHO/FAO Workshop on Food-Borne Trematode Infections in Asia, Hanoi, p. 1, 2004. 33. Rim, H.J. Clonorchiasis: An update. J. Helminthol. 79, 269, 2005. 34. Keiser, J., and Utzinger, J. Emerging food-borne trematodiasis. Emerg. Infect. Dis. 11, 1507, 2005. 35. Yossepowitch, O. et al. Opisthorchiasis from imported raw �sh. Emerg. Infect. Dis. 10, 2122, 2004.
36. Stauffer, W.M. et al. Biliary liver �ukes (Opisthorchiasis and Clonorchiasis) in immigrants in the United States: Often subtle and diagnosed years after arrival. J. Travel. Med. 11, 157, 2004.
37. Armignacco, O. et al. Human illnesses caused by Opisthorchis felineus �ukes, Italy. Emerg. Infect. Dis. 14, 1902, 2008.
38. Jongsuksuntigul, P., and Imsomboon, T. Opisthorchiasis control in Thailand. Acta Trop. 88, 229, 2003.
39. Rim, H.J. et al. Prevalence of intestinal parasite infections on a national scale among primary schoolchildren in Laos. Parasitol. Res. 91, 267, 2003.
40. WHO. Control of food-borne trematode infections. Report of a WHO Study Group. World Health Organ. Tech. Rep. Ser. 849, 1, 1995.
41. Zhu, Y.C. et al. Application of dipstick dye immunoassay (DDIA) kit for the diagnosis of schistosomiasis mekongi. Acta Trop. 96, 137, 2005.
42. Sadun CMEH. Studies on the epidemiology of the human intestinal �uke, Fasciolopsis buski (Lankester) in central Thailand. Am. J. Trop. Med. Hyg. 2, 1070, 1953.
43. Sithithaworn, P. et al. Epidemiology of food-borne trematodes and other parasite infections in a �shing community on the Nam Ngum reservoir, Lao PDR. Southeast Asian J. Trop. Med. Public Health 37, 1083, 2006.
44. Upatham, E.S. et al. Rate of re-infection by Opisthorchis viverrini in an endemic Northeast Thai community after chemotherapy. Int. J. Parasitol. 18, 643, 1988.
45. Sornmani, S. et al. A pilot project for controlling O. viverrini infection in Nong Wai, Northeast Thailand, by applying praziquantel and other measures. Arzneimittelforschung 34, 1231, 1984.
46. Saowakontha, S. et al. Field trials in the control of Opisthorchis viverrini with an integrated programme in endemic areas of Northeast Thailand. Parasitology 106(Pt 3), 283, 1993.
47. Lan-Anh, N.T. et al. Animal reservoir hosts and �sh-borne zoonotic trematode infections on �sh farms, Vietnam. Emerg. Infect. Dis. 15, 540, 2009.
48. Upatham, E.S. et al. Morbidity in relation to intensity of infection in Opisthorchiasis viverrini: Study of a community in Khon Kaen, Thailand. Am. J. Trop. Med. Hyg. 31, 1156, 1982.
49. Pungpak, S. et al. Ultrasonographic study of the biliary system in opisthorchiasis patients after treatment with praziquantel. Southeast Asian J. Trop. Med. Public Health 20, 157, 1989.
50. Upatham, E.S. et al. Relationship between prevalence and intensity of Opisthorchis viverrini infection, and clinical symptoms and signs in a rural community in North-east Thailand. Bull. World Health Organ 62, 451, 1984.
51. Mairiang, E. et al. Ultrasound screening for Opisthorchis viverrini-associated cholangiocarcinomas: Experience in an endemic area. Asian Pac. J. Cancer Prev. 7, 431, 2006.
52. Mairiang, E., and Mairiang, P. Clinical manifestation of opisthorchiasis and treatment. Acta Trop. 88, 221, 2003.
53. Mairiang, E. et al. Relationship between intensity of Opisthorchis viverrini infection and hepatobiliary disease detected by ultrasonography. J. Gastroenterol. Hepatol. 7, 17, 1992.
54. Choi, D. et al. Sonographic �ndings of active Clonorchis sinensis infection. J. Clin. Ultrasound 32,17, 2004.
55. Dhiensiri, T. et al. Roentgenographically controlled healing of gall bladder lesions in opisthorchiasis after praziquantel treatment. Arzneimittelforschung 34, 1175, 1984. 56. Haswell-Elkins, M.R. et al. Cross-sectional study of Opisthorchis viverrini infection and cholangiocarcinoma in communities within a high-risk area in Northeast Thailand. Int. J. Cancer 59, 505, 1994. 57. Elkins, D.B. et al. Cross-sectional patterns of hepatobiliary abnormalities and possible precursor conditions of cholangiocarcinoma associated with Opisthorchis viverrini infection in humans. Am. J. Trop. Med. Hyg. 55, 295, 1996. 58. Mairiang, E. et al. Reversal of biliary tract abnormalities associated with Opisthorchis viverrini infection following praziquantel treatment. Trans. R Soc. Trop. Med. Hyg. 87, 194, 1993. 59. Bhamarapravati, N. et al. Liver changes in hamsters infected with a liver �uke of man, Opisthorchis viverrini. Am. J. Trop. Med. Hyg. 27, 787, 1978. 60. Hou, P.C. The pathology of Clonorchis sinensis infestation of the liver. J. Pathol. Bacteriol. 70, 53, 1955. 61. Koompirochana, C. et al. Opisthorchiasis: A clinicopathologic study of 154 autopsy cases. Sotheast Asian J. Trop. Med. Public Health 9, 60, 1978. 62. Riganti, M. et al. Human pathology of Opisthorchis viverrini infection: A comparison of adults and children. Southeast Asian J. Trop. Med. Public Health 20, 95, 1989. 63. Tansurat, P. Opisthorchiasis. In: Marcial-Rojas, R.A. (ed.), Pathology of Protozoal and Helminthic Diseases. Williams & Wilkins, Baltimore, 536pp., 1971. 64. Sripa, B. et al. Liver �uke induces cholangiocarcinoma. PLoS Med. 4, e201, 2007. 65. Riganti, M. et al. Opisthorchis viverrini eggs and adult �ukes as nidus and composition of gallstones. Southeast Asian J. Trop. Med. Public Health 19, 633, 1988. 66. Sripa, B. et al. Opisthorchiasis-associated biliary stones: Light and scanning electron microscopic study. World J. Gastroenterol. 10, 3318, 2004. 67. Elkins, D.B. et al. A high frequency of hepatobiliary disease and suspected cholangiocarcinoma associated with heavy Opisthorchis viverrini infection in a small community in North-east Thailand. Trans. R. Soc. Trop. Med. Hyg. 84, 715, 1990. 68. Wongratanacheewin, S. et al. Immunology and molecular biology of Opisthorchis viverrini infection. Acta Trop. 88, 195, 2003. 69. Akai, P.S. et al. Serum antibody responses in opisthorchiasis. Int. J. Parasitol. 25, 971, 1995. 70. Akai, P.S. et al. Serum antibody response to Opisthorchis viverrini antigen as a marker for opisthorchiasis-associated cholangiocarcinoma. Trans. R Soc. Trop. Med. Hyg. 88, 471, 1994. 71. Itoh, M. et al. Association of antibodies to Opisthorchis viverrini with hepatobiliary disease in Northeastern Thailand. Am. J. Trop. Med. Hyg. 51, 424, 1994. 72. Wongratanacheewin, S., and Sirisinha, S. Analysis of Opisthorchis viverrini antigens: Physicochemical characterization and antigen localization. Southeast Asian J. Trop. Med. Public Health 18, 511, 1987. 73. Ruangkunaporn, Y. et al. Changes in serum antibodies to Opisthorchis viverrini in humans and hamsters following treatment of opisthorchiasis. Asian Pac. J. Allergy Immunol. 12, 83, 1994. 74. Suksumek, N. et al. TaqMan real-time PCR assay for speci�c detection of Opisthorchis viverrini DNA in Thai patients with hepatocellular carcinoma and cholangiocarcinoma. Exp. Parasitol. 119, 217, 2008. 75. Sripa, B., and Kaewkes, S. Localisation of parasite antigens and in�ammatory responses in experimental opisthorchiasis. Int. J. Parasitol. 30, 735, 2000.
76. Pinlaor, S. et al. Hepatobiliary changes, antibody response, and alteration of liver enzymes in hamsters re-infected with Opisthorchis viverrini. Exp. Parasitol. 108, 32, 2004.
77. Flavell, D.J., and Flavell, S.U. Opisthorchis viverrini: Pathogenesis of infection in immunodeprived hamsters. Parasite Immunol. 8, 455, 1986.
78. Wongratanacheewin, S. et al. Immunodepression in hamsters experimentally infected with Opisthorchis viverrini. J. Helminthol. 61, 151, 1987.
79. Thamavit, W. et al. Effects of dimethylnitrosamine on induction of cholangiocarcinoma in Opisthorchis viverrini-infected Syrian golden hamsters. Cancer Res. 38, 4634, 1978.
80. Haswell-Elkins, M.R. et al. Distribution patterns of Opisthorchis viverrini within a human community. Parasitology 103(Part 1), 97, 1991.
81. Flavell, D.J. et al. Opisthorchis viverrini: Partial success in adoptively transferring immunity with spleen cells and serum in the hamster. J. Helminthol. 54, 191, 1980.
82. Sirisinha, S. et al. Attempts to induce protective immunity in hamsters against infection by a liver �uke of man (Opisthorchis viverrini). Parasitology 86(Pt 1), 127, 1983.
83. Wongratanacheewin, S. et al. Effect of praziquantel treatment on antibody levels and lymphoproliferative responses in patients with opisthorchiasis. Southeast Asian J. Trop. Med. Public Health 19, 109, 1988.
84. Sirisinha, S. et al. Humoral immune responses in hamsters infected with Opisthorchis viverrini. Southeast Asian J. Trop. Med. Public Health 14, 243, 1983.
85. Sirisinha, S., and Wongratanacheewin, S. Immunization of hamsters against Opisthorchis viverrini infection. Southeast Asian J. Trop. Med. Public Health 17, 567, 1986.
86. Jittimanee, J. et al. Cytokine expression in hamsters experimentally infected with Opisthorchis viverrini. Parasite Immunol. 29, 159, 2007.
87. Sripa, B. et al. Advanced periductal �brosis from infection with the carcinogenic human liver �uke Opisthorchis viverrini correlates with elevated levels of interleukin-6. Hepatology 50, 1273, 2009.
88. IARC. IARC Monographs on the Evaluation of Carcinogenic Risks to Humans. International Agency for Research on Cancer, World Health Organization, Lyon, France, vol. 121, p. 61, 1994.
89. Parkin, D.M. et al. Cholangiocarcinoma: Epidemiology, mechanisms of carcinogenesis and prevention. Cancer Epidemiol. Biomarkers Prev. 2, 537, 1993.
90. Srivatanakul, P. et al. Epidemiology of liver cancer: An overview. Asian Pac. J. Cancer. Prev. 5, 118, 2004.
91. Lim, M.K. et al. Clonorchis sinensis infection and increasing risk of cholangiocarcinoma in the Republic of Korea. Am. J. Trop. Med. Hyg. 75, 93, 2006.
92. Honjo, S. et al. Genetic and environmental determinants of risk for cholangiocarcinoma via Opisthorchis viverrini in a densely infested area in Nakhon Phanom, Northeast Thailand. Int. J. Cancer 117, 854, 2005. 93. Uttaravichien, T. et al. Intrahepatic cholangiocarcinoma in Thailand. J. Hepatobiliary Pancreat. Surg. 6, 128, 1999.
94. Uttaravichien, T., and Buddhisawasdi, V. Experience of nonjaundiced cholangiocarcinoma. Hepatogastroenterology 37, 608, 1990.
95. Pairojkul, C. et al. Multistage carcinogenesis of liver�uke-associated cholangiocarcinoma in Thailand. Princess Takamatsu Symp. 22, 77, 1991.
96. Hou, P.C. The relationship between primary carinoma of the liver and infestation with Clonorchis sinesis. J. Pathol. Bact. 72, 239, 1956. 97. Haswell-Elkins, M.R. et al. Immune responsiveness and parasite-speci�c antibody levels in human hepatobiliary disease associated with Opisthorchis viverrini infection. Clin. Exp. Immunol. 84, 213, 1991. 98. Ohshima, H., and Bartsch, H. Chronic infections and in�ammatory processes as cancer risk factors: Possible role of nitric oxide in carcinogenesis. Mutat. Res. 305, 253, 1994. 99. Kawanishi, S. et al. Oxidative and nitrative DNA damage in animals and patients with in�ammatory diseases in relation to in�ammation-related carcinogenesis. Biol. Chem. 387, 365, 2006. 100. Ma, N. et al. Accumulation of 8-nitroguanine in human gastric epithelium induced by Helicobacter pylori infection. Biochem. Biophys. Res. Commun. 319, 506, 2004. 101. Pinlaor, S. et al. Nitrative and oxidative DNA damage in intrahepatic cholangiocarcinoma patients in relation to tumor invasion. World J. Gastroenterol. 11, 4644, 2005. 102. Srivatanakul, P. et al. Opisthorchis viverrini infestation and endogenous nitrosamines as risk factors for cholangiocarcinoma in Thailand. Int. J. Cancer 48,821, 1991. 103. Haswell-Elkins, M.R. et al. Liver �uke infection and cholangiocarcinoma: Model of endogenous nitric oxide and extragastric nitrosation in human carcinogenesis. Mutat. Res. 305, 241, 1994. 104. Satarug, S. et al. Thiocyanate-independent nitrosation in humans with carcinogenic parasite infection. Carcinogenesis 17,1075, 1996. 105. Satarug, S. et al. Relationships between the synthesis of N-nitrosodimethylamine and immune responses to chronic infection with the carcinogenic parasite, Opisthorchis viverrini, in men. Carcinogenesis 19, 485, 1998. 106. Satarug, S. et al. Induction of cytochrome P450 2A6 expression in humans by the carcinogenic parasite infection, Opisthorchiasis viverrini. Cancer Epidemiol. Biomarkers. Prev. 5, 795, 1996. 107. Ohshima, H. et al. Increased nitrosamine and nitrate biosynthesis mediated by nitric oxide synthase induced in hamsters infected with liver �uke (Opisthorchis viverrini). Carcinogenesis 15, 271, 1994. 108. Thanan, R. et al. Urinary 8-oxo-7,8-dihydro-2′deoxyguanosine in patients with parasite infection and effect of antiparasitic drug in relation to cholangiocarcinogenesis. Cancer Epidemiol. Biomarkers Prev. 17, 518, 2008. 109. Dechakhamphu, S. et al. Lipid peroxidation and etheno DNA adducts in white blood cells of liver �uke-infected patients: Protection by plasma alpha-tocopherol and praziquantel. Cancer Epidemiol. Biomarkers Prev. 19, 310, 2009. 110. Dechakhamphu, S. et al. High excretion of etheno adducts in liver �uke-infected patients: Protection by praziquantel against DNA damage. Cancer Epidemiol. Biomarker. Prev. 17, 1658, 2008. 111. Pinlaor, S. et al. Opisthorchis viverrini antigen induces the expression of Toll-like receptor 2 in macrophage RAW cell line. Int. J. Parasitol. 35, 591, 2005. 112. Pinlaor, S. et al. Repeated infection with Opisthorchis viverrini induces accumulation of 8-nitroguanine and 8-oxo-7, 8-dihydro-2′-deoxyguanine in the bile duct of hamsters via inducible nitric oxide synthase. Carcinogenesis 25, 1535, 2004. 113. Thuwajit, C. et al. Increased cell proliferation of mouse �broblast NIH-3T3 in vitro induced by excretory/secretory product(s) from Opisthorchis viverrini. Parasitology 129, 455, 2004.
114. Thuwajit, C. et al. Gene expression pro�ling de�ned pathways correlated with �broblast cell proliferation induced by Opisthorchis viverrini excretory/secretory product. World J. Gastroenterol. 12, 3585, 2006.
115. Pinlaor, S. et al. iNOS-dependent DNA damage via NF-kappaB expression in hamsters infected with Opisthorchis viverrini and its suppression by the antihelminthic drug praziquantel. Int. J. Cancer 119, 1067, 2006.
116. Boonmars, T. et al. Apoptosis-related gene expression in hamster opisthorchiasis post praziquantel treatment. Parasitol. Res. 102, 447, 2008.
117. Maeda, H. Carcinogenesis via microbial infection. Gan To Kagaku Ryoho 25, 1474, 1998.
118. Holzinger, F. et al. Mechanisms of biliary carcinogenesis: A pathogenetic multi-stage cascade towards cholangiocarcinoma. Ann. Oncol. 10(Suppl 4), 122, 1999.
119. Sithithaworn, P., Yongvanit, P., Tesana, S., and Pairojkul, C. Liver �ukes. In: Murrell, D., and Fried, B. (eds), Parasitic Foodborne Zoonoses. Springer-Verlag, Berlin, Germany, p. 1, 2008.
120. Hong, S.T. et al. The Kato–Katz method is reliable for diagnosis of Clonorchis sinensis infection. Diagn. Microbiol. Infect. Dis. 47, 345, 2003.
121. Viyanant, V. et al. A comparison of a modi�ed quick-Kato technique and the Stoll dilution method for �eld examination for Opisthorchis viverrini eggs. J. Helminthol. 57, 191, 1983.
122. Radomyos, P., Bunnag, D., and Harinasuta, T. Worms recovered in stools following praziquantel treatment. Drug Res. 34, 1215, 1984.
123. Radomyos, P. et al. Multi-infection with helminths in adults from Northeast Thailand as determined by post-treatment fecal examination of adult worms. Trop. Med. Parasitol. 45, 133, 1994.
124. Chai, J.Y. et al. High prevalence of liver and intestinal �uke infections among residents of Savannakhet Province in Laos. Korean J. Parasitol. 45, 213, 2007.
125. Poopyruchpong, N. et al. Diagnosis of opisthorchiasis by enzyme-linked immunosorbent assay using partially puri�ed antigens. Asian Pac. J. Allergy. Immunol. 8, 27, 1990.
126. Wongsaroj, T. et al. Af�nity puri�ed oval antigen for diagnosis of Opisthorchiasis viverrini. Asian Pac. J. Allergy Immunol. 19, 245, 2001.
127. Sirisinha, S. et al. Detection of Opisthorchis viverrini by monoclonal antibody-based ELISA and DNA hybridization. Am. J. Trop. Med. Hyg. 44, 140, 1991.
128. Ruangsittichai, J. et al. Opisthorchis viverrini: Identi�cation of a glycine-tyrosine rich eggshell protein and its potential as a diagnostic tool for human opisthorchiasis. Int. J. Parasitol. 36, 1329, 2006.
129. Thammapalerd, N. et al. Detection of antibodies against Opisthorchis viverrini in patients before and after treatment with praziquantel. Southeast Asian J. Trop. Med. Public Health 19, 101, 1988.
130. Laha, T. et al. The bandit, a new DNA transposon from a hookworm—Possible horizontal genetic transfer between host and parasite. PLoS Negl. Trop. Dis. 1, e35, 2007. 131. Laha, T. et al. Gene discovery for the carcinogenic human liver �uke, Opisthorchis viverrini. BMC Genomics 8, 189, 2007. 132. Laha, T. et al. Asparaginyl endopeptidase from the carcinogenic liver �uke, Opisthorchis viverrini, and its potential for serodiagnosis. Int. J. Infect. Dis. 12, e49, 2008. 133. Suttiprapa, S. et al. Ov-APR-1, an aspartic protease from the carcinogenic liver �uke, Opisthorchis viverrini: Functional expression, immunolocalization and subsite speci�city. Int. J. Biochem. Cell Biol., 41, 1148, 2009. 134. Chaicumpa, W. et al. Speci�c monoclonal antibodies to Opisthorchis viverrini. Int. J. Parasitol. 21, 969, 1991. 135. Sirisinha, S. et al. Evaluation of a monoclonal antibody-based enzyme linked immunosorbent assay for the diagnosis of Opisthorchis viverrini infection in an endemic area. Am. J. Trop. Med. Hyg. 52, 521, 1995. 136. Wongratanacheewin, S. et al. Development of a PCR-based method for the detection of Opisthorchis viverrini in experimentally infected hamsters. Parasitology 122, 175, 2001. 137. Wongratanacheewin, S. et al. Detection of Opisthorchis viverrini in human stool specimens by PCR. J. Clin. Microbiol. 40, 3879, 2002. 138. Stensvold, C.R. et al. Evaluation of PCR based coprodiagnosis of human opisthorchiasis. Acta Trop. 97, 26, 2006. 139. Duenngai, K. et al. Improvement of PCR for detection of Opisthorchis viverrini DNA in human stool samples. J. Clin. Microbiol. 46, 366, 2008. 140. Parvathi, A. et al. Development and evaluation of a polymerase chain reaction (PCR) assay for the detection of Opisthorchis viverrini in �sh. Acta Trop. 107, 13, 2008. 141. Ando, K. et al. Nucleotide sequence of mitochondrial CO I and ribosomal ITS II genes of Opisthorchis viverrini in Northeast Thailand. Southeast Asian J. Trop. Med. Public Health 32(Suppl 2), 17, 2001. 142. Le, T.H. et al. Clonorchis sinensis and Opisthorchis viverrini: Development of a mitochondrial-based multiplex PCR for their identi�cation and discrimination. Exp. Parasitol. 112, 109, 2006. 143. Pauly, A. et al. Molecular characterization and differentiation of opisthorchiid trematodes of the species Opisthorchis felineus (Rivolta, 1884) and Metorchis bilis (Braun, 1790) using polymerase chain reaction. Parasitol. Res. 90, 409, 2003. 144. Touch, S. et al. Discovery of Opisthorchis viverrini metacercariae in freshwater �sh in southern Cambodia. Acta Trop. 111, 108, 2009. 145. Traub, R.J. et al. A new PCR-based approach indicates the range of Clonorchis sinensis now extends to Central Thailand. PLoS Negl. Trop. Dis. 3, e367, 2009. 146. Parvathi, A. et al. Clonorchis sinensis: Development and evaluation of a nested polymerase chain reaction (PCR) assay. Exp. Parasitol. 115,291, 2007. 147. Sato, M. et al. Discrimination of O. viverrini, C. sinensis, H. pumilio and H. taichui using nuclear DNA-based PCR targeting ribosomal DNA ITS regions. Acta Trop. 109, 81, 2009. 148. Thaenkham, U. et al. Discrimination of Opisthorchis viverrini from Haplorchis taichui using COI sequence marker. Acta Trop. 103, 26, 2007. 39 Chapter 39 - Paragonimus
5. Keiser, J., and Utzinger, J. Emerging foodborne trematodiasis. Emerg. Infect. Dis. 11, 1507, 2005.
6. Narain, K., Agatsuma, T., and Blair, D. Paragonimus. In: Liu, D.Y. (ed.), Molecular Detection of Foodborne Pathogens. Chapter 61. CRC Press, Inc., Boca Raton, p. 827, 2009.
7. Miyazaki, I. Lung �ukes in the world–Morphology and life history. In: Sasa, M. (ed.), A Symposium on Epidemiology of Parasitic Diseases, International Medical Foundation of Japan, Tokyo, p. 101, 1974.
8. Blair, D., Agatsuma, T., and Wang, W. Paragonimiasis. In: Murrell, K.D., and Fried, B. (eds.), World Class Parasites: Food-borne Parasitic Zoonoses: Fish and Plant-Borne Parasites, Vol. 11. Springer, New York, p. 117, 2007.
9. Blair, D., Xu, Z.B., and Agatsuma, T. Paragonimiasis and the genus Paragonimus. Adv. Parasitol. 42, 113, 1999.
10. Blair, D. et al. A molecular perspective on the genera Paragonimus Braun, Euparagonimus Chen and Pagumogonimus Chen. J. Helminthol. 73, 295, 1999.
11. Blair, D. et al. Paragonimus skrjabini Chen, 1959 (Digenea: Paragonimidae) and related species in eastern Asia: A combined molecular and morphological approach to identi�cation and taxonomy. Syst. Parasitol. 60, 1, 2005.
12. Blair, D. et al. Geographical genetic structure within the human lung �uke, Paragonimus westermani, detected from DNA sequences. Parasitology 115, 411, 1997.
13. Terasaki, K. et al. Tetraploids of the lung �uke Paragonimus westermani found in China. Parasitol. Res. 81, 627, 1995.
14. Agatsuma, T., and Habe, S. Electrophoretic studies on enzymes of diploid and triploid Paragonimus westermani. Parasitology 91, 489, 1985.
15. Agatsuma, T. et al. Genetic differentiation between Malaysian and other Asian Paragonimus westermani. Trop. Biomed. 10, 45, 1993.
16. Iwagami, M. et al. Molecular phylogeographic studies on Paragonimus westermani in Asia. J. Helminthol. 74, 315, 2000.
17. Park, G.M., Im, K.I., and Yong, T.S. Phylogenetic relationship of ribosomal ITS2 and mitochondrial COI among diploid and triploid Paragonimus westermani isolates. Korean J. Parasitol. 41, 47, 2003.
18. Doanh, P.N. et al. Discovery of Paragonimus westermani in Vietnam and its molecular phylogenetic status in P. westermani complex. Parasitol. Res. 104, 1149, 2009.
19. Blair, D. et al. Phylogenetic relationships among the Thai species of Paragonimus inferred from DNA sequences. In: Tada, I., Kojima, S., and Tsuji, M. (eds.), IX International Congress of Parasitology (ICOPA IX), Monduzzi Editore, Bologna, p. 643, 1998.
20. Sugiyama, H. et al. New form of Paragonimus westermani discovered in Thailand: Morphological characteristics and host susceptibility. Southeast Asian J. Trop. Med. Public Health 38(Suppl. 1), 87, 2007.
21. Binchai, S. et al. Molecular systematics of a new form of Paragonimus westermani discovered in Thailand, Southeast Asian J. Trop. Med. Public Health 38(Suppl. 1), 92, 2007.
22. Singh, S.T., Singh, D.L., and Sugiyama, H. Possible discovery of Chinese lung �uke, Paragonimus skrjabini in Manipur, India. Southeast Asian J. Trop. Med. Public Health 37(Suppl. 3), 53, 2006.
23. Kamo, H. et al. On the occurrence of a new lung �uke, Paragonimus miyazakii n. sp. in Japan (Trematoda: Troglotrematidae). Yonago Acta Med. 5, 43, 1961.
24. Waikagul, J. A new species of Paragonimus (Trematoda: Troglotrematidae) from a cat infected with metacercariae from mountain crabs Larnaudia larnaudii. J. Parasitol. 93, 1496, 2007. 25. Thaenkham, U., and Waikagul, J. Molecular phylogenetic relationship of Paragonimus pseudoheterotremus. Southeast Asian J. Trop. Med. Public Health 39, 217, 2008. 26. Aka, N.A. et al. Human paragonimiasis in Africa. Ann. Afr. Med. 7, 153, 2008. 27. Zhong, H.L. et al. Recent progress in studies of Paragonimus and paragonimiasis control in China. Chin. Med. J. 94, 483, 1981. 28. Uchiyama, F., Morimoto, Y., and Nawa, Y. Re-emergence of paragonimiasis in Kyushu, Japan. Southeast Asian J. Trop. Med. Public Health 30, 686, 1999. 29. Singh, T.S. et al. Paragonimiasis in Manipur. Indian J. Med. Res. 97, 247, 1993. 30. Singh, T.S., Mutum, S.S., and Razaque, M.A. Pulmonary paragonimiasis: Clinical features, diagnosis and treatment of 39 cases in Manipur. Trans. R. Soc. Trop. Med. Hyg. 80, 967, 1986. 31. Singh, T.S., and Sugiyama, H. Paragonimiasis in India: A newly emerging food borne parasitic disease. Clin. Parasitol. 19, 95, 2008. 32. Razaque, M.A., Mutum, S.S., and Singh, T.S. Recurrent haemoptysis? Think of paragonimiasis. Trop. Doct. 21, 153, 1991. 33. Ogakwu, M., and Nwokolo, C. Radiological �ndings in pulmonary paragonimiasis as seen in Nigeria: A review based on one hundred cases. Br. J. Radiol. 46, 699, 1973. 34. Romeo, D.P., and Pollock, J.J. Pulmonary paragonimiasis: Diagnostic value of pleural �uid analysis. South. Med. J. 79, 241, 1986. 35. Watanabe, S. et al. Pulmonary paragonimiasis mimicking lung cancer on FDG-PET imaging. Anticancer Res. 23, 3437, 2003. 36. Yoshino, I. et al. Paragonimiasis westermani presenting as an asymptomatic nodular lesion in the lung: Report of a case. Surg. Today 28, 108, 1998. 37. Johnson, R.J., and Johnson, J.R. Paragonimiasis in Indochinese refugees. Roentgenographic �ndings with clinical correlations. Am. Rev. Respir. Dis. 128, 534, 1983. 38. Jeon, K. et al. Clinical features of recently diagnosed pulmonary paragonimiasis in Korea. Chest 128, 1423, 2005. 39. Chung, H.L., and Tsao, W.C. Paragonimus westermani (Szechuan variety) and a new species of lung �uke–paragonimiasis szechuanensis, part II. Studies on clinical aspects of Paragonimiasis szechuanensis: A new clinical entity. Chin. Med. J. 81, 419, 1962. 40. Singh, P.I., Singh, I.P., and Singh, L.B.M. Paragonimiasis: Review of 45 cases. Indian J. Med. Microbiol. 10, 243, 1992. 41. Jeong, M.G. et al. Retroperitoneal paragonimiasis: A case of ectopic paragonimiasis presenting as periureteral masses. J. Comput. Assist. Tomogr. 23, 696, 1999. 42. Singh, T.S., Khamo, V., and Sugiyama, H. Cerebral paragonimiasis mimicking tuberculoma: First case report in India. Trop. Parasitol. 1, 39, 2011. 43. Miyazaki, I., and Nishimura, K. Cerebral paragonimiasis. In: Hornabrook, R.W. (ed.), Topics on Tropical Neurology, Contemporary Neurology Series Vol. 12, Chapter 7. F.A. Davis Company, Philadelphia, p. 109, 1975. 44. Hughes, A.J., and Biggs, B.A. Parasitic worms of the central nervous system: An Australian perspective. Intern. Med. J. 32, 541, 2002. 45. Jaroonvesama, N. Differential diagnosis of eosinophilic meningitis. Parasitol. Today 4, 262, 1988.
46. Chang, K.H. et al. An imaging diagnosis of cerebral paragonimiasis: CT and MR �ndings and correlation with ELISA antibody test. J. Korean Radiol. Soc. 29, 345, 1993.
47. Choo, J.D. et al. Chronic cerebral paragonimiasis combined with aneurismal subarachnoid haemorrhage. Am. J. Trop. Med. Hyg. 69, 466, 2003.
48. Singh, T.S. et al. Paragonimus heterotremus infection in Nagaland: A new focus of paragonimiasis in India. Indian J. Med. Microbiol. 27, 123, 2009.
49. Komiya, Y., and Yokogawa, M. The recovering of Paragonimus eggs from stools of paragonimiasis patients by AMS III centrifuging technic. Jpn. J. Med. Sci. Biol. 6, 207, 1953.
50. Young, K.H. et al. Ethyl acetate as a substitute for diethyl ether in the formalin-ether sedimentation technique. J. Clin. Microbiol. 10, 852, 1979.
51. Kuroki, M. et al. High-resolution computed tomography �ndings of P. westermani. J. Thorac. Imaging 20, 210, 2005.
52. Song, H.O. et al. Skin test for paragonimiasis among schoolchildren and villagers in Namback District, Luangprabang province, Lao PDR. Korean J. Parasitol. 46, 179, 2008.
53. Sawada, T., Takei, K., and Yoneyama, K. Studies on the immunodiagnosis of paragonimiasis. II. Intradermal tests with fractionated antigens. J. Infect. Dis. 114, 315, 1964.
54. Yokogawa, M., and Tsuji, M. Immunological diagnosis as the screening method of paragonimiasis in the endemic area of paragonimiasis. In: First Regional Symposium on Scienti�c Knowledge of Tropical Parasites, University of Singapore, 5 to 9 November, 1962, UNESCO, Singapore, p. 194, 1962.
55. Biguet, J. et al. Contribution of immunoelectrophoretic analysis to the knowledge of worm antigens: Practical effects on their standardization, their puri�cation and diagnosis of helminthiasis by immunoelectrophoresis. Rev. Immunol. Ther. Antimicrob. 29, 5, 1965.
56. Katamine, D., Imai, J., and Iwamoto, I. Immunological study on paragonimiasis: Agar-gel precipitin reactions in paragonimiasis. Trop. Med. 10, 28, 1968.
57. Tsuji, M., Capron, A., and Yokogawa, M. Immunodiffusion test in the diagnosis of paragonimiasis. In: Second International Congress of Parasitology (ICOPA II), Colloquium on Immunodiagnosis, Washington, D.C., p. 6, 1970.
58. Tsuji, M. On the immunoelectrophoresis for helminthological researches. Jpn. J. Parasitol. 23, 335, 1974.
59. Pariyanonda, S. et al. Serodiagnosis of human paragonimiasis caused by Paragonimus heterotremus. Southeast Asian J. Trop. Med. Public Health 21, 103, 1990.
60. Quicho, L.P. et al. Humoral immune response of cats to Paragonimus infection. Southeast Asian J. Trop. Med. Public Health 12, 364, 1981.
61. Waikagul, J. Serodiagnosis of paragonimiasis by enzymelinked immunosorbent assay and immunoelectrophoresis. Southeast Asian J. Trop. Med. Public Health 20, 243, 1989.
62. Imai, J. Evaluation of ELISA for the diagnosis of paragonimiasis westermani Trans. R. Soc. Trop. Med. Hyg. 81, 3, 1987.
63. Zhang, Z. et al. Diagnosis of active Paragonimus westermani infections with monoclonal antibody-based antigen detection assay. Am. J. Trop. Med. Hyg. 49, 329, 1993.
64. Itoh, M., and Sato, S. Multi-dot enzyme-linked immunosorbent assay for serodiagnosis of trematodiasis. Southeast Asian J. Trop. Med. Public Health 21, 471, 1990.
65. Indrawati, I. et al. Studies on immunodiagnosis of human paragonimiasis and speci�c antigen of Paragonimus heterotremus. Int. J. Parasitol. 21, 395, 1991. 66. Narain, K., Devi, K.R., and Mahanta, J. Development of enzyme-linked immunosorbent assay for serodiagnosis of human paragonimiasis. Indian J. Med. Res. 121, 739, 2005. 67. Nakamura-Uchiyama, F., Onah, D.N., and Nawa, Y. Clinical features of paragonimiasis cases recently found in Japan: Parasite-speci�c immunoglobulin M and G antibody classes. Clin. Infect. Dis. 32, e171, 2001. 68. Ikeda, T. et al. Parasite-speci�c IgE and IgG levels in the serum and pleural effusion of Paragonimiasis westermani patients. Am. J. Trop. Med. Hyg. 47, 104, 1992. 69. Sugiyama, H. et al. Characterization and localization of Paragonimus westermani antigen stimulating antibody formation in both the infected cat and rat. J. Parasitol. 73, 363, 1987. 70. Maleewong, W. et al. Antigenic components of Paragonimus heterotremus recognized by infected human serum. Parasite Immunol. 13, 89, 1991. 71. Maleewong, W. et al. Analysis of antibody levels before and after praziquantel treatment in human paragonimiasis heterotremus. Asian Pac. J. Allergy Immunol. 10, 69, 1992. 72. Dekumyoy, P., Waikagul, J., and Eom, K.S. Human lung �uke Paragonimus heterotremus: Differential diagnosis between Paragonimus heterotremus and Paragonimus westermani infections by EITB. Trop. Med. Int. Health 3, 52, 1998. 73. Kong, Y. et al. Immunoglobulin G (IgG) subclass and IgE responses in human paragonimiasis caused by three different species. Clin. Diagn. Lab. Immunol. 5, 474, 1998. 74. Gan, X.X. et al. Development of rapid diagnostic kit (Dot Immunogold Filtration Assay) for detection of antibodies against Paragonimus westermani. Chin. J. Zoo. 21, 988, 2005. 75. Qian, B.Z., and Sugiyama, H. Development of an immunological diagnostic kit for paragonimiasis and its use for an epidemiological survey of zoonotic paragonimiasis in Mainland China, 1. Multi-antigen dot immunogold �ltration assay (Multi-DIGFA) for serodiagnosis of paragonimiasis. Final report of the JHSF project, The Japan Health Sciences Foundation, Tokyo, 2007. 76. Singh, T.S. et al. Morphological and molecular characterizations of Paragonimus heterotremus, the causative agent of human paragonimiasis in India. Southeast Asian J. Trop. Med. Public Health 38(Suppl. 1), 82, 2007. 77. Chang, Z.S. et al. Gene sequencing for identi�cation of Paragonimus eggs from a human case. Chin. J. Parasitol. Parasitic Dis. 18, 213, 2000. 78. Le, T.H. et al. Paragonimus heterotremus Chen and Hsia, (1964), in Vietnam: A molecular identi�cation and relationships of isolates from different hosts and geographical origins. Acta Trop. 98, 25, 2006. 79. Intapan, P.M. et al. Detection of Paragonimus heterotremus eggs in experimentally infected cats by a polymerase chain reaction-based method. J. Parasitol. 91, 195, 2005. 80. Nkouawa, A. et al. Paragonimiasis in Cameroon: Molecular identi�cation, serodiagnosis and clinical manifestations. Trans. R. Soc. Trop. Med. Hyg. 103, 255, 2009. 81. Agatsuma, T. et al. Mitochondrial DNA differentiation of Japanese diploid and triploid Paragonimus westermani. J. Helminthol. 68, 7, 1994. 82. Sugiyama, H. et al. Molecular discrimination between individual metacercariae of Paragonimus heterotremus and P. westermani occurring in Thailand. Southeast Asian J. Trop. Med. Public Health 36(Suppl. 4), 102, 2005. 83. Sugiyama, H. et al. A multiplex PCR for discrimination between Paragonimus westermani and P. miyazakii at the metacercarial stage. Southeast Asian J. Trop. Med. Public Health 35(Suppl. 1), 327, 2004.
84. Intapan, P.M. et al. Genomic characterization of lung �ukes, Paragonimus heterotremus, P. siamensis, P. harinasutai, P. westermani and P. bangkokensis by RAPD markers. Vet. Parasitol. 124, 55, 2004.
85. Maleewong, W. et al. Detection of Paragonimus heterotremus in experimentally infected cat feces by antigen capture-ELISA and by DNA hybridization. J. Parasitol. 83, 1075, 1997.
86. Sugiyama, H. et al. Polymerase chain reaction (PCR)-based molecular discrimination between Paragonimus westermani and P. miyazakii at the metacercarial stage. Mol. Cell. Probes 16, 231, 2002.
87. Rowan, W.B. The mode of hatching of the egg of Fasciola hepatica. Exp. Parasitol. 5, 118, 1956.
88. Vanijanonta, S. et al. Pulmonary paragonimiasis with expectoration of worms: A case report. Southeast Asian J. Trop. Med. Public Health 12, 104, 1981.
89. Miyazaki, I., Arellano, R.C., and Grados, O. The �rst demonstration of the �uke, Paragonimus from man in Peru. Jpn. J. Parasitol. 21, 168, 1972.
90. Miyazaki, I., and Fontan, R. Mature Paragonimus heterotremus found from a man in Laos. Jpn. J. Parasitol. 19, 109, 1970.
91. Miyazaki, I., and Harinasuta, T. The �rst case of human paragonimiasis caused by P. heterotremus Chen et Hsia, 1964. Ann. Trop. Med. Parasitol. 60, 509, 1966. 92. Sugiyama, H. et al. Studies on the geographical distribution of the lung �uke. Paragonimus westermani (Kerbert, 1878) in the southern prefectures of the Kinki District, Japan: Observations on the incidence of encysted larvae of P. westermani in Geothelphusa dehaani in Higashi-Yoshino area, Nara Prefecture. J. Vet. Med. Sci. 45, 227, 1983. 93. Maejima, J., Kamo, H., and Hatsushika, R. Studies on Paragonimus miyazakii Kamo, Nishida, Hatsushika et Tomimura, 1961. 2. Second intermediate host and intracrab stage. Yonago Acta Med. 15, 35, 1971. 94. Komalamisra, C., Asavisanu, R., and Setasuban, P. Distribution of Paragonimus heterotremus metacercariae in fresh water crab, Tiwaripotamon beusekomae Bott 1970. Southeast Asian J. Trop. Med. Public Health 19, 337, 1988. 95. Rangsiruji, A. et al. A new record of Paragonimus other than P. westermani in southern Thailand. Southeast Asian J. Trop. Med. Public Health 37(Suppl. 3), 57, 2006. 96. Bowles, J., Blair, D., and McManus, D.P. A molecular phylogeny of the human schistosomes. Mol. Phylogenet. Evol. 4, 103, 1995. 97. Rim, H.J. Paragonimiasis: Experimental and clinical experience with praziquantel in Korea. Arzneimittelforschung 34(9B), 1197, 1984. 98. Nawa, Y., and Doanh, P.N. Paragonimus bangkokensis and P. harinasutai: Reappraisal of phylogenetic status. J. Trop. Med. Parasitol. 32, 87, 2009. 40 Chapter 40 - Plagiorchis
1. Chai, J.Y., Shin, E.H., Lee, S.H., and Rim, H.J. Foodborne intestinal �ukes in Southeast Asia. Korean J. Parasitol. 47 (Suppl), S69–1S02, 2009.
2. Guk, S.M., Kim, J.L., Park, J.H., and Chai, J.Y. A human case of Plagiorchis vespertilionis (Digenea: Plagiorchiidae) infection in the Republic of Korea. J. Parasitol. 93(5), 1225– 1227, 2007.
3. Seo, B.S., Rim, H.J., and Lee, C.W. Studies on the parasitic helmiths of Korea: I. Trematodes of rodents. Korean J. Parasitol. 2(1), 20–26, 1964.
4. Hong, S.J. Surface ultrastructure of Plagiorchis muris growth and developmental stages in rats, the �nal host. Parasitol. Res. 105(4), 1077–1083, 2009.
5. MACY, R.W. The life cycle of Plagiorchis vespertilionis Parorchis. N. SSP. (Trematoda: Plagiorchidae), and observations on the effects of light on the emergence of the cercaria. J. Parasitol. 46:337–345, 1960.
6. Groschaft, J., and Tenora, F. Plagiorchis vespertilionis (Müller, 1780) (Trematoda)—A parasite of Meriones persicus (Rodentia) in Afghanistan. Folia Parasitol (Praha). 18(1), 40, 1971.
7. Hong, S.J., Woo, H.C., Lee, S.U., and Huh, S. Infection status of dragon�ies with Plagiorchis muris metacercariae in Korea. Korean J. Parasitol. 37(2), 65–70, 1999.
8. Rogan, M.T., Craig, P.S., Hide, G., Heath, S., Pickles, A., and Storey, D.M. The occurrence of the trematode Plagiorchis muris in the wood mouse Apodemus sylvaticus in North Yorkshire, UK. J. Helminthol. 81(1), 57–62, 2007.
9. Sato, H., and Suzuki, K. Gastrointestinal helminths of feral raccoons (Procyon lotor) in Wakayama Prefecture, Japan. J. Vet. Med. Sci. 68(4), 311–318, 2006. 10. Sohn, W.M., and Chai, J.Y. Infection status with helminthes in feral cats purchased from a market in Busan, Republic of Korea. Korean J. Parasitol. 43(3), 93–100, 2005. 11. Radomyos, P., Bunnag, D., and Harinasuta, T. A new intestinal �uke, Plagiorchis harinasutai n.sp. Southeast Asian J. Trop. Med. Public Health 20(1), 101–107, 1989. 12. Sandground, J.H. Plagiorchis javensis n. sp. a new trematode parasitic in man. Rev. Med. Trop. Parasitol. (Habana) 6, 207– 211, 1940. 13. Asada, J.I., Otagaki, H., and Morita, M. et al. A case report on the human infection with Plagiorchis muris Tanabe, 1922 in Japan. Jpn. J. Parasitol. 11, 512–516, 1962. 14. Hong, S.J., Woo, H.C., and Chai, J.Y. A human case of Plagiorchis muris (Tanabe, 1922: Digenea) infection in the Republic of Korea: Freshwater �sh as a possible source of infection. J. Parasitol. 82(4), 647–649, 1996. 15. Yu, S.H., and Mott, K.E. Epidemiology and morbidity of food-borne intestinal trematode infections. Trop. Dis. Bull. 91, R125–R152, 1994. 16. McMullen, D.B. An Experimental infection of Plagiorchis muris in man. J. Parasitol. 23, 113–115, 1937. 17. Chai, J.Y., Yu, J.R., Lee, S.H. et al. An egg-negative patient of acute metagonimiasis diagnosed serologically by ELISA. Seoul. J. Med. 30, 139–142, 1989. 18. Lee, S.U., Huh, S., and Sohn, W.M. Molecular phylogenic location of the Plagiorchis muris (Digenea, Plagiorchiidae) based on sequences of partial 28S D1 rDNA and mitochondrial cytochrome C oxidase subunit I. Korean J. Parasitol. 42(2), 71–75, 2004. 19. Qu, L.H., Nicoloso, M., and Bachellerie, J.P. Phylogenetic calibration of the 5′ terminal domain of large rRNA achieved by determining twenty eucaryotic sequences. J. Mol. Evol., 28(1–2), 113–124, 1988. 41 Chapter 41 - Schistosoma and Schistosomiasis
1. Vester, U. et al. Urinary tract morbidity due to Schistosoma haematobium infection in Mali. Kidney Int. 52, 478–481, 1997.
2. Cox, F.E. History of human parasitic diseases. Infect. Dis. Clin. North Am. 18, 171–188, table of contents, 2004.
3. Ohmae, H. et al. Schistosomiasis mekongi: From discovery to control. Parasitol. Int. 53, 135–142, 2004.
4. Webster, B.L., Southgate, V.R., and Littlewood, D.T. A revision of the interrelationships of Schistosoma including the recently described Schistosoma guineensis. Int. J. Parasitol. 36, 947–955, 2006.
5. Alarcón de Noya, B., and Noya, O. Schistosomosis. In: Parasitología Médica. México, 2008.
6. Rollinson, D., and Southgate, V.R. The genus Schistosoma: A taxonomic appraisal. In: Rollinson, D., and Simpson, A.J.G. (eds), The Biology of Schistosomes. From Genes to Latrines. Academic Press, London, 1987.
7. Morgan, J.A. et al. A newly-identi�ed lineage of Schistosoma. Int. J. Parasitol. 33, 977–985, 2003.
8. Berriman, M. et al. The genome of the blood �uke Schistosoma mansoni. Nature 460, 352–358, 2009.
9. Liu, F. et al. The Schistosoma japonicum genome reveals features of host-parasite interplay. Nature 460, 345–351, 2009.
10. LoVerde, P.T. et al. Schistosoma mansoni genome project: An update. Parasitol. Int. 53, 183–92, 2004.
11. Adams, M.D. et al. Complementary DNA sequencing: Expressed sequence tags and human genome project. Science 252, 1651–1656, 1991.
12. Verjovski-Almeida, S. et al. Transcriptome analysis of the acoelomate human parasite Schistosoma mansoni. Nat. Genet. 35, 148–157, 2003.
13. Hu, W. et al. Evolutionary and biomedical implications of a Schistosoma japonicum complementary DNA resource. Nat. Genet. 35, 139–147, 2003.
14. Wilkins, M.R. et al. Progress with proteome projects: Why all proteins expressed by a genome should be identi�ed and how to do it. Biotechnol. Genet. Eng. Rev. 13, 19–50, 1996. 15. Curwen, R.S. et al. The Schistosoma mansoni soluble proteome: A comparison across four life-cycle stages. Mol. Biochem. Parasitol. 138, 57–66, 2004. 16. Cheng, G.F. et al., Proteomic analysis of differentially expressed proteins between the male and female worm of Schistosoma japonicum after pairing. Proteomics 5, 511–521, 2005. 17. Braschi, S., and Wilson, R.A. Proteins exposed at the adult schistosome surface revealed by biotinylation. Mol. Cell Proteomics 5, 347–356, 2006. 18. Liu, F. et al. Insight into the host-parasite interplay by proteomic study of host proteins copuri�ed with the human parasite, Schistosoma japonicum. Proteomics 7, 450–462, 2007. 19. Guillou, F. et al. Excretory-secretory proteome of larval Schistosoma mansoni and Echinostoma caproni, two parasites of Biomphalaria glabrata. Mol. Biochem. Parasitol. 155, 45–56, 2007. 20. Dvorak, J. et al. Differential use of protease families for invasion by schistosome cercariae. Biochimie 90, 345–358, 2008. 21. Cass, C.L. et al. Proteomic analysis of Schistosoma mansoni egg secretions. Mol. Biochem. Parasitol. 155, 84–93, 2007. 22. King, C.H., Dickman, K., and Tisch, D.J. Reassessment of the cost of chronic helmintic infection: A meta-analysis of disability-related outcomes in endemic schistosomiasis. Lancet 365, 1561–1569, 2005. 23. Corachan, M. et al. Hematospermia: A new etiology of clinical interest. Am. J. Trop. Med. Hyg. 50, 580–584, 1994. 24. Freedman, D.O. et al. Spectrum of disease and relation to place of exposure among ill returned travelers. N. Engl. J. Med. 354, 119–130, 2006. 25. Pardo, J. et al. Utility of Schistosoma bovis adult worm antigens for diagnosis of human schistosomiasis by enzymelinked immunosorbent assay and electroimmunotransfer blot techniques. Clin. Diagn. Lab. Immunol. 11, 1165–1170, 2004. 26. Leshem, E. et al. Acute schistosomiasis outbreak: Clinical features and economic impact. Clin. Infect. Dis. 47, 1499–1506, 2008. 27. Jelinek, T., Nothdurft, H.D., and Loscher, T. Schistosomiasis in travelers and expatriates. J. Travel. Med. 3, 160–164, 1996. 28. Carranza-Rodriguez, C. et al. Cryptic parasite infection in recent West African immigrants with relative eosinophilia. Clin. Infect. Dis. 46, e48–e50, 2008. 29. Janeway, C.A., Jr., and Medzhitov, R. Innate immune recognition. Annu. Rev. Immunol. 20, 197–216, 2002. 30. Akira, S. Toll-like receptor signaling. J. Biol. Chem. 278, 38105–38108, 2003. 31. Beutler, B. Inferences, questions and possibilities in Toll-like receptor signalling. Nature 430, 257–263, 2004. 32. Jenkins, S.J. et al. Schistosome larvae stimulate macrophage cytokine production through TLR4-dependent and -independent pathways. Int. Immunol. 17, 1409–1418, 2005. 33. van Kooyk, Y., Engering, A., Lekkerkerker, A.N., Ludwig, I.S., and Geijtenbeek, T.B. Pathogens use carbohydrates to escape immunity induced by dendritic cells. Curr. Opin. Immunol. 16, 488–493, 2004. 34. Araujo, M.I. et al. Evidence of a T helper type 2 activation in human schistosomiasis. Eur. J. Immunol. 26, 1399–1403, 1996. 35. Caldas, I.R. et al. Human schistosomiasis mansoni: Immune responses during acute and chronic phases of the infection. Acta Trop. 108, 109–117, 2008. 36. van der Kleij, D. et al. A novel host-parasite lipid cross-talk. Schistosomal lyso-phosphatidylserine activates toll-like receptor 2 and affects immune polarization. J. Biol. Chem. 277, 48122–48189, 2002.
37. Montenegro, S.M. et al. Cytokine production in acute versus chronic human Schistosomiasis mansoni: The cross-regulatory role of interferon-gamma and interleukin-10 in the responses of peripheral blood mononuclear cells and splenocytes to parasite antigens. J. Infect. Dis. 179, 1502–1514, 1999.
38. Davies, S.J. et al. Modulation of blood �uke development in the liver by hepatic CD4 + lymphocytes. Science 294, 1358–1361, 2001.
39. Brunet, L.R. et al. IL-4 protects against TNF-alpha-mediated cachexia and death during acute schistosomiasis. J. Immunol. 159, 777–785, 1997.
40. Fallon, P.G. et al. Schistosome infection of transgenic mice de�nes distinct and contrasting pathogenic roles for IL-4 and IL-13: IL-13 is a pro�brotic agent. J. Immunol. 164, 2585–2591, 2000.
41. Cheever, A.W. et al. Analysis of egg granuloma formation in Schistosoma japonicum-infected mice treated with antibodies to interleukin-5 and gamma interferon. Infect. Immun. 59, 4071–4074, 1991.
42. de Jesus, A.R. et al. Association of type 2 cytokines with hepatic �brosis in human Schistosoma mansoni infection. Infect. Immun. 72, 3391–3397, 2004.
43. Hesse, M. et al. Differential regulation of nitric oxide synthase-2 and arginase-1 by type 1/type 2 cytokines in vivo: Granulomatous pathology is shaped by the pattern of l-arginine metabolism. J. Immunol. 167, 6533–6544, 2001.
44. Rumbley, C.A. et al. Activated eosinophils are the major source of Th2-associated cytokines in the schistosome granuloma. J. Immunol. 162, 1003–1009, 1999.
45. Pardo, J. et al. Helminth-related eosinophilia in African immigrants, Gran Canaria. Emerg. Infect. Dis. 12, 1587–1589, 2006.
46. Rutitzky, L.I., Lopes da Rosa, J.R., and Stadecker, M.J. Severe CD4 T cell-mediated immunopathology in murine schistosomiasis is dependent on IL-12p40 and correlates with high levels of IL-17. J. Immunol. 175, 3920–3926, 2005.
47. Rutitzky, L.I. et al. IL-23 is required for the development of severe egg-induced immunopathology in schistosomiasis and for lesional expression of IL-17. J. Immunol. 180, 2486–2495, 2008.
48. Hoffmann, K.F., Cheever, A.W., and Wynn, T.A. IL-10 and the dangers of immune polarization: Excessive type 1 and type 2 cytokine responses induce distinct forms of lethal immunopathology in murine schistosomiasis. J. Immunol. 164, 6406–6416, 2000.
49. Marquet, S. et al. Full results of the genome-wide scan which localises a locus controlling the intensity of infection by Schistosoma mansoni on chromosome 5q31-q33. Eur. J. Hum. Genet. 7, 88–97, 1999.
50. Burke, M.L. et al. Immunopathogenesis of human schistosomiasis. Parasite Immunol. 31, 163–176, 2009.
51. Dessein, A.J. et al. Severe hepatic �brosis in Schistosoma mansoni infection is controlled by a major locus that is closely linked to the interferon-gamma receptor gene. Am. J. Hum. Genet. 65, 709–721, 1999.
52. Blanton, R.E. et al. Schistosomal hepatic �brosis and the interferon gamma receptor: A linkage analysis using singlenucleotide polymorphic markers. Eur. J. Hum. Genet. 13, 660–668, 2005.
53. Henri, S. et al. Cytokine regulation of periportal �brosis in humans infected with Schistosoma mansoni: IFN-gamma is associated with protection against �brosis and TNF-alpha with aggravation of disease. J. Immunol. 169, 929–936, 2002. 54. Chevillard, C. et al. IFN-gamma polymorphisms (IFNgamma +2109 and IFN-gamma +3810) are associated with severe hepatic �brosis in human hepatic schistosomiasis (Schistosoma mansoni). J. Immunol. 171, 5596–5601, 2003. 55. Arnaud, V. et al. Regulatory role of interleukin-10 and interferon-gamma in severe hepatic central and peripheral �brosis in humans infected with Schistosoma japonicum. J. Infect. Dis. 198, 418–426, 2008. 56. Meltzer, E. et al. Schistosomiasis among travelers: New aspects of an old disease. Emerg. Infect. Dis. 12, 1696–1700, 2006. 57. Ross, A.G., Vickers, D., Olds, G.R., Shah, S.M., and McManus, D.P. Katayama syndrome. Lancet Infect. Dis. 7, 218–224, 2007. 58. Bica, I., Hamer, D.H., and Stadecker, M.J. Hepatic schistosomiasis. Infect. Dis. Clin. North Am. 14, 583–604, viii, 2000. 59. Gryseels, B. et al. Human schistosomiasis. Lancet 368, 1106–1118, 2006. 60. Bethlem, E.P., Schettino Gde, P., and Carvalho, C.R. Pulmonary schistosomiasis. Curr. Opin. Pulm. Med. 3, 361–365, 1997. 61. Barsoum, R. The changing face of schistosomal glomerulopathy. Kidney Int. 66, 2472–2484, 2004. 62. Poggensee, G. et al. Female genital schistosomiasis of the lower genital tract: Prevalence and disease-associated morbidity in northern Tanzania. J. Infect. Dis. 181, 1210–1213, 2000. 63. Ferrari, T.C. Involvement of central nervous system in the schistosomiasis. Mem. Inst. Oswaldo Cruz. 99, 59–62, 2004. 64. Nobre, V. et al. Schistosomal myeloradiculopathy due to Schistosoma mansoni: Report on 23 cases. Mem. Inst. Oswaldo Cruz. 96(Suppl), 137–141, 2001. 65. Yosry, A. Schistosomiasis and neoplasia. Contrib. Microbiol. 13, 81–100, 2006. 66. Rao, M.R. et al. Further evidence for association of hepatitis C infection with parenteral schistosomiasis treatment in Egypt. BMC Infect. Dis. 2, 29, 2002. 67. Kamal, S.M. et al. Kinetics of intrahepatic hepatitis C virus (HCV)-speci�c CD4 + T cell responses in HCV and Schistosoma mansoni coinfection: Relation to progression of liver �brosis. J. Infect. Dis. 189, 1140–1150, 2004. 68. Karanja, D. M. et al. Resistance to reinfection with Schistosoma mansoni in occupationally exposed adults and effect of HIV-1 co-infection on susceptibility to schistosomiasis: A longitudinal study. Lancet 360, 592–596, 2002. 69. Teixeira, R. et al. Schistosomiasis mansoni is associated with pyogenic liver abscesses in the state of Minas Gerais, Brazil. Mem. Inst. Oswaldo Cruz. 96(Suppl), 143–146, 2001. 70. Gendrel, D. et al. Nontyphoidal salmonellal septicemia in Gabonese children infected with Schistosoma intercalatum. Clin. Infect. Dis. 18, 103–105, 1994. 71. Katz, N., Chaves, A., and Pellegrino, J. A simple device for quantitative stool thick-smear technique in Schistosomiasis mansoni. Rev. Inst. Med. Trop. Sao Paulo. 14, 397–400, 1972. 72. Gyorkos, T.W. et al. Ef�cacy of new low-cost �ltration device for recovering Schistosoma haematobium eggs from urine. J. Clin. Microbiol. 39, 2681–2682, 2001. 73. Berhe, N. et al. Variations in helminth faecal egg counts in Kato-Katz thick smears and their implications in assessing infection status with Schistosoma mansoni. Acta Trop. 92, 205–212, 2004. 74. Oliver-Gonzalez, J. Anti-egg precipitins in the serum of humans infected with Schistosoma mansoni. J. Infect. Dis. 95, 86–91, 1954. 75. van Lieshout, L., Polderman, A.M., and Deelder, A.M. Immunodiagnosis of schistosomiasis by determination of the circulating antigens CAA and CCA, in particular in individuals with recent or light infections. Acta Trop. 77, 69–80, 2000.
76. Stothard, J.R. et al. An evaluation of urine-CCA strip test and �ngerprick blood SEA-ELISA for detection of urinary schistosomiasis in schoolchildren in Zanzibar. Acta Trop. 111, 64–70, 2009.
77. Doenhoff, M.J., Chiodini, P.L., and Hamilton, J.V. Speci�c and sensitive diagnosis of schistosome infection: Can it be done with antibodies? Trends Parasitol. 20, 35–39, 2004.
78. Pardo, J. et al. Application of an ELISA test using Schistosoma bovis adult worm antigens in travellers and immigrants from a schistosomiasis endemic area and its correlation with clinical �ndings. Scand. J. Infect. Dis. 39, 435–440, 2007.
79. Makarova, E. et al. Detection of IgG binding to Schistosoma mansoni recombinant protein RP26 is a sensitive and speci�c method for acute schistosomiasis diagnosis. Parasitol. Int. 54, 69–74, 2005.
80. Hamburger, J. et al. Polymerase chain reaction assay based on a highly repeated sequence of Schistosoma haematobium: A potential tool for monitoring schistosome-infested water. Am. J. Trop. Med. Hyg. 65, 907–911, 2001.
81. Rabello, A., Pontes, L.A., and Dias-Neto, E. Recent advances in the diagnosis of Schistosoma infection: The detection of parasite DNA. Mem. Inst. Oswaldo Cruz .97(Suppl 1), 171–172, 2002.
82. Pontes, L.A., Dias-Neto, E., and Rabello, A. Detection by polymerase chain reaction of Schistosoma mansoni DNA in human serum and feces. Am. J. Trop. Med. Hyg. 66, 157–162, 2002.
83. Pontes, L.A. et al. Comparison of a polymerase chain reaction and the Kato-Katz technique for diagnosing infection with Schistosoma mansoni. Am. J. Trop. Med. Hyg. 68, 652–656, 2003. 84. Sandoval, N. et al. A new PCR-based approach for the speci�c ampli�cation of DNA from different Schistosoma species applicable to human urine samples. Parasitology 133, 581–587, 2006. 85. Sandoval, N. et al. Schistosoma mansoni: A diagnostic approach to detect acute schistosomiasis infection in a murine model by PCR. Exp. Parasitol. 114, 84–88, 2006. 86. Akinwale, O.P. et al. Detection of schistosomes polymerase chain reaction ampli�ed DNA by oligochromatographic dipstick. Mol. Biochem. Parasitol. 160, 167–170, 2008. 87. Wichmann, D. et al. Diagnosing schistosomiasis by detection of cell-free parasite DNA in human plasma. PLoS Negl. Trop. Dis. 3, e422, 2009. 88. Fenwick, A. et al. Drugs for the control of parasitic diseases: Current status and development in schistosomiasis. Trends Parasitol. 19, 509–515, 2003. 89. Pérez Arellano, J.L. et al. Tratamiento de las enfermedades parasitarias (II): Helmintosis y ectoparasitosis. Información Terapéutica del Sistema Nacional de Salud 31, 55, 2007. 90. Xiao, S.H. Development of antischistosomal drugs in China, with particular consideration to praziquantel and the artemisinins. Acta Trop. 96, 153–167, 2005. 91. Keiser, J. et al. Me�oquine-an aminoalcohol with promising antischistosomal properties in mice. PLoS Negl. Trop. Dis. 3, e350, 2009. 92. McManus, D.P., and Loukas, A. Current status of vaccines for schistosomiasis. Clin. Microbiol. Rev. 21, 225–242, 2008. 42 Chapter 42 - Trichobilharzia
TABLE 42.3
PCR Conditions for the Amplification of Schistosome DNA
Cycle Step Temp. (°C) Time (s) Number of Cycles
1 1 95 240 1
2 1 94 30 3 2 55 30 3 72 120
3 1 94 30 3 2 54 30 3 72 120
4 1 94 30 3 2 53 30 3 72 120
5 1 94 30 3 2 52 30 3 72 120
6 1 94 30 3 2 51 30 3 72 120
7 1 94 30 20 2 50 30 3 72 150
8 1 72 300 1
3. Kouřilová, P., Syrůček, M., and Kolářová, L. The severity of mouse pathologies caused by the bird schistosome Trichobilharzia regenti in relation to host immune status. Parasitol. Res. 93, 534, 2004.
4. Skírnisson, K., and Kolářová, L. Diversity of bird schistosomes in anseriform birds in Iceland based on egg measurements and egg morphology. Parasitol. Res. 103, 43, 2008.
5. Podhorský, M. et al. Cercarial dimensions and surface structures as a tool for species determination of Trichobilharzia spp. Acta Parasitol. 54, 28, 2009.
6. Aldhoun, J.A. et al. Bird schistosome diversity in Iceland: Molecular evidence. J. Helminthol. 83, 173, 2009.
7. Brant, S.V., and Loker, E. S. Molecular systematics of the avian schistosome genus Trichobilharzia (Trematoda: Schistosomatidae) in North America. J. Parasitol. 95, 941, 2009.
8. Jouet, D. et al. Avian schistosomes in French aquatic birds: A molecular approach. J. Helminthol. 83, 181, 2009.
9. Rudolfová, J. et al. Validity reassessment of Trichobilharzia species using Lymnaea stagnalis as the intermediate host. Parasitol. Res. 95, 79, 2005.
10. Brant, S.V. et al. An approach to revealing blood �uke life cycles, taxonomy, and diversity: Provision of key reference data including DNA sequence from single life cycle stages. J. Parasitol. 92, 77, 2006.
11. Brant, S.V., and Loker, E.S. Schistosomes in the southwest United States and their potential for causing cercarial dermatitis or swimmer’s itch. J. Helminthol. 83, 191, 2009.
12. Aldhoun J.A. et al. Schistosomes in the North: A unique �nding from a prosobranch snail using molecular tools. Parasitol. Int. 58, 314, 2009.
13. Dvořák, J. et al. Comparison of European Trichobilharzia species based on ITS1 and ITS2 sequences. Parasitology 124, 307, 2002.
14. Bayssade-Dufour, C. et al. Seasonal morphological variations in bird schistosomes. Parasite 13, 205, 2006.
15. Kolářová, L. et al. Allobilharzia visceralis gen. nov. sp. nov. (Schistosomatidae-Trematoda) from Cygnus cygnus (L.) (Anatidae). Parasitol. Int. 55, 179, 2006.
16. Jouet, D. et al. Trichobilharzia spp. in natural conditions in Annecy Lake, France. Parasitol. Res. 103, 51, 2008.
17. Chrisanfova, G.G. et al. Genetic variability of bird schistosomes (class Trematoda, family Schistosomatidae) of Naroch lake: Identi�cation of a new species in the Trichobilharzia ocellata group. Doklady Biochem. Biophys. 428, 268, 2009.
18. Ferte, H. et al. Presence of Trichobilharzia szidati in Lymnaea stagnalis and T. franki in Radix auricularia in northeastern France: Molecular evidence. Parasitol. Res. 95, 150, 2005.
19. Picard, D., and Jousson, O. Genetic variability among cercariae of the Schistosomatidae (Trematoda: Digenea) causing swimmer’s itch in Europe. Parasite 8, 237, 2001.
20. Semyenova, S.K. et al. Individual and population variation in cercariae of bird schistosomes of the Trichobilharzia ocellata species group as revealed with the polymerase chain reaction. Russ. J. Genet. 41, 12, 2005.
21. Arnot, D.E., Roper, C., and Bayoumi, R.A.L. Digital codes from hypervariable tandemly repeated DNA sequences in the Plasmodium falciparum circumsporozoite gene can genetically barcode isolates. Mol. Biochem. Parasitol. 61, 15, 1993.
22. Nolan, M.J., and Cribb, T.J. The use and implications of ribosomal DNA sequencing for the discrimination of digenean species. Adv. Parasitol. 60, 101, 2005.
23. Vilas, R., Criscione, C., and Blouin, M. A comparison between mitochondrial DNA and the ribosomal internal transcribed regions in prospecting for cryptic species of platyhelminth parasites. Parasitology 131, 839, 2005. 24. Zarowiecki, M.Z., Huyse, T., and Littlewood, D.T.J. Making the most of mitochondrial genomes—Markers for phylogeny, molecular ecology and barcodes in Schistosoma (Platyhelminthes: Digenea). Int. J. Parasitol. 37, 1401, 2007. 25. Hamilton, P.B. et al. A novel, high-throughput technique for species identi�cation reveals a new species of tsetse-transmitted trypanosome related to the Trypanosoma brucei subgenus Trypanozoon. Infect. Genet. Evol. 8, 26, 2008. 26. Ferri, E. et al. Integrated taxonomy: Traditional approach and DNA barcoding for the identi�cation of �larioid worms and related parasites. Front. Zool. 6, 1, 2009. 27. Moszczynska, A. et al. Development of primers for the mitochondrial cytochrome c oxidase I gene in digenetic trematodes (Platyhelminthes) illustrates the challenge of barcoding parasitic helminths. Mol. Ecol. Resources 9, 75, 2009. 28. Besansky, N.J., Severson, D.W., and Ferdig, M.T. DNA barcoding of parasites and invertebrate disease vectors: What you don’t know can hurt you. Trends Parasitol. 19, 545, 2003. 29. Hansen, H., Bakke, T.A., and Bachmann, L. DNA taxonomy and barcoding of monogenean parasites: Lessons from Gyrodactylus. Trends Parasitol. 23, 363, 2007. 30. Cort, W.W. Schistosome dermatitis in the United States (Michigan). J. Am. Med. Assoc. 90, 1027, 1928. 31. Loken, B.R., Spencer, C.N., and Granath, W.R. Jr., Prevalence and transmission of cercariae causing schistosome dermatitis in Flathead Lake, Montana. J. Parasitol. 81, 646, 1995. 32. Kolářová, L., Horák, P., and Sitko, J. Cercarial dermatitis in focus: Schistosomes in the Czech Republic. Helminthologia 34, 127, 1997. 33. Farahnak, A., and Essalat, M. A study on cercarial dermatitis in Khuzestan province, south western Iran. BMC Public Health 3, 35, 2003. 34. Voronin, M.V., and Beer, S.A. Morphological peculiarities of Schistosomatidae cercariae of Trichobilharzia cf. ocellata group occurring in Moscow and Saint-Petersburg populations. Parazitologiia 36, 60, 2002. 35. Bourée, P., and Caumes, E. La dermatitie cercarienne. Presse Med. 33, 490, 2004. 36. Sheng, S.C. et al. Preliminary study on the ecology of Trichobilharzia cercariae in the Huaihe river system. Chin. J. Parasitol. Parasitol. Dis. 22, 349, 2004. 37. Z˙bikowska, E. Infection of snails with bird schistosomes and the threat of swimmer’s itch in selected Polish lakes. Parasitol. Res. 92, 30, 2004. 38. Coady, N.R. et al. Ubiquitous variability in the prevalence of Trichobilharzia stagnicolae (Schistosomatidae) infecting Stagnicola emarginata in three northern Michigan lakes. J. Parasitol. 92, 10, 2006. 39. Cort, W.W. Studies on schistosome dermatitis XI. Status of knowledge after more than twenty years. Am. J. Hyg. 52, 251, 1950. 40. Penner, L.R. Cercaria littorinalinae sp. nov., a dermatitisproducing schistosome larva from the marine snail, Littorina planaxis Philippi. J. Parasitol. 36, 466, 1950. 41. Stunkard, H.W., and Hinchliffe, M.C. The morphology and life history of Microbilharzia variglandis (Miller and Northup, 1926) Stunkard and Hinchliffe, 1951, avian blood �ukes whose larvae cause “swimmer’s itch” of ocean beaches. J. Parasitol. 38, 248, 1952. 42. Leigh, W.H. Cercaria huttoni sp. nov.: A dermatitis-producing schistosome larva from the marine snail, Haminoea antillarum guadalupensis Sowerby. J. Parasitol. 39, 625, 1953.
43. Chu, G.W.T.C. Paci�c area distribution of freshwater and marine cercarial dermatitis. Pac. Sci. 12, 299, 1958.
44. Leedom, W.S., and Short, R.B. Cercaria pomaceae sp. n., a dermatitis-producing schistosome cercaria from Pomacea paludosa, the Florida apple snail. J. Parasitol. 67, 257, 1981.
45. Kolářová, L. Schistosomes causing cercarial dermatitis: A mini-review of current trends in systematics and host speci�city and pathogenicity. Folia Parasitol. 54, 81, 2007.
46. Loy, C., and Haas, W. Prevalence of cercariae from Lymnaea stagnalis snails in a pond system in Southern Germany. Parasitol. Res. 87, 878, 2001.
47. Schets, F.M., Lodder, W.J., and de Roda Husman, A.M. Con�rmation of the presence of Trichobilharzia by examination of water samples and snails following reports of cases of cercarial dermatitis. Parasitology 137, 77, 2010.
48. Skírnisson, K., Aldhoun, J.A., and Kolářová, L. A review on swimmers itch and the occurrence of bird schistosomes in Iceland. J. Helminthol. 83, 165, 2009.
49. Valdovinos, C., and Balboa, C. Cercarial dermatitis and lake eutrophication in South-Central Chile. Epidemiol. Infect. 136, 391, 2008.
50. Kolářová, L. et al. Avian schistosomes of the genus Trichobilharzia in Europe. Bull. Scandinavian-Baltic Soc. Parasitol. 14, 85, 2005.
51. Rudolfová, J., Sitko, J., and Horák, P. Nasal schistosomes in the Czech Republic. Parasitol. Res. 88, 1093, 2002.
52. Caumes, E. et al. Failure of an ointment based on IR3535 (ethylbutylacetylaminopropionate) to prevent an outbreak of cercarial dermatitis during swimming races across Lake Annecy, France. Ann. Trop. Med. Parasitol. 97, 157, 2003.
53. De Gentile, L. et al. La dermatite cercarienne en Europe: Un problème de santé publique nouveau? (Cercarial dermatitis in Europe: A new public health problem?). WHO Bull. 74, 159, 1996.
54. Kolářová, L., Skírnisson, K., and Horák, P. Schistosome cercariae as the causative agent of swimmer’s itch in Iceland. J. Helminthol. 73, 215, 1999.
55. Rao, V.G. et al. Cercarial dermatitis in Central India: An emerging health problem among tribal communities. Ann. Trop. Med. Parasitol. 101, 409, 2007.
56. Hörweg, C., Sattmann, H., and Auer, H. Cercarial dermatitis in Austria: Questionnaires as useful tools to estimate risk factors? Wien. Klin. Wochenschr. 118, 177, 2006.
57. Fraser, S.J. et al. Cercarial dermatitis in the UK. Clin. Exp. Dermatol. 34, 344, 2009.
58. Schets, F.M. et al. Cercarial dermatitis in the Netherlands caused by Trichobilharzia spp. J. Water Health 06, 187, 2008.
59. Wang, C.R. et al. Orientobilharzia species: Neglected zoonotic agents. Acta Trop. 109, 171, 2008.
60. Dubois, J.-P. et al. Epidemiological studies related to cercarial dermatitis in lakes of the Savoy District (France). Helminthologia 38, 244, 2001.
61. Thors, C., and Linder, E. Swimmer’s itch in Sweden. Helminthologia 38, 244, 2001.
62. Lévesque, B. et al. Investigation of an outbreak of cercarial dermatitis. Epidemiol. Infect. 129, 379, 2002.
63. Larsen, A.H., Bresciani, J., and Buchmann, K. Increasing frequency of cercarial dermatitis in higher latitudes. Acta Parasitol. 49, 217, 2004.
64. Mas-Coma, S., Valero, M.A., and Bargues, M.D. Effect of climate change on animal and zoonotic helminthiases. Rev. Sci. Tech. 27, 443, 2008. 65. Skírnisson, K., and Kolářová, L. Are nasal Trichobilharzia cercariae potential threat to human health? Iceland Med. J. 88, 739, 2002. 66. Lindblade, K.A. The epidemiology of cercarial dermatitis and its association with limnological characteristics of a northern Michigan lake. J. Parasitol. 84, 19, 1998. 67. Chamot, E., Toscani, L., and Rougemont, A. Public health importance and risk factors for cercarial dermatitis associated with swimming in Lake Leman at Geneva, Switzerland. Epidemiol. Infect. 120, 305, 1998. 68. Verbrugge, L.M. et al. Prospective study of swimmer’s itch incidence and severity. J. Parasitol. 90, 697, 2004. 69. Hertel, J. et al. Detection of bird schistosomes in lakes by PCR and �lter-hybridisation. Exp. Parasitol. 101, 57, 2002. 70. Kouřilová, P., and Kolářová, L. Variations in immuno�uorescent antibody response against Trichobilharzia and Schistosoma antigens in compatible and noncompatible hosts. Parasitol. Res. 88, 513, 2002. 71. Graczyk, T.K., and Shiff, C.J. Recovery of avian schistosome cercariae from water using penetration stimulant matrix with an unsaturated fatty acid. Am. J. Trop. Med. Hyg. 63, 174, 2000. 72. Peterson, A.T. et al. The need for proper vouchering in phylogenetic studies of birds. Mol. Phylogen. Evol. 45, 1042, 2007. 73. Olsson, U. et al. What is the proper vouchering in phylogenetic studies of birds?—A reply to Peterson et al., 2007. Mol. Phylogen. Evol. 48, 383, 2008. 74. Hoberg, E.P., Pilitt, P.A., and Galbreath, K.E. Why museums matter: A tale of pinworms (Oxyuroidea: Heteroxynematidae) among pikas (Ochotona princeps and O. collaris) in the American West. J. Parasitol. 95, 790, 2009. 75. Olivier, L. Schistosome dermatitis, a sensitization phenomenon. Am. J. Hyg. 49, 290, 1949. 76. Oda, T. Schistosome dermatitis in Japan. In: Morishita, K., Komiya, Y., and Matsubayashi, H. (eds), Progress in Medical Parasitology in Japan, Vol. 2, Meguro Parasitological Museum, Tokyo, pp. 5–53, 1973. 77. Hunter, G.W., Ritchie, L.S., and Tanabe, H. The epidemiology of schistosome dermatitis (“Koganbyo”) in Japan. Trans. R. Soc. Trop. Med. Hyg. 45, 103, 1951. 78. Haemmerli, U. Schistosomen-Dermatitis am Zürichsee. Dermatologica 107, 302, 1953. 79. Kouřilová, P. et al. Cercarial dermatitis caused by bird schistosomes comprises both immediate and late phase cutaneous hypersensitivity reactions. J. Immunol. 172, 3766, 2004. 80. Lichtenbergová, L. et al. Antibody responses induced by Trichobilharzia regenti antigens in murine and human hosts exhibiting cercarial dermatitis. Parasite Immunol. 30, 585, 2008. 81. Kolářová, L., Horák, P., and Čada, F. Histopathology of CNS and nasal infections caused by Trichobilharzia regenti in vertebrates Parasitol. Res. 87, 644, 2001. 82. Chanová, M., Vuong, S., and Horák, P. Trichobilharzia szidati: The lung phase of migration within avian and mammalian hosts. Parasitol. Res. 100, 1243, 2007. 83. Bayssade-Dufour, C., Martins, C., and Vuong, P.N. Histopathologie d’un modèle mammifère et dermatite cercarienne humaine. Méd. Mal. Infect. 31, 713, 2001. 84. Hoef�er, D.F. Swimmers’ itch (cercarial dermatitis). Cutis 19, 461, 1977. 85. Appleton, C.C. Schistosome dermatitis—An unrecognized problem in South Africa. South Afr. Med. J. 65, 467, 1984.
86. Sonnenberg, R., Nolte, A.W., and Tautz, D. An evaluation of LSU rDNA D1-D2 sequences for their use in species identi�cation. Front. Zool. 4, 6, 2007.
87. Webster, B.L. et al. The complete mitochondrial genome of the bird schistosome Trichobilharzia regenti (Platyhelminthes: Digenea), causative agent of cercarial dermatitis. J. Parasitol. 93, 553, 2007.
88. Brant, S.V. The occurrence of the avian schistosome Allobilharzia visceralis Kolářová, Rudolfová, Hampl et Skirnisson, 2006 (Schistosomatidae) in the tundra swan, Cygnus columbianus (Anatidae), from North America. Folia Parasitol. 54, 99, 2007.
89. Buhay, J.E. “CO1-LIKE” sequences are becoming problematic in molecular systematics and DNA barcoding studies. J. Crust. Biol. 29, 96, 2009. 90. Truett, G.E. et al. Preparation of PCR-quality mouse genomic DNA with hot sodium hydroxide and Tris (Hotshot). Biotechniques 29, 52, 2000. 91. Lockyer, A.E. et al. The phylogeny of the Schistosomatidae based on three genes with emphasis on the interrelation ships of Schistosoma Weinland, 1858. Parasitology 126, 203, 2003. 92. Tkach, V.V., Pawlowski, J., and Mariaux, J. Phylogenetic analysis of the suborder Plagiorchiata (Platyhelminthes, Digenea) based on partial lsrDNA sequences. Int. J. Parasitol. 30, 83,, 2000. 93. Mollaret, I. et al. Phylogenetic analysis of the Monogenea and their relationships with Digenea and Eucestoda inferred from 28S rDNA sequences. Mol. Biochem. Parasitol. 90, 433, 1997.
Section V
Nematoda 43 Chapter 43 - Ancylostoma
10. Blotkamp, J. et al. Observations on the morphology of adults and larval stages of Oesophagostomum sp. isolated from man in northern Togo and Ghana. J. Helminthol. 67, 49, 1993.
11. Krepel, H.P. et al. Quantitative interpretation of coprocultures in a population infected with Oesophagostomum bifurcum. Trop. Geogr. Med. 47, 157, 1995.
12. de Gruijter, J.M. et al. Polymerase chain reaction-based differential diagnosis of Ancylostoma duodenale and Necator americanus infections in humans in northern Ghana. Trop. Med. Int. Health 10, 574, 2005.
13. Verweij, J.J. et al. Determining the prevalence of Oesophagostomum bifurcum and Necator americanus infections using speci�c PCR ampli�cation of DNA from faecal samples. Trop. Med. Int. Health 6, 726, 2001.
14. Verweij, J.J. et al. Simultaneous detection and quanti�cation of Ancylostoma duodenale, Necator americanus, and Oesophagostomum bifurcum in fecal samples using multiplex real-time PCR. Am. J. Trop. Med. Hyg. 77, 685, 2007.
15. Harada, Y., and Mori, O. A new method for culturing hookworm. Yonago Acta Med. 1, 177, 1955.
16. Lindo, J.F. et al. Prospective evaluation of enzyme-linked immunosorbent assay and immunoblot methods for the diagnosis of endemic Strongyloides stercoralis infection. Am. J. Trop. Med. Hyg. 51, 175, 1994.
17. Gasser, R.B. Molecular tools—Advances, opportunities and prospects. Vet. Parasitol. 136, 69, 2006.
18. Gasser, R.B. et al. DNA technological progress toward advanced diagnostic tools to support human hookworm control. Biotechnol. Adv. 26, 35, 2008.
19. Hawdon, J.M. Differentiation between the human hookworms Ancylostoma duodenale and Necator americanus using PCRRFLP. J. Parasitol. 82, 642, 1996.
20. Zhan, B. et al. Species-speci�c identi�cation of human hookworms by PCR of the mitochondrial cytochrome oxidase I gene. J. Parasitol. 87, 1227, 2001. 21. Monti, J.R. et al. Speci�c ampli�cation of Necator americanus or Ancylostoma duodenale DNA by PCR using markers in ITS-1 rDNA, and its implications. Mol. Cell Probes 12, 71, 1998. 22. Gasser, R.B. et al. Improved molecular diagnostic tools for human hookworms. Expert Rev. Mol. Diagn. 9, 17, 2009. 23. Hu, M. et al. Single-strand conformation polymorphism-based analysis of mitochondrial cytochrome c oxidase subunit 1 reveals signi�cant substructuring in hookworm populations. Electrophoresis 23, 27, 2002. 24. de Gruijter, J.M. et al. AFLP �ngerprinting for the analysis of genetic diversity within Necator americanus. Mol. Cell Probes 20, 317, 2006. 25. de Gruijter, J.M. et al. Genetic substructuring within Oesophagostomum bifurcum (Nematoda) from human and non-human primates from Ghana based on random ampli�ed polymorphic DNA analysis. Am. J. Trop. Med. Hyg. 71, 227, 2004. 26. de Gruijter, J.M. et al. High resolution DNA �ngerprinting by AFLP to study the genetic variation among Oesophagostomum bifurcum (Nematoda) from human and non-human primates from Ghana. Parasitology 130, 229, 2005. 27. Kaplan, R.M. Drug resistance in nematodes of veterinary importance: A status report. Trends Parasitol. 20, 477, 2004. 28. Schwenkenbecher, J.M. et al. Characterization of beta-tubulin genes in hookworms and investigation of resistance-associated mutations using real-time PCR. Mol. Biochem. Parasitol. 156, 167, 2007. 29. Verweij, J.J. et al. Prevalence of Entamoeba histolytica and Entamoeba dispar in northern Ghana. Trop. Med. Int. Health 8, 1153, 2003. 30. Gasser, R.B. et al. Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res. 21, 2525, 1993. 31. Niesters, H.G. Clinical virology in real time. J. Clin. Virol. 25(Suppl 3), S3, 2002. 44 Chapter 44 - Angiostrongylus
1. Graeff-Teixeira, C., da Silva, A.C.A., and Yoshimura, K., Update on eosinophilic meningoencephalitis and its clinical relevance. Clin. Microbiol. Rev., 22, 322, 2009. 2. Lv, S. et al., Angiostrongylus cantonensis: Morphological and behavioral investigation within the freshwater snail Pomacea canaliculata. Parasitol. Res., 104, 1351, 2009. 3. Guilhon, J., and Cens, B., Angiostrongilus vasorum (Baillet, 1866). Biological and morphological study. Ann. Parasitol. Hum. Comp., 48, 567, 1973. 4. Bolt, G. et al., Canine angiostrongylosis: A review. Vet. Rec., 135, 447, 1994. 5. Ash, L.R., Diagnostic morphology of the third-stage larvae of Angiostrongylus cantonensis, Angiostrongylus vasorum, Aelurostrongylus abstrusus, and Ana�laroides rostratus (Nematoda: Metastrongyloidea). J. Parasitol., 56, 249, 1970. 6. Bonetti, V.C., and Graeff-Teixeira, C., Angiostrongylus costaricensis and the intermediate hosts: Observations on elimination of L3 in the mucus and inoculation of L1 through the tegument of mollucs. Rev. Soc. Bras. Med. Trop., 31, 289, 1998. 7. Ubelaker, J.E., Bullick, G.R., and Caruso, J., Emergence of third-stage larvae of Angiostrongylus costaricensis Morera and Cespedes 1971 from Biomphalaria glabrata (Say). J. Parasitol., 66, 856, 1980. 8. Tesana, S. et al., Prevalence and intensity of infection with third stage larvae of Angiostrongylus cantonensis in mollusks from Northeast Thailand. Am. J. Trop. Med. Hyg., 80, 983, 2009. 9. Thiengo, S.C. et al., The giant African snail Achatina fulica as natural intermediate host of Angiostrongylus cantonensis in Pernambuco, northeast Brazil. Acta Trop., 115, 194, 2010. 10. Vitta, A. et al., Survey of Angiostrongylus cantonensis in rats and giant African land snails in Phitsanulok province, Thailand. Asian Pac. J. Trop. Med., 4, 597, 2011. 11. Ash, L.R., Observations on the role of mollusks and planarians in the transmission of Angiostrongylus cantonensis infection to man in New Caledonia. Rev. Biol. Trop., 24, 163, 1976. 12. Kliks, M.M., and Palumbo, N.E., Eosinophilic meningitis beyond the Paci�c Basin: The global dispersal of a peridomestic zoonosis caused by Angiostrongylus cantonensis, the nematode lungworm of rats. Soc. Sci. Med., 34, 199, 1992. 13. Chung, H., A letter regarding a case of eosinophilic meningitis. Hawaii Med. J., 64, 60, 2005. 14. Hochberg, N.S. et al., Distribution of eosinophilic meningitis cases attributable to Angiostrongylus cantonensis, Hawaii. Emerg. Infect. Dis., 13, 1675, 2007. 15. Hochberg, N.S. et al., Eosinophilic meningitis attributable to Angiostrongylus cantonensis infection in Hawaii: Clinical characteristics and potential exposures. Am. J. Trop. Med. Hyg., 85, 685, 2011. 16. Lindo, J.F. et al., Enzootic Angiostrongylus cantonensis in rats and snails after an outbreak of human eosinophilic meningitis, Jamaica. Emerg. Infect. Dis., 8, 324, 2002. 17. Slom, T.J. et al., An outbreak of eosinophilic meningitis caused by Angiostrongylus cantonensis in travelers returning from the Caribbean. N. Engl. J. Med., 346, 668, 2002. 18. Raccurt, C.P., Blaise, J., and Durette-Desset, M.C., Presence of Angiostrongylus cantonensis in Haiti. Trop. Med. Int. Health, 8, 423, 2003. 19. Chen, X.G., Li, H., and Lun, Z.R., Angiostrongyliasis, mainland China. Emerg. Infect. Dis., 11, 1645, 2005. 20. Lv, S. et al., Emerging angiostrongyliasis in Mainland China. Emerg. Infect. Dis., 14, 161, 2008. 21. Lv, S. et al., Human angiostrongyliasis outbreak in Dali, China. PLoS Negl. Trop. Dis., 3, e520, 2009. 22. Lv, S. et al., Invasive snails and an emerging infectious disease: Results from the �rst national survey on Angiostrongylus cantonensis in China. PLoS Negl. Trop. Dis., 3, e368, 2009.
23. Wang, J. et al., An outbreak of angiostrongyliasis cantonensis in Beijing. J. Parasitol., 96, 377, 2010.
24. Chen, D. et al., Epidemiological survey of Angiostrongylus cantonensis in the west-central region of Guangdong Province, China. Parasitol. Res., 109, 305, 2011.
25. Deng, Z.H. et al., First provincial survey of Angiostrongylus cantonensis in Guangdong Province, China. Trop. Med. Int. Health, 2011 Sep 9.
26. Hu, X. et al., Epidemic status of Angiostrongylus cantonensis in Hainan Island, China. Asian Pac. J. Trop. Med., 4, 275, 2011.
27. Campbell, B.G., and Little, M.D., The �nding of Angiostrongylus cantonensis in rats in New Orleans. Am. J. Trop. Med. Hyg., 38, 568, 1988.
28. Costa, L. et al., Verminous meningoencephalomyelitis by Angiostrongylus (= Parastrongylus) cantonensis in an American miniature horse. Equine Vet. Educ., 12, 2, 2000.
29. Duffy, M.S. et al., Parastrongylus cantonensis in a nonhuman primate, Florida. Emerg. Infect. Dis., 10, 2207, 2004.
30. Maldonado, Jr., A. et al., First report of Angiostrongylus cantonensis (Nematoda: Metastrongylidae) in Achatina fulica (Mollusca: Gastropoda) from Southeast and South Brazil. Mem. Inst. Oswaldo Cruz, 105, 938, 2010.
31. Archer, C.E. et al., The rat lung-worm Angiostrongylus cantonensis: A �rst report in South Africa. S. Afr. Med. J., 101, 174, 2011.
32. New, D., Little, M.D., and Cross, J., Angiostrongylus cantonensis infection from eating raw snails. N. Engl. J. Med., 332, 1105, 1995.
33. Marsh, C.M., Eosinophilic meningitis/angiostrongyliasis from eating aquaculture-raised snails: A case report. Hawaii Med. J., 57, 652, 1998.
34. Lai, C.H. et al., Eosinophilic meningitis caused by Angiostrongylus cantonensis after ingestion of raw frogs. Am. J. Trop. Med. Hyg., 76, 399, 2007.
35. Sawanyawisuth, K. et al., Clinical factors predictive of encephalitis caused by Angiostrongylus cantonensis. Am. J. Trop. Med. Hyg., 81, 698, 2009.
36. Tsai, H.C. et al., Encephalitis caused by Angiostrongylus cantonensis after eating raw frogs mixed with wine as a health supplement. Intern. Med., 50, 771, 2011.
37. Wang, J.J. et al., Eosinophilic meningitis risk associated with raw Ampullarium canaliculatus snails consumption. Kaohsiung J. Med. Sci., 27, 184, 2011.
38. Anonimous, Predictive factors for encephalitis caused by Angiostrongylus cantonensis infection. Am. J. Trop. Med. Hyg., 82, 516, 2010; author reply, 82, 517, 2010.
39. Gelis, S., Spratt, D.M., and Raidal, S.R., Neuroangiostrongyliasis and other parasites in tawny frogmouths (Podargus strigoides) in south-eastern Queensland. Aust. Vet. J., 89, 47, 2011.
40. Mota, E.M., and Lenzi, H.L., Angiostrongylus costaricensis life cycle: A new proposal. Mem. Inst. Oswaldo Cruz, 90, 707, 1995.
41. Mota, E.M., and Lenzi, H.L., Angiostrongylus costaricensis: Complete redescription of the migratory pathways based on experimental Sigmodon hispidus infection. Mem. Inst. Oswaldo Cruz, 100, 407, 2005.
42. Hulbert, T.V., Larsen, R.A., and Chandrasoma, P.T., Abdominal Angiostrongyliasis mimicking acute appendicitis and Meckel’s diverticulum: Report of a case in the United States and review. Clin. Infect. Dis., 14, 836, 1992.
43. Graeff-Teixeira, C. et al., Longitudinal clinical and serological survey of abdominal angiostrongyliasis in Guaporé, southern Brazil, from 1995 to 1999. Rev. Soc. Bras. Med. Trop., 38, 310, 2005. 44. Alicata, J.E., Angiostrongylus cantonensis and eosinophilic meningitis. Lancet, 2, 1367, 1988. 45. Panackel, C. et al., Eosinophilic meningitis due to Angiostrongylus cantonensis. Indian J. Med. Microbiol., 24, 220, 2006. 46. Li, H. et al., A severe eosinophilic meningoencephalitis caused by infection of Angiostrongylus cantonensis. Am. J. Trop. Med. Hyg., 79, 568, 2008. 47. Tsai, H.C. et al., Dynamic changes of hepatocyte growth factor in eosinophilic meningitis caused by Angiostrongylus cantonensis infection. Am. J. Trop. Med. Hyg., 80, 980, 2009. 48. Lee, I.C., Angiostrongylus cantonensis meningitis in two developmentally delayed children: Findings in brain images. Pediatr. Infect. Dis. J., 29, 90, 2010. 49. Cui, Y., Shen, M., and Meng, S., Lung CT �ndings of angiostrongyliasis cantonensis caused by Angiostrongylus cantonensis. Clin. Imaging, 35, 180, 2011. 50. Prociv, P., Spratt, D.M., and Carlisle, M.S., Neuroangiostrongyliasis: Unresolved issues. Int. J. Parasitol., 30, 1295, 2000. 51. Maretić, T. et al., Meningitis and radiculomyelitis caused by Angiostrongylus cantonensis. Emerg. Infect. Dis., 15, 996, 2009. 52. Dorta-Contreras, A.J. et al., Neuroimmunological �ndings of Angiostrongylus cantonensis meningitis in Ecuadorian patients. Arq. Neuropsiquiatr., 69, 466, 2011. 53. Wang, Q.P. et al., Human Angiostrongylus cantonensis: An update. Eur. J. Clin. Microbiol. Infect. Dis., 2011 Jul 5 [Epub ahead of print]. 54. Alto, W., Human infections with Angiostrongylus cantonensis. Pac. Health Dialog., 8, 176, 2001. 55. Diao, Z. et al., Human ocular angiostrongyliasis: A literature review. Trop. Doct., 41, 76, 2011. 56. Liu, I.H. et al., Necrotizing retinitis induced by Angiostrongylus cantonensis. Am. J. Ophthalmol., 141, 577, 2006. 57. Diao, Z. et al., International symposium on Angiostrongylus and Angiostrongyliasis, 2010. Emerg. Infect. Dis., 17, e1, 2011. 58. Loría-Cortés, R., and Lobo-Sanahuja, J.F., Clinical abdominal angiostrongyliasis. A study of 116 children with intestinal eosinophilic granuloma caused by Angiostrongylus costaricensis. Am. J. Trop. Med. Hyg., 29, 538, 1980. 59. Morera, P., Abdominal angiostrongyliasis: A problem of public health. Parasitol. Today, 1, 173, 1985. 60. Kramer, M.H. et al., First reported outbreak of abdominal angiostrongyliasis. Clin. Infect. Dis., 26, 365, 1998. 61. Rodriguez, R. et al., Abdominal angiostrongyliasis: Report of two cases with different clinical presentations. Rev. Inst. Med. Trop. Sao Paulo, 50, 339, 2008. 62. Incani, R.N. et al., Human infection by Angiostrongylus costaricensis in Venezuela: First report of a con�rmed case. Rev. Inst. Med. Trop. Sao Paulo, 49, 197, 2007. 63. Quirós, J.L. et al., Abdominal angiostrongyliasis with involvement of liver histopathologically con�rmed: A case report. Rev. Inst. Med. Trop. Sao Paulo, 53, 219, 2011. 64. Sawanyawisuth, K., and Sawanyawisuth, K., Drug target in eosinophilic meningitis caused by Angiostrongylus cantonensis. Infect. Disord. Drug Targets, 10, 322, 2010. 65. Liu, J., Gao, J., and Zhou, C., A case report on eosinophilic meningitis caused by Angiostrongylus cantonensis. Int. J. Med. Sci., 8, 510, 2011. 66. He, H.J. et al., Ef�cacy of combined treatment with albendazole and baicalein against eosinophilic meningitis induced by Angiostrongylus cantonensis in mice. J. Helminthol., 85, 92, 2011.
67. Tsai, H.C. et al., Brain magnetic resonance imaging abnormalities in eosinophilic meningitis caused by Angiostrongylus cantonensis infection. Vector Borne Zoonotic Dis., 12, 161, 2012.
68. Walker, M.D., and Zunt, J.R., Neuroparasitic infections: Nematodes. Semin. Neurol., 25, 252, 2005.
69. Ramirez-Avila, L. et al., Eosinophilic meningitis due to Angiostrongylus and Gnathostoma species. Clin. Infect. Dis., 48, 322, 2009.
70. Kuberski, T. et al., Recovery of Angiostrongylus cantonensis from cerebrospinal �uid of a child with eosinophilic meningitis. J. Clin. Microbiol., 9, 629, 1979.
71. Graeff-Texeira, C., Camillo-Coura, L., and Lenzi, H.L., Histopathological criteria for the diagnosis of Angiostrongyliasis. Parasitol. Res., 77, 606, 1991.
72. Vitta, A. et al., Application of recombinant SMR-domain containing protein of angiostrongylus cantonensis in immunoblot diagnosis of human angiostrongyliasis. Southeast Asian J. Trop. Med. Public Health, 41, 785, 2010.
73. Ben, R. et al., Use of heterologous antigens for the immunodiagnosis of abdominal angiostrongyliasis by an enzyme-linked immunosorbent assay. Mem. Inst. Oswaldo Cruz, 105, 914, 2010.
74. Chen, M.X. et al., Seroprevalence of Angiostrongylus cantonensis infection in humans in China. J. Parasitol., 97, 144, 2011.
75. Chen, M.X. et al., Development of a double antibody sandwich ELISA assay for the diagnosis of angiostrongyliasis. J. Parasitol., 97, 721, 2011.
76. Eamsobhana, P., and Hong, H.S., Immunological diagnosis of human angiostrongyliasis due to Angiostrongylus cantonensis (Nematoda: Angiostrongylidae). Int. J. Infect. Dis., 13, 425, 2009.
77. Caldeira, R.L., Molecular differentiation of Angiostrongylus costaricensis, A. cantonensis, and A. vasorum by polymerase chain reaction-restriction fragment length polymorphism. Mem. Inst. Oswaldo Cruz, 98, 1039, 2003.
78. da Silva, A.C., Graeff-Teixeira, C., and Zaha, A., Diagnosis of abdominal angiostrongyliasis by PCR from sera of patients. Rev. Inst. Med. Trop. Sao Paulo, 45, 295, 2003.
79. Qvarnstrom, Y. et al., PCR-based detection of Angiostrongylus cantonensis in tissue and mucus secretions from molluscan hosts. Appl. Environ. Microbiol., 73, 1415, 2007.
80. Fontanilla, I.K., and Wade, C.M., The small subunit (SSU) ribosomal (r) RNA gene as a genetic marker for identifying infective 3rd juvenile stage Angiostrongylus cantonensis. Acta Trop., 105, 181, 2008.
81. He, H. et al., Preliminary molecular characterization of the human pathogen Angiostrongylus cantonensis. BMC Mol. Biol., 10, 97, 2009.
82. Helm, J. et al., A case of canine Angiostrongylus vasorum in Scotland con�rmed by PCR and sequence analysis. J. Small Anim. Pract., 50, 255, 2009.
83. Jefferies, R., Morgan, E.R., and Shaw, S.E., A SYBR green real-time PCR assay for the detection of the nematode Angiostrongylus vasorum in de�nitive and intermediate hosts. Vet. Parasitol., 166, 112, 2009. 84. Jefferies, R. et al., Improved detection of canine Angiostrongylus vasorum infection using real-time PCR and indirect ELISA. Parasitol. Res., 109, 1577, 2011. 85. Al-Sabi, M.N. et al., PCR detection of Angiostrongylus vasorum in faecal samples of dogs and foxes. Parasitol. Res., 107, 135, 2010. 86. Eamsobhana, P. et al., Molecular differentiation of Angiostrongylus taxa (Nematoda: Angiostrongylidae) by cytochrome c oxidase subunit I (COI) gene sequences. Acta Trop., 116, 152, 2010. 87. Eamsobhana, P. et al., Molecular differentiation and phylogenetic relationships of three Angiostrongylus species and Angiostrongylus cantonensis geographical isolates based on a 66-kDa protein gene of A. cantonensis (Nematoda: Angiostrongylidae). Exp. Parasitol., 126, 564, 2010. 88. Fang, W. et al., PCR assay for the cell-free copro-DNA detection of Angiostrongylus cantonensis in rat faeces. Vet. Parasitol., 183, 299, 2012. 89. Chye, S.M. et al., Immuno-PCR for detection of antigen to Angiostrongylus cantonensis circulating �fth-stage worms. Clin. Chem., 50, 51, 2004. 90. Liu, C.Y. et al., Speci�c detection of Angiostrongylus cantonensis in the snail Achatina fulica using a loop-mediated isothermal ampli�cation (LAMP) assay. Mol. Cell. Probes, 25, 164, 2011. 91. Qvarnstrom, Y. et al., Improved molecular detection of Angiostrongylus cantonensis in mollusks and other environmental samples with a species-speci�c internal transcribed spacer 1-based TaqMan assay. Appl. Environ. Microbiol., 76, 5287, 2010. 92. Gerrikagoitia, X., Barral, M., and Juste, R.A., Angiostrongylus species in wild carnivores in the Iberian Peninsula. Vet. Parasitol., 174, 175, 2010. 93. Ubelaker, J.E., Systematics of species referred to the genus Angiostrongylus. J. Parasitol., 72, 237, 1986. 94. Diao, Z., Jin, E., and Yin, C., Angiostrongylus cantonensis: Lesions in brain and spinal cord. Am. J. Trop. Med. Hyg., 82, 519, 2010. 95. Diao, Z., Yin, C., and Jin, E., Images in clinical tropical medicine. Myelitis caused by infection of Angiostrongylus cantonensis. Am. J. Trop. Med. Hyg., 83, 1176, 2010. 96. Tseng, Y.T. et al., Clinical manifestations of eosinophilic meningitis caused by Angiostrongylus cantonensis: 18 years’ experience in a medical center in southern Taiwan. J. Microbiol. Immunol. Infect., 44, 382, 2011. 97. Sawanyawisuth, K. et al., How can clinicians ensure the diagnosis of meningitic angiostrongyliasis? Vector Borne Zoonotic Dis., 12, 73, 2012. 98. Abrahams-Sandi, E. et al., An indirect immuno�uorescence antibody test employing whole eggs as the antigen for the diagnosis of abdominal angiostrongyliasis. Mem. Inst. Oswaldo Cruz, 106, 390, 2011. 99. Sawanyawisuth, K. et al., Speci�city of immunoblotting analyses in eosinophilic meningitis. Mem. Inst. Oswaldo Cruz, 106, 570, 2011. 45 Chapter 45 - Anisakis
1. Fagerholm, H.P. Systematic implications of male caudal morphology in ascaridoid nematode parasites. Syst. Parasitol. 19, 215, 1991.
2. Mattiucci, S. et al. Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology 93, 83, 1986.
3. Mattiucci, S. et al. Genetic and ecological data on the Anisakis simplex complex, with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J. Parasitol. 86, 401, 1997.
4. Nascetti, G. et al. Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North-East Atlantic. Int. J. Parasitol. 16, 633, 1986.
5. Berland, B. Nematodes from some Norwegian marine �shes. Sarsia 2, 1, 1961.
6. Mattiucci, S. et al. Allozyme and morphological identi�cation of Anisakis, Contracaecum and Pseudoterranova from Japanese waters (Nematoda, Ascaridoidea). Syst. Parasitol. 40, 81, 1998.
7. Mattiucci, S. et al. Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). Int. J. Parasitol. 31, 9, 2001.
8. Mattiucci, S. et al. Genetic markers in the study of Anisakis typica (Diesing, 1860): Larval identi�cation and genetic relationships with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae). Syst. Parasitol. 51, 159, 2002.
9. Mattiucci, S. et al. Evidence for a new species of Anisakis Dujardin, 1845: Morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Syst. Parasitol. 61, 157, 2005.
10. Mattiucci, S. et al. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: Morphological description, genetic relationships between congeners and ecological data. Syst. Parasitol. 74, 199, 2009.
11. Paggi, L. et al. A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whales (Ziphiidae): Allozyme and morphological evidence. Syst. Parasitol. 40, 161, 1998.
12. D’Amelio, S. et al. Genetic markers in ribosomal DNA for the identi�cation of members of the genus Anisakis (Nematoda: Ascaridoidea) de�ned by polymerase chain reaction based restriction fragment length polymorphism. Int. J. Parasitol. 30, 223, 2000.
13. Abollo, E. et al. Occurrence of recombinant genotypes of Anisakis simplex s.s. and Anisakis pegref�i (Nematoda: Anisakidae) in an area of sympatry. Infect. Genet. Evol. 3, 175, 2003.
14. Pontes, T. et al. Molecular characterization of larval anisakid nematodes from marine �shes of Madeira by a PCR-based approach, with evidence for a new species. J. Parasitol. 91, 1430, 2005.
15. Marques, J.F. et al. Molecular identi�cation of Anisakis species from Pleuronectiformes off the Portuguese coast. J. Helminthol. 80, 47, 2006. 16. Palm, H.W. et al. Molecular genotyping of Anisakis Dujardin, 1845 (Nematoda: Ascaridoidea: Anisakidae) larvae from marine �sh of Balinese and Javanese waters, Indonesia. Helminthologia 45, 3, 2008. 17. Davey, J.T. A revision of the genus Anisakis Dujardin, 1845 (Nematoda: Ascaridida). J. Helminthol. 45, 51, 1971. 18. Kurochkin, Y.V. Parasites of the Caspian seal Pusa caspica. Rapport et proces-verbaux des reunion. Cons. Int. Expl. Mer. 169, 363, 1975. 19. Mattiucci, S., and Nascetti, G. Molecular systematics, phylogeny and ecology of anisakid nematodes of the genus Anisakis Dujardin, 1845: An update. Parasite 13, 99, 2006. 20. Mattiucci, S., and Nascetti, G. Genetic diversity and infection levels of anisakid nematodes parasitic in �sh and marine mammals from Boreal and Austral hemispheres. Vet. Parasitol. 148, 43, 2007. 21. D’Amelio, S. et al. Molecular genotyping of anisakid nematodes from cetaceans of Florida waters. European Cetacean Society Annual Conference, Cetacean Social Organization. La Rochelle (France) 2–7 Aprile 32, 2005. 22. Umehara, A. et al. Molecular identi�cation of Anisakis simplex sensu stricto and Anisakis pegref�i (Nematoda: Anisakidae) from �sh and cetacean in Japanese waters. Parasitol. Int. 55, 267, 2006. 23. Farjallah, S. et al. Occurrence and molecular identi�cation of Anisakis spp. from the North African coasts of Mediterranean Sea. Parasitol. Res. 102, 371, 2008. 24. Iglesias, R. et al. Molecular and morphological evidence of a new taxon of Anisakis Dujardin, 1845 (Nematoda, Anisakidae) from the Blainville’s beaked whale (Mesoplodon densirostris). J. Helminthol. 82, 305, 2008. 25. Oshima, T. Anisakis-Is the sushi bar guilty? Parasitol. Today 3, 44, 1987. 26. Kasuya, S. et al. Mackerel-induced urticaria and Anisakis. Lancet 335, 665, 1990. 27. Kagei, N., and Isogaki, H. A case of abdominal syndrome caused by the presence of a large number of Anisakis larvae. Int. J. Parasitol. 22, 251, 1992. 28. Audicana, M.T. et al. Anisakis simplex: Dangerous dead and alive? Trends Parasitol. 18, 20, 2002. 29. Audicana, M.T., and Kennedy, M.W. Anisakis simplex: From obscure infectious worm to inducer of immune hypersensitivity. Clin. Microbiol. Rev. 21, 360, 2008. 30. Verhamme, M.A., and Ramboer, C.H. Anisakiasis caused by herring in vinegar: A little known medical problem. Gut. 29, 843, 1988. 31. Daschner, A. et al. Gastric anisakiasis: An underestimated cause of acute urticaria and angioedema. Br. J. Dermatol. 139, 822, 1998. 32. Fernández de Corres, L. et al. Anisakis simplex induces not only anisakiasis: Report n 28 cases of allergy caused by this nematode. J. Investig. Allergol. Clin. Immunol. 6, 315,, 1996. 33. Del Pozo, M.D. et al. Anisakis simplex, a relevant etiologic factor in acute urticaria. Allergy 52, 576, 1997. 34. Ohtaki, H., and Ohtaki, R. Clinical manifestation of gastric anisakisis. In: Ishikura, H., and Namiki, M. (eds), Gastric Anisakiasis in Japan, Springer-Verlag, Tokyo, p. 37, 1989. 35. Shibata, O., Uchida, Y., and Furusawa, T. Acute gastric anisakiasis with special analysis of the location of the worms penetrating the gastric mucosa. In: Ishikura, H., and Namiki, M. (eds) Gastric Anisakiasis in Japan Springer-Verlag, Tokyo, p. 53, 1989.
36. Ishikura, H., Kikuchi, Y., and Hayasaka, H. Pathological and clinical observation on intestinal anisakiasis. Arch. Jpn. Chir. 36, 663, 1967.
37. Rosales, M. et al. Acute intestinal anisakiasis in Spain: A fourth stage Anisakis simplex larva. Mem. Inst. Oswaldo Cruz 94, 823, 1999.
38. Moschella, C.M. et al. Intestinal anisakiasis in Italy: A case treated by emergency surgery. G. Chir. 26, 201, 2005.
39. Ishikura, H., and Kikuchi K. Intestinal Anisakiasis in Japan, Springer-Verlag, Tokyo, 1990. 40. Rodriguéz, E. et al. Novel sequences and epitopes of diagnostic value derived from the Anisakis simplex Ani s 7 major allergen. Allergy 63, 219, 2008.
41. Anibarro, B., and Seoane, F.J. Occupational conjunctivitis caused by sensitizazion to Anisakis simplex. J. All. Clin. Immunol. 102, 331, 1998.
42. Armentia, A. et al. Anisakis simplex allergy after eating chicken meat. J. Investig. Allergol. Clin. Immunol. 16, 258, 2006.
43. Kikuchi, Y., Ishikura, H., and Kikuchi, K. Pathology of intestinal anisakiasis. Ishikura, H., and Kikuchi K. (eds), In Intestinal Anisakiasis in Japan. Springer-Verlag, Tokyo, p. 129, 1990.
44. Daschner, A. Usefulness of early serial measurement of speci�c and total IgE in the diagnostico of gastro-allergic anisakiasis. Clin. Exp. All. 29.1260, 1999.
45. Daschner, A. What does Anisakis simplex parasitism teach us about interpretating speci�c and total IgE values? Allergol. Immunopathol. 28, 67, 2000.
46. Moneo, I. et al. Isolation and characterization of a major allergen from the �sh parasite Anisakis simplex. J. All. Clin. Immunol. 106, 177, 2000.
47. Shikamura, K. et al. Puri�cation and molecular cloning of a major allergen from Anisakis simplex. Mol. Biochem. Parasitol. 135, 69, 2004.
48. Moneo, I. et al. Sensitization to the �sh parasite Anisakis simplex: Clinical and laboratory aspects. Parasitol. Res. 101, 1051, 2007.
49. Moneo, I. et al. Isolation of a heat-resistant allergen from the �sh parasite Anisakis simplex. Parasitol. Res. 96, 285, 2005.
50. Anádon, A.M. et al. The Anisakis simplex Ani s 7 major allergen as an indicator of true Anisakis infections. Clin. Exp. Immunol. 156, 471, 2009.
51. Paggi, L. et al. Biochemical taxonomy of ascaridoid nematodes. Parassitologia 27, 105, 1985.
52. Paggi, L. et al. Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the North Atlantic and Norwegian and Barents Seas. Int. J. Parasitol. 21, 195, 1991.
53. Paggi, L. et al. Nematodi del genere Anisakis in pesci, cefalopodi e cetacei del Mar Mediterraneo e dell’Oceano Atlantico e Paci�co. Biol. Mar. Medit. 5, 1585, 1998.
54. Paggi, L. et al. Pseudoterranova decipiens species A and B (Nematoda, Ascaridoidea): Nomenclatural designation, morphological diagnostic characters and genetic markers. Syst. Parasitol. 45, 185, 2000.
55. Bullini, L. et al. Genetic variation of ascaridoid worms with different life cycles. Evolution 40, 437, 1986. 46 Chapter 46 - Ascaris
1. Tenora, F., and Borkovcová, M. Corrections in the review of helminthofauna of Ondatra zibethica (L., 1785) (Rodentia) in the Czech Republic. Acta Univ. Agric. Silvic. Mendelianae Brunensis 51, 65, 2003.
2. Maung, M. An ascaridoid nematode from a hippopotamus. Int. J. Parasitol. 5, 431, 1975.
3. Sprent, J.F.A. Ascaridoid nematodes of South American mammals, with a de�nition of a new genus. J. Helminthol. 56, 275, 1982. 4. Sprent, J.F.A. Notes on Ascaris and Toxascaris, with a de�nition of Baylisascaris gen. nov. Parasitology 58, 185, 1968. 5. Warren, G. Studies on the morphology and taxonomy of the genera Toxocara stiles, 1905 and Neoascaris travassos, 1927. Zool. Anz. 185, 393, 1970. 6. Tenora, F., and Shalayeva, N. Scanning electron microscopy of Ascaris tarbagan (Nematoda). Acta Soc. Zool. Bohemoslov. 54, 130, 1990. 7. Anderson, R.C. Nematode Parasites of Vertebrates, Their Development and Transmission, 2nd edn. CABI, Wallingford, UK, 650pp., 2000. 8. Meldal, B.H. et al. An improved molecular phylogeny of the Nematoda with special emphasis on marine taxa. Mol. Phylogen. Evol. 42, 622, 2007. 9. Sprent, J.F.A. Anatomical distinction between human and pig strains of Ascaris. Nature 170, 627, 1952. 10. Abdulrachman, S., and Joe, L.K. Morphological differences between Ascaris from man and pigs. Doc. Med. Geo. Tropica 6, 342, 1954. 11. Ubelaker, J.E., and Allison, V.F. Scanning electron microscopy of the denticles and eggs of Ascaris lumbricoides and Ascaris suum. Thirtieth Annual Proceedings Electron Microscopy Society America, 1972. 12. Maung, M. Ascaris lumbricoides Linne, 1758 and Ascaris suum Goeze, 1782: Morphological differences between specimens obtained from man and pig. Southeast Asian J. Trop. Med. Public Health 4, 41, 1973. 13. Weise, R.W. Ascaris suum: A scanning electron microscope study. J. Parasitol. 59, 141, 1973. 14. Kurimoto, H. Morphological, biochemical and immunological studies on the differences between Ascaris lumbricoides Linnaeus, 1758 and Ascaris suum Goeze, 1782. Jpn. J. Parasitol. 23, 251, 1974. 15. Little, M.D. Comparison of human and pig Ascaris. Eighth International Congress Tropical Medicine Malaria. Abstracts & Reviews, pp. 181, 1968. 16. Yi, K. et al. Dephosphorylation of major sperm protein (MSP) �ber protein 3 by protein phosphatase 2A during cell body retraction in the MSP-based amoeboid motility of Ascaris sperm. Mol. Biol. Cell 20, 3200, 2009. 17. Beaver, P.C., and Little, M.D. The genital girdle in relation to estrus and mating in Ascaris lumbricoides. J. Parasitol. 50, 128, 1964. 18. Brown, H.W., and Cort, W.W. The egg production of Ascaris lumbricoides. J. Parasitol. 14, 88, 1927. 19. Mello, D.A. A note on egg production of Ascaris lumbricoides. J. Parasitol. 60, 380, 1974. 20. Roepstorff, A. The earthworm Lumbricus terrestris as a possible paratenic or intermediate host of the pig parasite Ascaris suum. Comp. Parasitol. 69, 206, 2002. 21. Murrell, K.D. et al. Ascaris suum: A revision of its early migratory path and implications for human ascariasis. J. Parasitol. 83, 255, 1997. 22. Seo, B.S. Ascariasis and its control problems in Korea. Seoul J. Med. 22, 323, 1981. 23. Hobo, B. Epidemiological studies on Ascaris infection among prisoners and the length of life of Ascaris lumbricoides in human host. Jpn. J. Natl Health 25, 1, 1956. 24. Galvin, T.J. Development of human and pig ascaris in the pig and rabbit. J. Parasitol. 54, 1085, 1968. 25. Beaver, P.C., Jung, R.C., and Cupp, E.W. Clinical Parasitology. Lea and Febiger, Philadelphia, 825pp., 1986.
26. Orihel, T.C. Primates as models for parasitological research. In: Goldsmith, E.I. and Moor-Jankowski, J. (eds), Medical Primatology, S. Karger, Basel, 772pp., 1971.
27. Kmetec, E., Beaver, P.C., and Bueding, E. Succinoxidase activities and cytochrome oxidase of extracts of Ascaris eggs. Comp. Biochem. Physiol. 9, 115, 1963.
28. Brudastov, A.N. et al. Infectivity of Ascaris lumbricoides eggs to man and guinea-pigs after 10 years in the soil. Med. Parasitol. Parasit. Dis. 39, 447, 1970.
29. Krasnonos, L.L. Many-year viability of ascarid eggs (Ascaris lumbricoides L. 1758) in soil of Sarkand. Med. Parasitol. Parasit. Dis. 47, 103, 1978.
30. Crompton, D.W. Ascaris and ascariasis. Adv. Parasitol. 48, 285, 2001.
31. Stephenson, L.S. Pathophysiology of intestinal nematodes, In Holland, C.V. and Kennedy, M.W. (eds), World Class Parasites: Volume 2. The Geohelminths: Ascaris, Trichuris, and Hookworm, Springer, Heidelberg, Germany, pp. 39–61, 2002.
32. Hadju, V. et al. Improvements of growth, appetite, and physical activity in helminth-infected schoolboys 6 months after single dose of albendazole. Asia Pac. J. Clin. Nutr. 7, 170, 1998.
33. Hadju, V. et al. Improvements in appetite and growth in helminth-infected schoolboys three and seven weeks after a single dose of pyrantel pamoate. Parasitology 113, 497, 1996.
34. Tripathy, K. et al. Malabsorption syndrome in ascariasis. Am. J. Clin. Nutr. 25, 1276, 1972.
35. McOrist, S., Blunt, R., and Wiseman, J. Ascaris suum parasitism in �nisher pigs: Welfare and economic impacts and control options. Pig J. 61, 74, 2008.
36. Crofton, H.D. A quantitative approach to parasitism. Parasitology 62, 179, 1971.
37. Holland, C., and Boes, J. Distribution and predisposition: People and pigs, in Holland, C.V. and Kennedy, M.W. (eds), World Class Parasites: Volume 2. The Geohelminths: Ascaris, Trichuris, and Hookworm, Springer, Heidelberg, Germany, pp. 1–24, 2002.
38. Elkins, D.B., Haswell-Elkins, M., and Anderson, R.M. The epidemiology and control of intestinal nematodes in the Pulicat Lake region of Southern India. I. Study design and pre- and post-treatment observations of Ascaris lumbricoides infection. Trans. R. Soc. Trop. Med. Hyg. 80, 774, 1986.
39. Boes, J. et al. Distribution of Ascaris suum in experimentally and naturally infected pigs and comparison with Ascaris lumbricoides infections in humans. Parasitology 117, 589, 1998.
40. Little, M.D. Nematodes of the digestive tract and related species. In: Beaver, P.C. and Jung, R.C. (eds), Animal Agents and Vectors of Human Disease, Lea & Febiger, Philadelphia, pp. 127–170, 1985.
41. Crompton, D.W.T. How much human helminthiasis is there in the world? J. Parasitol. 85, 397, 1999.
42. Kurosawa, C.M. et al. The half-life of Ascaris lumbricoides prevalence in Japanese school children. Acta Med. Okayama 62, 303, 2008.
43. Chai, J.Y. et al. Reduction in the incidence of biliary and other surgical complications of ascariasis according to the decrease of its national egg prevalence in Korea. Korean J. Parasitol. 29, 101, 1991.
44. Garcia, L,S. Diagnostic Medical Parasitology, 5th edn. ASM Press, Washington, DC, pp. 1092, 2006.
45. WHO. Bench Aids for the Diagnosis of Intestinal Parasites. Prepared by LR Ash, TC Orihel, L Savioli. World Health Organization, Geneva. 46. Komiya, Y., and Kobayashi, A. Evaluation of Kato’s thick smear technic with a cellophane cover for helminth eggs in feces. Jpn. J. Med. Sci. Biol. 19, 59, 1966. 47. Martin, L.K., and Beaver, P.C. Evaluation of Kato thick-smear technique for quantitative diagnosis of helminth infections. Am. J. Trop. Med. Hyg. 17, 382, 1968. 48. Katz, N., Coelho, P.M.Z., and Pellegrino, J. Evaluation of Kato’s quantitative method through the recovery of Schistosoma mansoni eggs added to human feces. J. Parasitol. 56, 1032, 1970. 49. Leles, D. et al. Molecular diagnosis of ascariasis from human feces and description of a new Ascaris sp. genotype in Brazil. Vet. Parasitol. 163, 167, 2009. 50. Anderson, T.J.C., Romero-Abal, M.E., and Jaenike, J. Genetic structure and epidemiology of Ascaris populations: Patterns of host af�liation in Guatemala. Parasitology 107, 319, 1993. 51. Anderson, T.J.C., Romero-Abal, M., and Jaenike, J. Mitochondrial DNA and Ascaris microepidemiology: The composition of parasite populations from individual hosts, families and villages. Parasitology 110, 221, 1995. 52. Anderson, T.J.C., and Jaenike, J. Host speci�city, evolutionary relationships and macrogeographic differentiation among Ascaris populations from humans and pigs. Parasitology 115, 325, 1997. 53. Anderson, T.J.C. The dangers of using single locus markers in parasite epidemiology: Ascaris as a case study. Trends Parasitol. 17, 183, 2001. 54. Criscione, C.D. et al. Microsatellite markers for the human nematode parasite Ascaris lumbricoides: Development and assessment of utility. J. Parasitol. 93, 704, 2007. 55. Criscione, C.D. et al. Disentangling hybridization and host colonization in parasitic roundworms of humans and pigs. Proc. R. Soc. Lond. B. 274, 2669, 2007. 56. Nadler, S.A., Lindquist, R.L., and Near, T.J. Genetic structure of midwestern Ascaris suum populations: A comparison of isoenzyme and RAPD markers. J. Parasitol. 81, 385, 1995. 57. Nejsum, P. et al. Population structure in Ascaris suum (Nematoda) among domestic swine in Denmark as measured by whole genome DNA �ngerprinting. Hereditas 142, 7, 2005. 58. Peng, W. et al. Genetic variation in sympatric Ascaris populations from humans and pigs in China. Parasitology 117, 355, 1998. 59. Peng, W. et al. Molecular epidemiological investigation of Ascaris genotypes in China based on single-strand conformation polymorphism analysis of ribosomal DNA. Electrophoresis 24, 2308, 2003. 60. Peng, W. et al. Mutation scanning-coupled analysis of haplotypic variability in mitochondrial DNA regions reveals low gene �ow between human and porcine Ascaris in endemic regions of China. Electrophoresis 26, 4317, 2005. 61. Peng, W. et al. Experimental infections of pigs and mice with selected genotypes of Ascaris. Parasitology 133, 651, 2006. 62. Peng, W. et al. Recent insights into the epidemiology and genetics of Ascaris in China using molecular tools. Parasitology 134, 325, 2007. 63. Zhu, X. et al. Characterisation of Ascaris from human and pig hosts by nuclear ribosomal DNA sequences. Int. J. Parasitol. 29, 469, 1999. 64. Anderson T.J.C. Ascaris infections in humans from North America: Molecular evidence for cross-infection. Parasitology 110, 215, 1995. 65. Nejsum, P. et al. Ascariasis is a zoonosis in Denmark. J. Clin. Microbiol. 43, 1142, 2005.
66. Nejsum, P. et al. A novel technique for identi�cation of Ascaris suum cohorts in pigs. Vet. Parasitol. 154, 171, 2008.
67. Anderson, T.J.C. et al. Are mitochondria inherited paternally in Ascaris? Int. J. Parasitol. 25, 1001, 1995.
68. Pecson, B.M. et al. A real-time PCR method for quantifying viable Ascaris eggs using the �rst internally transcribed spacer region of ribosomal DNA. Appl. Environ. Microbiol. 72, 7864, 2006.
69. Gonçalves, M.L.C., Araujo, A., and Ferreira, L.F. Human intestinal parasites in the past: New �ndings and a review. Mem. Inst. Oswaldo Cruz 98, 103, 2003. 70. Loreille, O. et al. Ancient DNA from Ascaris: Extraction ampli�cation and sequences from eggs collected in coprolites. Int. J. Parasitol. 31, 1101, 2001. 71. Leles, D. et al. Molecular paleoparasitological diagnosis of Ascaris sp. from coprolites: New scenery of ascariasis in preColombian South America times. Mem. Inst. Oswaldo Cruz 103, 106, 2008. 72. Leles, D. et al. ITS1 intra-individual variability of Ascaris isolates from Brazil. Parasitol. Int. 59, 93, 2010. 73. Hall, A., and Holland, C. Geographical variation in Ascaris lumbricoides fecundity and its implications for helminth control. Parasitol. Today 16, 540, 2000. 47 Chapter 47 - Baylisascaris
21. Page, L.K., Swihart, R.K., and Kazacos, K.R., Implications of raccoon latrines in the epizootiology of baylisascariasis. J. Wildl. Dis., 35, 474, 1999.
22. Sorvillo, F. et al., Baylisascaris procyonis: An emerging helminthic zoonosis. Emerg. Infect. Dis., 8, 355, 2002.
23. Weller, P.F., Eosinophilic meningitis. Am. J. Med., 95, 250, 1993.
24. Moertel, C.L. et al., Eosinophil-associated in�ammation and elaboration of eosinophil-derived proteins in two children with raccoon roundworm (Baylisascaris procyonis) encephalitis. Pediatrics, 108, e93, 2001.
25. Cunningham, C.K. et al., Diagnosis and management of Baylisascaris procyonis infection in an infant with nonfatal meningoencephalitis. Clin. Infect. Dis., 18, 868, 1994.
26. Boyce, W.M., Branstetter, B.A., and Kazacos, K.R., In vitro culture of Baylisascaris procyonis and initial analysis of larval excretory-secretory antigens. Proc. Helminthol. Soc. Washington, 55, 15, 1988.
27. Boyce, W.M. et al., Physiochemical characterization and monoclonal and polyclonal antibody recognition of Baylisascaris procyonis larval excretory-secretory antigens. J. Parasitol., 75, 540, 1989.
28. Hamann, K.J. et al., Immuno�uorescent localization of eosinophil granule major basic protein in fatal human cases of Baylisascaris procyonis infection. Am. J. Trop. Med. Hyg., 40, 291, 1989.
29. Dangoudoubiyam, S. et al., Molecular cloning of an immunogenic protein of Baylisascaris procyonis and expression in Escherichia coli for use in developing improved serodiagnostic assays. Clin. Vaccine Immunol., 17, 1933, 2010.
30. Dangoudoubiyam, S. et al., Recombinant antigen-based enzyme-linked immunosorbent assay for diagnosis of Baylisascaris procyonis larva migrans. Clin. Vaccine Immunol., 18, 1650, 2011. 31. Pontes, T. et al., Molecular characterization of larval anisakid nematodes from marine �shes of Madeira by a PCR-based approach, with evidence for a new species. J. Parasitol., 91, 1430, 2005. 32. Abe, N., Tominaga, K., and Kimata, I., Usefulness of PCRrestriction fragment length polymorphism analysis of the internal transcribed spacer region of rDNA for identi�cation of Anisakis simplex complex. Jpn. J. Infect. Dis., 59, 60, 2006. 33. Chen, Q. et al., Speci�c PCR assays for the identi�cation of common anisakid nematodes with zoonotic potential. Parasitol. Res., 104, 79, 2008. 34. Umehara, A. et al., Multiplex PCR for the identi�cation of Anisakis simplex sensu stricto, Anisakis pegref�i and the other anisakid nematodes. Parasitol. Int., 57, 49, 2008. 35. Dangoudoubiyam, S., Vemulapalli, R., and Kazacos, K.R., PCR assays for detection of Baylisascaris procyonis eggs and larvae. J. Parasitol., 95, 571, 2009. 36. Fang, W. et al., Multiple primer PCR for the identi�cation of anisakid nematodes from Taiwan Strait. Exp. Parasitol., 124, 197, 2010. 37. Leles, D. et al., ITS1 intra-individual variability of Ascaris isolates from Brazil. Parasitol. Int., 59, 93, 2010. 38. Testini, G. et al., New insights into the morphology, molecular characterization and identi�cation of Baylisascaris transfuga (Ascaridida, Ascarididae). Vet. Parasitol., 175, 97, 2011. 39. Gatcombe, R.R. et al., Evaluation of a molecular beacon realtime PCR assay for detection of Baylisascaris procyonis in different soil types and water samples. Parasitol. Res., 106, 499, 2010. 40. Walker, M.D., and Zunt, J.R., Neuroparasitic infections: Nematodes. Semin. Neurol., 25, 252, 2005. 41. Page, K. et al., Reducing Baylisascaris procyonis roundworm larvae in raccoon latrines. Emerg. Infect. Dis., 17, 90, 2011. 48 Chapter 48 - Brugia
1. WHO Lymphatic �lariasis. Fourth report of the WHO Expert Committee on Filariasis. World Health Organ. Tech. Rep. Ser. 702, 3–112, 1984.
2. Baird, J.K. et al. North American brugian �lariasis: Report of nine infections of humans. Am. J. Trop. Med. Hyg. 35, 1205, 1986.
3. Cox, F.E. Elimination of lymphatic �lariasis as a public health problem. Parasitol. Today 16, 135, 2000.
4. Despommier, D. et al. Lymphatic �lariae: Wuchereria bancrofti, Cobbold 1877) and Brugia malayi (Brug 1927). In: Despommier, D.D. and Gwadz, R.W. (eds.), Parasitic Diseases, 3 edn. Springer-Verlag, New York, pp. 40–47, 1995. 5. Markell, E. et al. The blood and tissue dwelling nematodes. In: Markell, E.K., John, D.T., and Williams, A. (eds.), Medical Parasitology.W.B. Saunders Company, London, p. 304, 1999. 6. Arcari, M., Baxendine, A., and Bennett, C.E. The blood nematodes. http://www.southampton.ac.uk/~ceb/Diagnosis/ Vol10.htm 7. Chansiri, K. et al. PCR based method for identi�cation of zoonostic Brugia malayi micro�lariae in domestic cats. Mol. Cell. Probes 16, 129, 2002. 8. Sucharit, S. et al. The differentiation of brugia malayi, B. pahangi, B. tupaiae and Wuchereria bancrofti. Southeast Asian J. Trop. Med. Public Health 6, 549, 1975. 9. Saitongdee, P. Scanning electron microscopic study of micro�laria, third stage larva and adult of Brugia malayi, B. pahangi and third stage larva of Wuchereria bancrofti. In: Anatomy, Mahidol University, pp. 148, 1989. 10. Sivandam, S., and Fredericks, H. The Innenkorper in differentation between the micro�lariae of Brugia pahangi and B. malayi (superiodic form). Med. J. Malaya 20, 337, 1966. 11. Buckley, J.J., and Edeson J.F. On the adult morphology of Wuchereria sp. (malayi?) from a monkey (Macaca irus) and from cats in Malaya, and on Wuchereria pahangi n. sp. from a dog and a cat. J. Helminthol. 30, 1, 1956. 12. Suguri, S. Ultrastructure of the micro�laria of Brugia pahangi (Buckley and Edeson, 1956) Buckley, 1958. Acta Med. Okayama 31, 295, 1977. 13. Lim P. et al. Scanning electron microscope study of subperiodic Brugia malayi and Brugia pahangi infective larvae (Filarioidae: Onchocercidae). Trop. Bio Med. 1, 29, 1984. 14. Aoki Y. et al., Scanning electron microscopy of third- and fourth-stage larvae and adults of Brugia pahangi (Nematoda: Filarioidea). J. Parasitol. 66, 449, 1980. 15. Leeuwin, R.S. Micro�laraemia in Surinamese living in Amsterdam. Trop. Geogr. Med. 14, 355, 1962. 16. Esslinger, J.H. Development of Porocephalus crotali (Humboldt, 1808) (Pentastomida) in experimental intermediate hosts. J. Parasitol. 48, 452, 1962. 17. The World Health Report 1995—Bridging the gaps. World Health Forum 16, 377, 1995. 18. Schacher, J.F. Morphology of the micro�laria of Brugia pahangi and of the larval stages in the mosquito. J. Parasitol. 48, 679, 1962. 19. Smyth, J. Introduction to Animal Parasitology: Nematoda: General Account, 3rd edn. Cambridge University Press, Cambridge, p. 369, 1994. 20. Life cycle of B. malayi. In: from URL:http://www.dpd.cdc. gov/ dpdx/HTML/ImageLibrary/Filariasis_il.asp?body = AF/ Filariasis/body_Filariasis_il17.htm. Filariasis, ed. Image Library, 2005. 21. Jordan, P. Notes on elephantiasis and hydrocoele due to Wuchereria bancrofti. J. Trop. Med. Hyg. 58, 113, 1955. 22. Chandrashekar, R. et al. Molecular cloning of Brugia malayi antigens for diagnosis of lymphatic �lariasis. Mol. Biochem. Parasitol. 64, 261, 1994. 23. Dissanayake, S. et al. Evaluation of a recombinant parasite antigen for the diagnosis of lymphatic �lariasis. Am. J. Trop. Med. Hyg. 50, 727, 1994. 24. Fischer, P. et al. PCR-based detection and identi�cation of the �larial parasite Brugia timori from Alor Island, Indonesia. Ann. Trop. Med. Parasitol. 96, 809, 2002. 25. Nuchprayoon, S. et al. Detection and differentiation of �larial parasites by universal primers and polymerase chain reactionrestriction fragment length polymorphism analysis. Am. J. Trop. Med. Hyg. 73, 895, 2005.
26. Rishniw, M. et al. Discrimination between six species of canine micro�lariae by a single polymerase chain reaction. Vet. Parasitol. 135, 303, 2006.
27. Thanomsub, B. et al. Differential diagnosis of human lymphatic �lariasis using PCR-RFLP. Mol. Cell. Probes 14, 41, 2000.
28. Areekit, S. et al. Intraspecies variation of Brugia spp. in cat reservoirs using complete ITS sequences. Parasitol. Res. 104, 1465, 2009.
29. Areekit, S. et al. Molecular genetics analysis for co- infection of Brugia malayi and Brugia pahangi in cat reservoirs based on internal transcribed spacer region 1. Southeast Asian J. Trop. Med. Public Health 40, 30, 2009.
30. Wilhelm, J., and Pingoud, A. Real-time polymerase chain reaction. Chembiochem 4, 1120, 2003. 31. Ririe K.M. et al. Product differentiation by analysis of DNA melting curves during the polymerase chain reaction. Anal. Biochem. 245, 154, 1997. 32. Areekit, S. et al. High resolution melting real-time PCR for rapid discrimination between Brugia malayi and Brugia pahangi. J. Med. Assoc. Thai. 92, 24, 2009. 33. Swofford, D.L. PAUP (version 4.0b10). Sinauer Associates, Sunderland, MA, 2002. 34. Nuchprayoon, S. et al. Detection of �larial parasites in domestic cats by PCR-RFLP of ITS1. Vet. Parasitol. 140, 366, 2006. 35. Yao, C. et al. Hybridization assay of hepatitis B virus by QCM peptide nucleic acid biosensor. Biosens. Bioelectron. 23, 879, 2008 36. Zhou, X. et al. Microgravimetric DNA sensor based on quartz crystal microbalance: Comparison of oligonucleotide immobilization methods and the application in genetic diagnosis. Biosens. Bioelectron. 16, 85, 2001. 49 Chapter 49 - Capillaria
1. Butterworth, E.W., and Beverley-Burton, M., The taxonomy of Capillaria spp. (Nematoda: Trichuroidea) in carnivorous mammals from Ontario, Canada. Syst. Parasitol., 1, 211, 1980.
2. Moravec, F., Proposal of a new systematic arrangement of nematodes of the family Capillariidae. Folia Parasitol. (Praha), 29, 119, 1982.
3. Moravec, F., Nagasawa, K., and Madinabeitia, I., A new species of Capillaria (Nematoda: Capillariidae) from the intestine of the marine �sh Acanthopagrus schlegelii schlegelii (Sparidae) from Japan. J. Parasitol., 96, 771, 2010.
4. Moravec, F., Redescription and systematic status of Capillaria philippinensis, an intestinal parasite of human beings. J. Parasitol., 87, 161, 2001.
5. Traversa, D. et al., New insights into morphological and biological features of Capillaria aerophila (Trichocephalida, Trichuridae). Parasitol. Res., 109, S97, 2011.
6. Camargo, L.M. et al., Capillariaisis (Trichurida, Trichinellidae, Capillaria hepatica) in the Brazilian Amazon: Low pathogenicity, low infectivity and a novel mode of transmission. Parasit. Vectors, 3, 11, 2010.
7. Jeong, W.I. et al., Macrophages, myo�broblasts and mast cells in a rat liver infected with Capillaria hepatica. J. Vet. Sci., 9, 211, 2008.
8. Fan, P.C., Chung, W.C., and Chen, E.R., Capillaria hepatica: A spurious case with a brief review. Kaohsiung J. Med. Sci., 16, 360, 2000.
9. Gomes, A.T. et al., Capillaria hepatica in rats: Focal parasitic hepatic lesions and septal �brosis run independent courses. Mem. Inst. Oswaldo Cruz, 101, 895, 2006.
10. Hamir, A.N., and Rupprecht, C.E., Hepatic capillariasis (Capillaria hepatica) in porcupines (Erethizon dorsatum) in Pennsylvania. J. Vet. Diagn. Invest., 12, 463, 2000.
11. Oliveira, L., de Souza, M.M., and Andrade, Z.A., Capillaria hepatica-induced hepatic �brosis in rats: Paradoxical effect of repeated infections. Rev. Soc. Bras. Med. Trop., 37, 123, 2004.
12. Redrobe, S.P., and Patterson-Kane, J.C., Calodium hepaticum (syn. Capillaria hepatica) in captive rodents in a zoological garden. J. Comp. Pathol., 133, 73, 2005.
13. Reperant, L.A., and Deplazes, P., Cluster of Capillaria hepatica infections in non-commensal rodents from the canton of Geneva, Switzerland. Parasitol. Res., 96, 340, 2005.
14. Santos, C.C., Onofre-Nunes, Z., and Andrade, Z.A., Role of partial hepatectomy on Capillaria hepatica-induced hepatic �brosis in rats. Rev. Soc. Bras. Med. Trop., 40, 495, 2007. 15. Pizzi, R. et al., Capillaria hepatica (syn. Calodium hepaticum) in primates in a zoological collection in the UK. Vet. Rec., 163, 690, 2008. 16. Mowat, V. et al. Histopathological features of Capillaria hepatica infection in laboratory rabbits. Toxicol. Pathol., 37, 661, 2009. 17. Quadros, R.M. et al., Capillaria hepatica in Puma concolor: First report in Brazil. J. Zoo Wildl. Med., 40, 586, 2009. 18. Resendes, A.R. et al., Prevalence of Calodium hepaticum (syn. Capillaria hepatica) in house mice (Mus musculus) in the Azores archipelago. Vet. Parasitol., 160, 340, 2009. 19. Stidworthy, M.F. et al., Capillaria hepatica in primates in zoological collections in the British Isles. Vet. Rec., 164, 66, 2009. 20. Sekikawa, H. et al., First human case of Capillaria hepatica infection in Japan. Jpn. J. Parasitol., 40, 528, 1991. 21. Choe, G. et al., Hepatic capillariasis: First case report in the Republic of Korea. Am. J. Trop. Med. Hyg., 48, 610, 1993. 22. Bhattacharya, D. et al., Capillaria hepatica, a parasite of zoonotic importance–a brief overview. J. Commun. Dis., 31, 267, 1999. 23. Tesana, S., Puapairoj, A., and Saeseow, O., Granulomatous, hepatolithiasis and hepatomegaly caused by Capillaria hepatica infection: First case report of Thailand. Southeast Asian J. Trop. Med. Public Health, 38, 636, 2007. 24. Li, C.D., Yang, H.L., and Wang, Y., Capillaria hepatica in China. World J. Gastroenterol., 16, 698, 2010. 25. Carvalho-Costa, F.A. et al., Pseudoparasitism by Calodium hepaticum (syn. Capillaria hepatica; Hepaticola hepatica) in the Negro River, Brazilian Amazon. Trans. R. Soc. Trop. Med. Hyg., 103, 1071, 2009. 26. Mansour, N.S., Anis, M.H., and Mikhail, E.M., Human intestinal capillariasis in Egypt. Trans. R. Soc. Trop. Med. Hyg., 84, 114, 1990. 27. Chichino, G., Bernuzzi, A.M., and Bruno, A., Intestinal capillariasis (Capillaria philippinensis) acquired in Indonesia: A case report. Am. J. Trop. Med. Hyg., 47, 10, 1992. 28. Dronda, F. et al., Human intestinal capillariasis in an area of nonendemicity: Case report and review. Clin. Infect. Dis., 17, 909, 1993. 29. Kang, G. et al., Human intestinal capillariasis: First report from India. Trans. R. Soc. Trop. Med. Hyg., 88, 204, 1994. 30. Hoghooghi-Rad, N., Maraghi, S., and Narenj-Zadeh, A., Capillaria philippinensis infection in Khoozestan Province, Iran: Case report. Am. J. Trop. Med. Hyg., 37, 135, 1987. 31. Ahmed, L. et al., Capillaria philippinensis: An emerging parasite causing severe diarrhoea in Egypt. J. Egypt. Soc. Parasitol., 29, 483, 1999. 32. Austin, D.N. et al., Intestinal capillariasis acquired in Egypt. Eur. J. Gastroenterol. Hepatol., 11, 935, 1999. 33. Belizario, V.Y. Jr. et al., Intestinal capillariasis, western Mindanao, the Philippines. Emerg. Infect. Dis., 16, 736, 2010. 34. Wongsawasdi, L., Ukarapol, N., and Lertprasertsuk, N., The endoscopic diagnosis of intestinal capillariasis in a child: A case report. Southeast Asian J. Trop. Med. Public Health, 33, 730, 2002. 35. Lu, L.H. et al., Human intestinal capillariasis (Capillaria philippinensis) in Taiwan. Am. J. Trop. Med. Hyg., 74, 810, 2006. 36. Soukhathammavong, P. et al., Three cases of intestinal capillariasis in Lao People’s Democratic Republic. Am. J. Trop. Med. Hyg., 79, 735, 2008. 37. Lalosević, D. et al., Pulmonary capillariasis miming bronchial carcinoma. Am. J. Trop. Med. Hyg., 78, 14, 2008.
38. Berger, T. et al., Hepatic capillariasis in a 1-year-old child. Eur. J. Pediatr., 149, 333, 1990.
39. Kokai, G.K. et al., Capillaria hepatica infestation in a 2-year old girl. Histopathology, 17, 275, 1990.
40. Sawamura, R. et al., Hepatic capillariasis in children: Report of 3 cases in Brazil. Am. J. Trop. Med. Hyg., 61, 642, 1999.
41. Gonzalez-Barranco, D. et al., Third case of human hepatic capillariasis in Mexico. Rev. Investig. Clin., 48, 301, 1996.
42. Govil, H., and Desai, M., Capillaria hepatica parasitism. Indian J. Pediatr., 63, 698, 1996.
43. Pampiglione, S., and Gustinelli, A., Human hepatic capillariasis: A second case occurred in Korea. J. Korean Med. Sci., 23, 560, 2008.
44. Nabi, F. et al., Capillaria hepatica infestation. Indian Pediatr., 44, 781, 2007.
45. Fuehrer, H.P., Igel, P., and Auer, H., Capillaria hepatica in man-an overview of hepatic capillariosis and spurious infections. Parasitol. Res., 109, 969, 2011.
46. Bustamente-Sarabia, J., Suswillo, R., and Contreras-Moreno, E., Capillaria hepatica in a 19-month old boy, third case in Mexico. Bol. Med. Hosp. Infant Mex., 53, 395, 1996.
47. Ferreira, L.A., and Andrade, Z.A., Capillaria hepatica: A cause of septal �brosis of the liver. Mem. Inst. Oswaldo Cruz, 88, 441, 1993.
48. Lemos, Q.T., Magalhães-Santos, I.F., and Andrade, Z.A., Immunological basis of septal �brosis of the liver in Capillaria hepatica-infected rats. Braz. J. Med. Biol. Res., 36, 1201, 2003.
49. Andrade, S.B., and Andrade, Z.A., Experimental hepatic �brosis due to Capillaria hepatica infection (differential features presented by rats and mice). Mem. Inst. Oswaldo Cruz, 99, 399, 2004.
50. Kohatsu, H. et al., A space-occupying lesion in the liver due to Capillaria infection. Am. J. Trop. Med. Hyg., 52, 414, 1995.
51. Attah, E.B. et al., Hepatic capillariasis. Am. J. Clin. Pathol., 79, 127, 1983.
52. Chunlertrith, K., Mairiang, P., and Sukeepaisarnjaroen, W., Intestinal capillariasis: A cause of chronic diarrhea and hypoalbuminemia. Southeast Asian J. Trop. Med. Public Health, 23, 433, 1992.
53. Lee, S.H. et al., A case of intestinal capillariasis in the Republic of Korea. Am. J. Trop. Med. Hyg., 48, 542, 1993.
54. Hong, S.T. et al., Two cases of intestinal capillariasis in Korea. Korean J. Parasitol., 32, 43, 1994.
55. Wilairatana, P., Capillariasis with chronic intestinal pseudoobstruction. Southeast Asian J. Trop. Med. Public Health, 25, 600, 1994. 56. Sangchan, A. et al., The endoscopic-pathologic �ndings in intestinal capillariais: A case report. J. Med. Assoc. Thai., 90, 175, 2007. 57. Kusolsuk, T. et al., Capillariasis: Chronic watery diarrhea–not only from microorganisms. Southeast Asian J. Trop. Med. Public Health, 39, 1000, 2008. 58. Cross, J.H., Intestinal capillariasis. Parasitol. Today, 6, 26, 1990. 59. Cross, J.H., and Basaca-Sevilla, V., Capillariasis philippinensis: A �sh-borne parasitic zoonosis. Southeast Asian J. Trop. Med. Public Health, 22, S153, 1991. 60. Cross, J.H., Intestinal capillariasis. Clin. Microbiol. Rev., 5, 120, 1992. 61. Bair, M.J. et al., Clinical features of human intestinal capillariasis in Taiwan. World J. Gastroenterol., 10, 2391, 2004. 62. el-Karaksy, H. et al., Capillaria philippinensis: A cause of fatal diarrhea in one of two infected Egyptian sisters. J. Trop. Pediatr., 50, 57, 2004. 63. Saichua, P., Nithikathkul, C., and Kaewpitoon, N., Human intestinal capillariasis in Thailand. World J. Gastroenterol., 14, 506, 2008. 64. Pereira, V.G., and Mattosinho Franca, L.C., Successful treatment of Capillaria hepatica infection in an acutely ill adult. Am. J. Trop. Med. Hyg., 32, 1272, 1983. 65. Rana, S.S. et al., Human intestinal capillariasis: Diagnosis by jejunal �uid analysis obtained at enteroscopy and reversal of subtotal villous atrophy after treatment. Endoscopy, 41 (suppl 2), E102, 2009. 66. Juncker-Voss, M. et al., Serological detection of Capillaria hepatica by indirect immuno�uorescence assay. J. Clin. Microbiol., 38, 431, 2000. 67. Assis, B.C. et al., A contribution to the diagnosis of Capillaria hepatica infection by indirect immuno�uorescence test. Mem. Inst. Oswaldo Cruz, 99, 173, 2004. 68. El Dib, N.A. et al., Evaluation of Capillaria philippinensis coproantigen in the diagnosis of infection. J. Egypt. Soc. Parasitol., 34, 97, 2004. 69. Intapan, P.M. et al., Potential use of Trichinella spiralis antigen for serodiagnosis of human capillariasis philippinensis by immunoblot analysis. Parasitol. Res., 98, 227, 2006. 70. Intapan, P.M. et al., An enzyme-linked immunosorbent assay as screening tool for human intestinal capillariasis. Southeast Asian J. Trop. Med. Public Health, 41, 298, 2010. 71. Zhu, X. et al., Mitochondrial DNA polymorphism within and among species of Capillaria sensu lato from Australian marsupials and rodents. Int. J. Parasitol., 30, 933, 2000. 50 Chapter 50 - Dioctophyme
1. Bailly, M.L., Leuzinger, U., and Bouchet, F., Dioctophymidae eggs in coprolites from neolithic site of Arbon-Bleiche 3 (Switzerland). J. Parasitol., 89, 1073, 2003.
2. Tokiwa, T. et al., Dioctophyme renale (Nematoda: Dioctophymatoidea) in the abdominal cavity of Rattus norvegicus in Japan. Parasitol. Int., 60, 324, 2011.
3. Pedrassani, D. et al., Morphology of eggs of Dioctophyme renale Goeze, 1782 (Nematoda: Dioctophymatidae) and in�uences of temperature on development of �rst-stage larvae in the eggs. Rev. Bras. Parasitol. Vet., 18, 15, 2009.
4. Measures, L.N., and Anderson, R.C., Centrarchid �sh as paratenic hosts of the giant kidney worm, Dioctophyma renale (Goeze, 1782). J. Wildl. Dis., 21, 11, 1985.
5. Mace, T.F., and Anderson, R.C., Development of the giant kidney worm, Dioctophyma renale (Goeze, 1782) (Nematoda: Dioctophymatoidea). Can. J. Zool., 53, 1552, 1975.
6. Hodesson, D.L., Daniels, R.B., and Richards, B., Dioctophyma renale in a dog (a case report). Vet. Med. Small Anim. Clin., 75, 825, 1980.
7. Alvarenga, J. et al., Dioctophyma renale in a dog. Mod. Vet. Pract., 65, 125, 1984.
8. Barros, D.M., Lorini, M.L., and Persson, V.G., Dioctophymosis in the Little Grison (Galictis cuja). J. Wildl. Dis., 26, 538, 1990.
9. Dyer, N.W., Dioctophyma renale in ranch mink. J. Vet. Diagn. Invest., 10, 111, 1998. 10. Mech, L.D., and Tracy, S.P., Prevalence of giant kidney worm (Dioctophyma renale) in wild mink (Mustela vison) in Minnesota. Am. Midland Natural., 145, 206, 2001. 11. Nakagawa, T.L. et al., Giant kidney worm (Dioctophyma renale) infections in dogs from Northern Paraná, Brazil. Vet. Parasitol., 145, 366, 2007. 12. Ribeiro, C.T., Verocai, G.G., and Tavares, L.E., Dioctophyme renale (Nematoda, Dioctophymatidae) infection in the crabeating fox (Cerdocyon thous) from Brazil. J. Wildl. Dis., 45, 248, 2009. 13. Verocai, G.G. et al., Dioctophyme renale (Goeze, 1782) in the abdominal cavity of a domestic cat from Brazil. Vet. Parasitol., 161, 342, 2009. 14. Ishizaki, M.N. et al., Dioctophyma renale (Goeze, 1782) in the abdominal cavity of a capuchin monkey (Cebus apella), Brazil. Vet. Parasitol., 173, 340, 2010. 15. Jorde, D.G., Occurrence of Dioctophyma renale (Goeze, 1782) in mink from North Dakota. J. Wildl. Dis., 16, 381, 1980. 16. Hanjani, A.A. et al., The �rst report of human infection with Dioctophyma renale in Iran. Trans. R. Soc. Trop. Med. Hyg., 62, 647, 1968. 17. Myers, B.J., Nematodes transmitted to man by �sh and aquatic mammals. J. Wildl. Dis., 6, 266, 1970. 18. Beaver, P.C., and Khamboonruang, C., Dioctophyma-like nematode in a subcutaneous nodule from man in northern Thailand. Am. J. Trop. Med. Hyg., 33, 1032, 1984. 19. Beaver, P.C. and Theis, J.H., Dioctophymatid larval nematode in a subcutaneous nodule from man in California. Am. J. Trop. Med. Hyg., 28, 206, 1979. 20. Fernando, S.S., The giant kidney worm (Dioctophyma renale) infection in man in Australia. Am. J. Surg. Pathol., 7, 281, 1983. 21. Sun, T. et al., Giant kidney worm (Dioctophyma renale) infection mimicking retroperitoneal neoplasm. Am. J. Surg. Pathol., 10, 508, 1986. 22. Narváez, J.A. et al., Hyperdense renal cystic lesions caused by Dioctophyma renale. Am. J. Roentgenol., 163, 997, 1994. 23. Urano, Z. et al., Dioctophymatid nematode larva found from human skin with creeping eruption. J. Parasitol., 87, 462, 2001. 24. Ignjatovic, I. et al., Infestation of the human kidney with Dioctophyma renale. Urol. Int., 70, 70, 2003. 25. Pilsczek, F.H., Helminthic infections mimicking malignancy: A review of published case reports. J. Infect. Dev. Ctries., 4, 425, 2010. 26. Li, G. et al., Fatal bilateral dioctophymatosis. J. Parasitol., 96, 1152, 2010. 27. Gutierrez, Y., Cohen, M., and Machicao, C.N., Dioctophyme larva in the subcutaneous tissues of a woman in Ohio. Am. J. Surg. Pathol., 13, 800, 1989. 28. Sardjono, T.W. et al., Dioctophymatosis renalis in humans: First case report from Indonesia. Proc. ASEAN Congr. Trop. Med. Parasitol., 3, 90, 2008. 29. Ferreira, V.L. et al., Dioctophyma renale in a dog: Clinical diagnosis and surgical treatment. Vet. Parasitol., 168, 151, 2010. 30. Anderson, R.C., and Bain, O., Keys to genera of the superfamilies Rhabditoidea, Dioctophymatoidea, Trichinelloidea and Muspiceoidea. In: Anderson, R.C., Chabaud, A.G., and. Willmott, S. (eds.). CIH Keys to the Nematode Parasites of Vertebrates. No. 9, Commonwealth Agricultural Bureaux, Farnham Royal, Bucking-hamshire, U.K. pp. 1–26, 1982. 51 Chapter 51 - Dirofilaria
1. Addario, C. Su un nematode dell’occhio umano. Ann. Oftalm. 14, 134, 1885.
2. Dashiell, G.F. A case of diro�lariasis involving the lung. Am. J. Trop. Med. Hyg. 10, 37, 1961.
3. Anderson, R.C. Description and relationship of Diro�laria ursi Yamaguti, 1941, and a review of the genus Diro�laria Railliet and Henry, 1911. Trans. Royal Can. Inst. 29, 35, 1952.
4. Faust, E.C. Mammalian heart worms of the genus Diro�laria, In: Festschrift Berhanrd Nocht zum 80. Geburtstag, Hamburg, p. 131, 1937.
5. Canestri Trotti, G. et al. The species of genus Diro�laria Railliet & Henry, 1911. Parassitologia, 39, 369, 1997.
6. Manfredi, M.T. et al. Phylogeny, systematics and structural aspects. In: Simon, F., and Genchi, C. (eds), Heartworm Infection in Humans and Animals, Ediciones Universidad de Salamanca, Salamanca, España, p. 19, 2001.
7. Wong, M.M., and Brummer, M.E.G. Cuticular morphology of �ve species of Diro�laria: A scanning electron microscope study. J. Parasitol. 64, 108, 1978. 8. Anderson, R.C. Nematode Parasites of Vertebrates. Their Development and Transmission. CAB International Publishing, Wallingford, Oxon, UK, 467pp, 2000. 9. Manfredi, M.T. et al. Biology of �larial worms parasitizing dogs and cats. Proceedings First European Diro�laria Days, Zagreb, February 22–25, 2007. 10. Nozais, J.P. et al. A case of subcutaneous Diro�laria (Nochtiella) repens with micro�laremia originating in Corsica. Bull. Soc. Pathol. Exot. 87, 183, 1994. 11. Coluzzi, M. Osservazioni sperimentali sul comportamento di Diro�laria repens in diversi gruppi di artropodi ematofagi. Parassitologia 6, 57, 1964. 12. Ludlam, K.W. et al. Potential vectors of Diro�laria immitis. JAVMA 157, 1354, 1970. 13. Genchi, C. et al. Climate and Diro�laria infection in Europe. Vet. Parasitol. 163, 286, 2009. 14. Prieto, G. et al. Seroepidemiology of Diro�laria immitis and Diro�laria repens in humans from three areas of southern Europe. Res. Rev. Parasitol. 60, 95, 2000. 15. Donahoe, J.M.R. Clinical aspects of feline diro�lariasis. In: Proceedings of Heartworm Symposium. VM Publishing, Bonner Spring, p. 59, 1975. 16. Otto, G.F. Changing geographic distribution of heartworm disease in the United States. In: Morgan, H.C. Proceedings Heartworm Symposium ‘74. Veterinary Medicine, Bonner Springs, Kansas, USA, 1975. 17. Genchi, C. et al. Epidemiology of canine and feline diro�lariasis: A global view. In: Simon, F., and Genchi, C. (eds), Heartworm Infection in Humans and Animals. Ediciones Universidad de Salamanca, Salamanca, España, p. 121, 2001. 18. Bolio-Gonzalez, M.E. et al. Prevalence of Diro�laria immitis infection in dogs from Meridia, Yucatan, Mexico. Vet. Parasitol. 148, 166, 2007. 19. Ryan, W.G., and Newcomb, K.M. Prevalence of feline heartworm disease—A global review. In: Soll, M.D. (ed.), Proceedings of the Heartworm Symposium ‘95, Auburn, AL, The American Heartworm Society, Batavia, IL, p. 79, 1995. 20. McCall, J.W. et al. Heartworm disease in animals and humans: A review. Adv. Parasitol. 66, 193, 2008. 21. Milanez de Campos, J.R. et al. Human pulmonary diro�lariasis: Analysis of 24 cases from Sao Paulo, Brazil. Chest, 112, 729, 1997. 22. Vezzani, D. et al. Diro�lariasis in Argentina: Historical review and �rst report of Diro�laria immitis in a natural mosquito population. Vet. Parasitol. 136, 259, 2006. 23. Miyoshi, T. et al. Human pulmonary diro�lariasis: A case report and review of the recent Japanese literature. Respirology 11, 343, 2006. 24. Simón, F. et al. What is new about animal and human diro�lariosis? Trends Parasitol. 25, 404, 2009. 25. Pampiglione, S. et al. Human diro�lariasis due to Diro�laria (Nochtiella) repens: A review of world literature. Parassitologia 37, 149, 1995. 26. Pampiglione, S., and Rivasi, F. Human diro�lariasis due to Diro�laria (Nochtiella) repens: An update of world literature from 1995 to 2000. Parassitologia 42, 231, 2000. 27. Dissanaike, A.S. et al. Human infection caused by Diro�laria (Nochtiella) repens in Sri Lanka. Parassitologia 39, 375, 1997. 28. Orihel, T.C., and Ash, L. Parasites in Human Tissues. ASCPL [American Society of Clinical Pathologists] Press, Chicago, p. 386, 1995. 29. Beaver, P.C. et al. Diro�laria ursi-like parasites acquired by humans in the northern United States and Canada: Report of two cases and brief review. Am. J. Trop. Med. Hyg. 37, 357, 1987.
30. Orihel, T.C., and Eberhard, M.L. Zoonotic �lariasis. Clin. Microbiol. Rev. 11, 366, 1998.
31. Orihel, T.C., and Isbey, E.K. Diro�laria striata infection in a North Carolina child. Am. J. Trop. Med. Hyg. 42, 124, 1990.
32. Beaver, P.C. et al. In: Faust, E.C., Campbell, H.E., and Kellogg, C.R. (eds), Clinical Parasitology. 9th edn. Lea & Febiger, Philadelphia, PA, 1984.
33. Theis, J.H. et al. Distribution, prevalence, and relative risk of �lariasis in dogs from the state of Washington (1997–1999). J. Am. An. Hosp. Ass. 37, 339, 2001.
34. Pampiglione, S. et al. Human Diro�laria repens infection in Hungary: A case in the spermatic cord and a review of the literature. Acta Vet. Acad. Sci. Hung. 47, 77, 1999.
35. Collins, B.M. et al. Diro�laria tenuis infection of the oral mucosa and cheek. J. Oral Maxillofac. Surg. 51, 1037, 1993.
36. Marty, A.M., and Nea�e, R.C. Pathology of Infectious Diseases, Vol. I, Helminthiases. Washington, DC, Armed Forces Institute of Pathology, 2000.
37. Pampiglione, S. et al. Pitfalls and dif�culties in histological diagnosis of human diro�lariasis due to Diro�laria (Nochtiella) repens. Diag. Microbiol. Infect. Dis. 34, 57, 1999.
38. Pampiglione, S. et al. Diro�larial human cases in the Old World, attributed to Diro�laria immitis: A critical analysis. Histopathology 54, 192, 2009.
39. Simon, F. et al. Human humoral immune response to Diro�laria species. Parassitologia 39, 397, 1997.
40. Santamaria, B. et al. Evaluation of Diro�laria immitis excretory/secretory products for seroepidemiological studies on human diro�lariasis. Parasite 2, 269, 1995.
41. Simón, F. et al. Immunity in the vertebrate hosts. Simón, F., and Genchi, C. (eds), Ediciones Universidad de Salamanca. Salamanca, España, 83, 2001.
42. Song, K.H. et al. Cross-reactivity between sera from dogs experimentally infected with Diro�laria immitis and crude extract of Toxocara canis. Korean J. Parasitol. 40, 195, 2002.
43. Simón, F. et al. Immunoglobulin G antibodies against the endosymbionts of �larial nematodos (Wolbachia) in patients with pulmonary diro�lariasis. Clin. Diagn. Lab. Immunol. 10, 180, 2003.
44. Grandi, G. Wolbachia in Diro�laria repens causing human subcutaneous diro�lariosis. J. Parasitol. 94, 1421, 2008.
45. Cancrini, G. et al. Identi�cation of Diro�laria in man by multi locus electrophoretic analysis. Ann. Trop. Med. Pad. 85, 529, 1991. 46. Thanomsub, B.W. et al. Differential diagnosis of human lymphatic �lariasis using PCR-RFLP. Mol. Cell. Probes 14, 41, 2000. 47. Favia, G. et al. Polymerase chain reaction-identi�cation of Diro�laria repens and Diro�laria immitis. Parasitology, 113, 567, 1996. 48. Poole, C.B. et al. Cloning of a cuticular antigen that contains multiple tandem repeats from the �larial parasite Diro�laria immitis. Proc. Natl. Acad. Sci. USA 89, 5986, 1992. 49. Scoles, A.G., and Kambhamptati, S. Polymerase chain reaction-based method for detection of canine heartworm (Filaria: Onchocercidae) in mosquitoes (Diptera: Culicidae) and vertebrate hosts. J. Med. Entomol. 32, 864, 1995. 50. Chandrasekharan, N.V. et al. Diro�laria repens: Cloning and characterization of a repeated DNA sequence for the diagnosis of diro�lariasis in dogs, Canis familiaris. Exp. Parasitol. 78, 279, 1994. 51. Vakalis, N. et al. Improved detection of Diro�laria repens DNA by direct polymerase chain reaction. Parasitol. Int. 48, 145, 1999. 52. Rivasi, F. et al. Detection of Diro�laria (Nochtiella) repens DNA by polymerase chain reaction in embedded paraf�n tissues from two human pulmonary locations. APMIS, 114, 566, 2006. 53. Mar, P.H. et al. Speci�c polymerase chain reaction for differential diagnosis of Diro�laria immitis and Diro�laria reconditum using primers derived from internal transcribed spacer region 2 (ITS2). Vet. Parasitol. 106, 243, 2002. 54. Nuchprayoon, S. et al. Detection of �larial parasites in domestic cats by PCR-RFLP of ITS1. Vet. Parasitol. 140, 366, 2006. 55. Huang, H. et al. Molecular characterization and phylogenetic analysis of Diro�laria immitis of China based on COI and 12S rDNA genes. Vet. Parasitol. 160, 175, 2009. 56. Wright, D.K., and Manos, M. Sample preparation from paraf�n-embedded tissue. In: Innis, M.A., Glefand, D.H., and Sninsky, J.J. (eds), PCR Protocol: A Guide to Methods and Application. Academic, San Diego, pp. 153–158, 1990. 57. Strauss, W.M. Preparation of genomic DNA from mammalian tissue, in Current Protocols. In: Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidnan, J.G., Smith, J.A., and Struhl, K. (eds), Molecular Biology. John Wiley & Sons, New York, pp. 2.2.1–2.2.3, 1995. 58. Sambrook, J., Fritsch, E.F., and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp. 9.14–9.23, 1989.
52 Chapter 52 - Dracunculus
1. Muller, R., Dracunculus and dracunculiasis. Adv. Parasitol., 9, 73, 1971.
2. Muller, R., Guinea worm disease; epidemiology, control and treatment. Bull. WHO, 57, 683, 1979.
3. Jones, H.I., and Mulder, E., Dracunculus mulbus n. sp. (Nematoda: Spirurida) from the water python Liasis fuscus (Serpentes: Boidae) in northern Australia. Syst. Parasitol., 66, 195, 2007.
4. Moravec, F., and Santos, C.P., Dracunculus brasiliensis sp. n. (Nematoda: Dracunculidae) from the anaconda, Eunectes murinus (Ophidia: Boidae). Parasitol. Res., 104, 589, 2009.
5. Watts, S.J., Dracunculiasis in Africa: Its geographic extent, incidence, and at-risk population. Am. J. Trop. Med. Hyg., 37, 119, 1987.
6. Adamson, P., Dracontiasis in antiquity. Med. Hist., 32, 204, 1988.
7. Eberhard, M.L., Rab, M.A., and Dilshad, M.N., Red Dracunculus medinensis. Am. J. Trop. Med. Hyg., 41, 479, 1989.
8. Greenaway, C., Dracunculiasis (guinea worm disease). CMAJ, 170, 495, 2004.
9. Moravec, F., Some aspects of the taxonomy and biology of dracunculoid nematodes parasitic in �shes: A review. Folia Parasitol., 51, 1, 2004.
10. Moravec, F., and Little, M.D., Redescription of Dracunculus globocephalus Mackin, 1927 (Nematoda: Dracunculidae), a parasite of the snapping turtle, Chelydra serpentina. Folia Parasitol., 51, 339, 2004.
11. Bimi, L., Potential vector species of Guinea worm (Dracunculus medinensis) in Northern Ghana. Vector Borne Zoonotic Dis., 7, 324, 2007.
12. Bimi, L. et al., Differentiating Dracunculus medinensis from D. insignis, by the sequence analysis of the 18S rRNA gene. Ann. Trop. Med. Parasitol., 99, 511, 2005.
13. Birare, S.D. et al., Guinea worm infection of urinary bladder manifesting as obstructive uropathy in rural Maharashtra. Trop. Doct., 35, 242, 2005.
14. Eberhard, M.L., and Brandt, F.H., The role of tadpoles and frogs as paratenic hosts in the life cycle of Dracunculus insignis (Nematoda: Dracunculoidea). J. Parasitol., 81, 792, 1995.
15. Edungbola, L.D. et al., A method of rapid assessment of the distribution and endemicity of dracunculiasis in Nigeria. Soc. Sci. Med., 23, 550, 1986.
16. Cairncross, S., Anemana, S.D., and Olsen, A., Towards the eradication of guinea worm; a Danish-Ghanaian collaboration. Parasitol. Today, 15, 127, 1999. 17. Cairncross, S., Muller, R., and Zagaria, N., Dracunculiasis (guinea worm disease) and the eradication initiative. Clin. Microbiol. Rev., 15, 223, 2002. 18. Watts, S.J., Dracunculiasis in the Caribbean and South America: A contribution to the history of dracunculiasis eradication. Med. Hist., 45, 227, 2000. 19. Menon, T., Incidental �nding of Dracunculus medinensis in Australia. Med. J. Aust., 183, 51, 2005. 20. Ruiz-Tiben, E., and Hopkins, D.R., Dracunculiasis (Guinea worm disease) eradication. Adv. Parasitol., 61, 275, 2006. 21. Rwakimari, J.B., Hopkins, D.R., and Ruiz-Tiben, E., Uganda’s successful Guinea Worm Eradication Program. Am. J. Trop. Med. Hyg., 75, 3, 2006. 22. Mathur, S., and Gahlot, R.S., Calci�ed guinea worms in breast. J. Assoc. Physicians India, 54, 201, 2006. 23. Miri, E. et al., Nigeria’s triumph: Dracunculiasis eradicated. Am. J. Trop. Med. Hyg., 83, 215, 2010. 24. Centers for Disease Control and Prevention (CDC), Progress toward global eradication of dracunculiasis, January 2009– June 2010. Morb. Mortal. Wkly. Rep., 59, 1239, 2010. 25. Fabiansen, C., Harboe, Z.B., and Christensen, V., Dracunculiasis in South Sudan. Am. J. Trop. Med. Hyg., 82, 757, 2010. 26. Barry, M., The tail end of guinea worm—global eradication without a drug or a vaccine. N. Engl. J. Med., 356, 2561, 2007. 27. Iriemenam, N.C. et al., Dracunculiasis—the saddle is virtually ended. Parasitol. Res., 102, 343, 2008. 28. Brieger, W.R., Watts, S., and Yacoob, M., Guinea worm, maternal morbidity and child health. J. Trop. Paediatr., 35, 285, 1989. 29. Adeyeba, O.A., and Kale, O.O., Epidemiology of dracunculiasis and its socio-economic impact in a village in south-west Nigeria. West Afr. J. Med., 10, 208, 1991. 30. Hakim, F.A., and Khan, N.N., Dracunculiasis. An incidental diagnosis in a Saudi female. Saudi Med. J., 28, 1438, 2007. 31. Hopkins, D.R. et al., Dracunculiasis, onchocerciasis, schistosomiasis, and trachoma. Ann. N.Y. Acad. Sci., 1136, 45, 2007. 32. Adeyeba, O.A., Secondary infection in dracunculiasis: Bacteria and morbidity. Int. J. Zoonoses, 12, 147, 1985. 33. Smith, G.S. et al., Disability from dracunculiasis: Effect on mobility. Ann. Trop. Med. Parasitol., 83, 151, 1989. 34. Imtiaz, R., Hopkins, D.R., and Ruiz-Tiben, E., Permanent disability from dracunculiasis. Lancet, 336, 630, 1990. 35. Glenshaw, M.T. et al., Guinea worm disease outcomes in Ghana: Determinants of broken worms. Am. J. Trop. Med. Hyg., 81, 305, 2009. 36. Chippaux, J.P., Mebendazole treatment of dracunculiasis. Trans. R. Soc. Trop. Med. Hyg., 85, 280, 1991. 37. Bloch, P., Simonsen, P.E., and Vennervald, B.J., The antibody response to Dracunculus medinensis in an endemic human population of northern Ghana. J. Helminthol., 67, 37, 1993. 38. Bloch, P., and Simonsen, P.E., Studies on immunodiagnosis of dracunculiasis. I. Detection of speci�c serum antibodies. Acta Trop., 70, 73, 1998. 39. Bloch, P., Vennervald, B.J., and Simonsen, P.E., Studies on immunodiagnosis of dracunculiasis. II. Search for circulating antigens. Acta Trop., 70, 303, 1998. 40. Bapna, S., and Renapurkar, D.M., Immunodiagnosis of early dracunculiasis. J. Commun. Dis., 28, 33, 1996. 41. Bloch, P., and Simonsen, P.E., Immunoepidemiology of Dracunculus medinensis infections I. Antibody responses in relation to infection status. Am. J. Trop. Med. Hyg., 59, 978, 1998.
42. Bloch, P., and Simonsen, P.E., Immunoepidemiology of Dracunculus medinensis infections II. Variation in antibody responses in relation to transmission season and patency. Am. J. Trop. Med. Hyg., 59, 985, 1998.
43. Knopp, S. et al., Antibody and cytokine responses in Dracunculus medinensis patients at distinct states of infection. Trans. R. Soc. Trop. Med. Hyg., 102, 277, 2008.
44. Wijová, M. et al., Phylogenetic position of Dracunculus medinensis and some related nematodes inferred from 18S rRNA. Parasitol. Res., 96, 133, 2005. 45. Elsasser, S.C. et al., Species identi�cation of North American guinea worms (Nematoda: Dracunculus) with DNA barcoding. Mol. Ecol. Resour., 9, 707, 2009. 46. Centers for Disease Control and Prevention (CDC), Renewed transmission of dracunculiasis—Chad, 2010. Morb. Mortal. Wkly. Rep., 60, 744, 2011. 53 Chapter 53 - Enterobius
TABLE 53.1
Primers Targeting Enterobius vermicularis CO1, ITS2, and 5S rRNA Genes
Gene Primer Sequence (5′-3′) PCR Conditions
CO1 pr-a TGGTTTTTTGTGCATCCTGAGGTTTA 1 × 94°C for 1 min; 30 × 94°C for 1 min, 40°C for 1 min, 72°C for 2 min; 1 × 72°C for 7 min pr-b AGAAAGAACGTAATGAAAATGAGCAAC
ITS2 LC1 CGAGTATCGATGAAGAACGCAGC 1 × 94°C for 1 min; 30 × 94°C for 1 min, 52°C for 3 min, 72°C for 2 min; 1 × 72°C for 7 min HC2 ATATGCTTAAGTTCAGCGGG
5S rRNA Ent 5SF GCGAATTCTTGGATCGGAGACGGCCTG 1 × 94°C for 1 min; 30 × 94°C for 1 min, 40°C for 1 min, 72°C for 2 min; 1 × 72°C for 7 min Ent 5SR GCTCTAGACGAGATGTCGTGCTTTCAACG
7. Hasegawa, H., and Udono, T., Chimpanzee pinworm, Enterobius anthropopitheci (Nematoda: Oxyuridae), maintained for more than twenty years in captive chimpanzees in Japan. J. Parasitol., 93, 850, 2007.
8. Vermund, S.H., and Wilson, C.M., Pinworm (Enterobius vermicularis). Sem. Pediat. Infect. Dis., 11, 252, 2000.
9. Grencis, R.K., and Cooper, E.S., Enterobius, Trichuris, Capillaria, and hookworm including Ancylostoma caninum. Gastroenterol. Clin. North. Am., 25, 579, 1996.
10. Ashford, R.W., Hart, C.A., and Williams, R.G., Enterobius vermicularis infection in a children’s ward. J. Hosp. Infect., 12, 221, 1988.
11. Herrstrom, P. et al., Enterobius vermicularis and �nger sucking in young Swedish children. Scand. J. Prim. Health Care, 15, 146, 1997.
12. Lohiya, G.S. et al., Epidemiology and control of enterobiasis in a developmental center. West J. Med., 172, 305, 2000.
13. Ogunji, F.O., Post coital pinworm infection. J. Hyg. Epidemiol. Microbiol. Immunol., 27, 103, 1983.
14. Williams, D.J., and Dixon, M.F., Sex, Enterobius vermicularis and the appendix. Br. J. Surg., 75, 1225, 1988.
15. Abdolrasouli, A., and Hart, J., Oral-anal intercourse and sexual transmission of Enterobius vermicularis; do we need to screen for other intestinal parasites? Int. J. STD AIDS, 20, 739, 2009.
16. Mahdi, N.K., and al Khfaji, A.A., Prevalence and seasonal variation of enterobiasis in children of Iraq. Southeast Asian J. Trop. Med. Public Health, 21, 135, 1990.
17. Fan, P.C., Review of enterobiasis in Taiwan and offshore islands. J. Microbiol. Immunol. Infect., 31, 203, 1998.
18. Lee, K.J., Lee, I.Y., and Im, K.I., Enterobius vermicularis egg positive rate in a primary school in Chungchongnam-do (Province) in Korea. Korean J. Parasitol., 38, 177, 2000.
19. Sirivichayakul, C. et al., Prevalence of enterobiasis and its incidence after blanket chemotherapy in a male orphanage. Southeast Asian J. Trop. Med. Public Health, 31, 144, 2000.
20. Sung, J.F. et al., Pinworm control and risk factors of pinworm infection among primary-school children in Taiwan. Am. J. Trop. Med. Hyg., 65, 558, 2001.
21. Kim, B.J. et al., Egg positive rate of Enterobius vermicularis of primary school children in Geoje island. Korean J. Parasitol., 41, 75, 2003.
22. Song, H.J. et al., Prevalence and risk factors for enterobiasis among preschool children in a metropolitan city in Korea. Parasitol. Res., 91, 46, 2003.
23. Park, J.H. et al., A survey of Enterobius vermicularis infection among children on western and southern coastal islands of the Republic of Korea. Korean J. Parasitol., 43, 129, 2005.
24. Remm, M., Distribution of enterobiasis among nursery school children in SE Estonia and of other helminthiases in Estonia. Parasitol. Res., 99, 729, 2006.
25. Jang, M.H. et al., Prevalence of Enterobius vermicularis infection among children attending preschools in Taipei City. Taipei City Med. J., 4, 565, 2007.
26. Chang, T.K. et al., Prevalence of Enterobius vermicularis infection among preschool children in kindergartens of Taipei City, Taiwan in 2008. Korean J. Parasitol., 47, 185, 2009.
27. Wang, C.C. et al., Current status of Enterobius vermicularis infection in primary schoolchildren in Miaoli County and Taichung County, Taiwan. J. Microbiol. Immunol. Infect., 42, 420, 2009.
28. Wang, L.C., Hwang, K.P., and Chen, E.R., Enterobius vermicularis infection in schoolchildren: A large-scale survey 6 years after a population-based control. Epidemiol. Infect., 138, 28, 2010. 29. Iñiguez, A.M. et al., Enterobius vermicularis: Ancient DNA from North and South American human coprolites. Mem. Inst. Oswaldo Cruz, 98, S67, 2003. 30. Iñiguez, A.M. et al., SL1 RNA gene recovery from Enterobius vermicularis ancient DNA in pre-Columbian human coprolites. Int. J. Parasitol., 36, 1419, 2006. 31. Ganor, S., In whom does pinworm infection itch? Int. J. Dermnatol., 26, 667, 1987. 32. Raghallaigh, S.N., and Powell, F.C., Enterobius vermicularis dermatitis. Clin. Exp. Dermatol., 35, e32, 2010. 33. Cook, G.C., Enterobius vermicularis infection. Gut, 35, 1159, 1994. 34. Vafai, M., and Mohit, P., Granuloma of the anal canal due to Enterobius vermicularis. Report of a case. Dis. Colon Rectum, 26, 349, 1983. 35. Mayayo, E. et al., Pelvic oxyuriasis. Acta Obstet. Gynecol. Scand., 65, 805, 1986. 36. Gupta, S.C. et al., Pathology of tropical appendicitis. J. Clin. Pathol., 42, 1169, 1989. 37. Mondou, E.N., and Gnepp, D.R., Hepatic granuloma resulting from Enterobius vermicularis. Am. J. Clin. Pathol., 91, 97, 1989. 38. Goldenberg, S.P., and Marignani, P., The endoscopic diagnosis of colonic enterobiasis. Gastrointest. Endosc., 36, 309, 1990. 39. Sinniah, B. et al., Enterobiasis: A histopathological study of 259 patients. Ann. Trop. Med. Parasitol., 85, 625, 1991. 40. Wiebe, B.M., Appendicitis and Enterobius vermicularis. Scand. J. Gastroenterol., 26, 336, 1991. 41. Kollias, G., Kyriakopoulos, M., and Tiniakos, G., Epididymitis from Enterobius vermicularis: Case report. J. Urol., 147, 1114, 1992. 42. Mattia, A.R., Perianal mass and recurrent cellulitis due to Enterobius vermicularis. Am. J. Trop. Med. Hyg., 47, 811, 1992. 43. Tornieporth, N.G. et al., Ectopic enterobiasis: A case report and review. J. Infect., 24, 87, 1992. 44. Dalimi, A., and Khoshzaban, F., Comparative study of two methods for the diagnosis of Enterobius vermicularis in the appendix. J. Helminthol., 67, 85, 1993. 45. Patterson, L.A. et al., Perforation of the ileum secondary to Enterobius vermicularis: Report of a rare case. Mod. Pathol., 6, 781, 1993. 46. Dahlstrom, J.E., and Macarthur, E.B., Enterobius vermicularis: A possible cause symptoms resembling appendicitis. Aust. N. Z. J. Surg., 64, 692, 1994. 47. Sterba, J., and Vicek, M., Appendiceal enterobiasis—its incidence and relationships to appendicitis. Folia Parasitol., 31, 311, 1984. 48. Liu, L.X. et al., Eosinophilic colitis associated with larvae of the pinworm Enterobius vermicularis. Lancet, 346, 410, 1995. 49. Surmont, I., and Liu, L.X., Enteritis, eosinophilia, and Enterobius vermicularis. Lancet, 346, 1167, 1995. 50. Beattie, R., Walker-Smith, A., and Domizio, P., Ileal and colonic ulceration due to Enterobius. J. Paediatr., 132, 1055, 1995. 51. Gazvani, M.R., and Emery, S.J., Intraperitoneal Enterobius vermicularis infection: A case report. Infect. Dis. Obstet. Gynecol., 4, 28, 1996. 52. Zoorob, R.J., Appendiceal colic caused by Enterobius vermicularis. J. Am. Board Fam. Pract., 9, 57, 1996. 53. Ajao, O.G. et al., Enterobius vermicularis (pin worm) causing symptoms of appendicitis. Trop. Doct., 27, 182, 1997. 54. Cacopardo, B. et al., Eosinophilic ileocolitis by Enterobius vermicularis: A description of two rare cases. Ital. J. Gastroenterol. Hepatol., 29, 51, 1997.
55. Arora, V.K. et al., Fine needle aspiration diagnosis of a subcutaneous abscess from Enterobius vermicularis infestation, A case report. Acta Cytol., 41, 1845, 1997.
56. Gatti, S. et al., Intestinal parasitic infections in an institution for the mentally retarded. Ann. Trop. Med. Parasitol., 94, 453, 2000.
57. Macedo, T., and MacCarty, R., Eosinophilic ileocolitis secondary to Enterobius vermicularis case report, Abdom. Imaging, 25, 530, 2000.
58. Saxena, A.K. et al., Laparoscopic appendectomy in children with Enterobius vermicularis. Surg. Laparosc. Endosc. Percutan. Tech., 11, 284, 2001.
59. Kumar, N. et al., Perianal nodule due to enterobiasis: An aspiration cytologic diagnosis. Diagn. Cytopathol., 28, 58, 2003.
60. Mahomed, A.A. et al., Enterobius vermicularis and perianal sepsis in children. Pediat. Surg. Int., 19, 740, 2003.
61. Arca, M.J. et al., Clinical manifestations of appendiceal pinworms in children: An institutional experience and a review of literature. Pediatr. Surg. Int., 20, 372, 2004.
62. Nackley, A.C. et al., Appendiceal Enterobius vermicularis infestation associated with right-sided chronic pelvic pain. JSLS, 8, 171, 2004.
63. Nordstrand, A.J., and Jayasekera, L.K., Enterobius vermicularis and clinical appendicitis: Worms in the vermiform appendix. Aust. N. Z. J. Surg., 74, 1024, 2004.
64. Petro, M., Iavu, K., and Minocha, A., Unusual endoscopic and microscopic view of Enterobius vermicularis: A case report with a review of the literature. South Med. J., 98, 927, 2005.
65. Jardine, M., Kokai, G., and Dalzell, M., Enterobius vermicularis and colitis in children. J. Pediatr. Gastroenterol. Nutr., 43, 610, 2006.
66. Quasem, A., and Salam, A., Ectopic enterobiasis: A case report and review of literature. Pak. J. Med. Sci., 23, 785, 2007.
67. Ariyarathenam, A.V. et al., Enterobius vermicularis infestation of the appendix and management at the time of laparoscopic appendicectomy: Case series and literature review. Int. J. Surg., 8, 466, 2010.
68. Cateau, E. et al., Enterobius vermicularis in the kidney: An unusual location. J. Med. Microbiol., 59, 860, 2010.
69. Hirai, Y. et al., Unusual colonoscopic view of Enterobius vermicularis. Intern. Med., 50, 657, 2011.
70. Beckman, E.N., and Holland, J.B., Ovarian enterobiasis—a proposed pathogenesis. Am. J. Trop. Med. Hyg., 30, 74, 1981.
71. Neri, A. et al., Enterobius (Oxyuris) vermicularis of the pelvic peritoneum—a cause of infertility. Eur. J. Obstet. Gyn. RB, 23, 239, 1986.
72. Mendoza, E. et al., Invasion of human embryo by Enterobius vermicularis. Arch. Pathol. Lab. Med., 111, 761, 1987.
73. Knuth, K.R. et al., Pinworm infestation of the genital tract. Am. Fam. Phys., 38, 127, 1988.
74. Gokalp, A. et al., Relation between Enterobius vermicularis infestation and dysuria, nocturia, enuresis nocturna and bacteriuria in primary school girls. Indian Pediatr., 28, 948, 1991.
75. Sun, T. et al., Enterobius egg granuloma of the vulva and peritoneum: Review of the literature. Am. J. Trop. Med. Hyg., 45, 249, 1991.
76. Schnell, V.L. et al., Enterobius vermicularis salpingitis: A distant episode from precipitating appendicitis. Obstet. Gynecol., 80, 553, 1992.
77. Hamblin, J., and Connor, P.D., Pinworms in pregnancy. J. Am. Board Fam. Pract., 8, 321, 1995.
78. McCabe, K. et al., Enterobiasis of the ovary in a patient with cervical carcinoma in situ. Infect. Dis. Obstet. Gynecol., 2, 231, 1995. 79. Chung, D.I. et al., Live female Enterobius vermicularis in the posterior fornix of the vagina of a Korean woman. Korean J. Parasitol., 35, 67, 1997. 80. al Rufaie, H.K. et al., Pinworms and postmenopausal bleeding. J. Clin. Pathol., 51, 401, 1998. 81. Avolio, L. et al., Perianal granuloma caused by Enterobius vermicularis: Report of a new observation and review of the literature. J. Paediatr., 132, 1055, 1998. 82. Dundas, K.C., Calder, A.A., and Alyusuf, R., Enterobius vermicularis threadworm infestation of paraovarian tissue in a woman who has had a hysterectomy. Brit. J. Obstet. Gynaecol., 106, 605, 1999. 83. Ok, U.Z. et al., Relationship between pinworm and urinary tract infections in young girls. APMIS, 107, 474, 1999. 84. Erhan, V. et al., Unilateral salpingitis due to Enterobius vermicularis. Int. J. Gynecol. Pathol., 19, 188, 2000. 85. Das, D.K. et al., Pelvic abscess from Enterobius vermicularis: Report of a case with cytologic detection of eggs and worms. Acta Cytol., 45, 425, 2001. 86. Tandan, T. et al., Pelvic in�ammatory disease associated with Enterobius vermicularis. Arch. Dis. Child., 86, 439, 2002. 87. Smolyakov, R. et al., Enterobius vermicularis infection of female genital tract: A report of three cases and review of the literature. Eur. J. Obstet. Gynecol. Reprod. Biol., 107, 220, 2003. 88. Sizer, A.R. et al., A pelvic mass due to infestation of the fallopian tube with Enterobius vermicularis. J. Obstet. Gynaecol., 24, 462, 2004. 89. Thomson, J.C., Pelvic pain caused by intraperitoneal Enterobius vermicularis (threadworm) ova with an associated systemic autoimmune reaction. J. Obstet. Gynaecol. Res., 30, 90, 2004. 90. Burkhart, C.N., and Burkhart, C.G., Assessment of frequency, transmission, and genitourinary complications of enterobiasis (pinworms). Int. J. Dermatol., 44, 837, 2005. 91. Siochou, A., Birtsou, H., and Papazahariadou, M., Enterobius vermicularis infection of female genital tract. Int. J. Immunopathol. Pharmacol., 21, 1031, 2008. 92. Craggs, B. et al., Enterobius vermicularis infection with tuboovarian abscess and peritonitis occurring during pregnancy. Surg. Infect. (Larchmt), 10, 545, 2009. 93. Konanahalli, P. et al., Enterobious vermicularis (pinworm) infestation of the vulva: Report of 2 cases of a pseudoneoplastic lesion mimicking squamous carcinoma. Int. J. Gynecol. Pathol., 29, 490, 2010. 94. Young, C. et al., Enterobius vermicularis infection of the fallopian tube in an infertile female. Pathol. Res. Pract., 206, 405, 2010. 95. Pampiglione, S., and Rivasi, F., Enterobiasis in ectopic locations mimicking tumor-like lesions. Int. J. Microbiol., 2009, 642481, 2009. 96. Murata, K. et al., Fatal infection with human pinworm, Enterobius vermicularis, in a captive chimpanzee. J. Med. Primatol., 31, 104, 2002. 97. Singh, S., and Samantaray, J.C., Topical anthelmintic treatment of recurrent genitourinary enterobiasis. Genitourin. Med., 65, 284, 1989. 98. Yang, Y.S. et al., Chemotherapeutic trial to control enterobiasis in schoolchildren. Korean J. Parasitol., 35, 265, 1997. 99. St Georgiev, V., Chemotherapy of enterobiasis (oxyuriasis). Expert Opin. Pharmacother., 2, 267, 2001. 100. Adungo, N.I., Ondijo, S.O., and Pamba, H.O., Observation of Enterobius vermicularis ova in urine: 3 case reports. East Afr. Med. J., 63, 676, 1986.
101. Cho, S.Y., A female Enterobius vermicularis picked up in a cellotape anal swab. Korean J. Parasitol., 35, 299, 1997.
102. Khalil, H.M. et al., Modi�cations in diagnosis and treatment of Enterobius vermicularis. J. Egypt. Soc. Parasitol., 17, 643, 1987.
103. Liu, L.X., Blaxter, M.L., and Shi, A., The 5S ribosomal RNA intergenic region of parasitic nematodes: Variation in size and presence of SL1 RNA. Mol. Biochem. Parasitol., 83, 235, 1996.
104. Iñiguez, A.M. et al., Enterobius vermicularis: Speci�c detection by ampli�cation of an internal region of 5S ribosomal RNA intergenic spacer and trans-splicing leader RNA analysis. E. vermicularis: Speci�c detection by PCR and SL1 RNA analysis. Exp. Parasitol., 102, 218, 2002. 105. Kang, S. et al., The mitochondrial genome sequence of Enterobius vermicularis (Nematoda: Oxyurida)–an idiosyncratic gene order and phylogenetic information for chromadorean nematodes. Gene, 429, 87, 2009. 106. Zelck, U.E., Bialek, R., and Weiß, M., Molecular phylogenetic analysis of Enterobius vermicularis and development of an 18S ribosomal DNA-rargeted diagnostic PCR. J. Clin. Microbiol., 49, 1602, 2011. 107. Piperaki, E.T. et al., Characterization of Enterobius vermicularis in a human population, employing a molecular-based method from adhesive tape samples. Mol. Cell. Probes, 25, 121, 2011. 54 Chapter 54 - Gnathostoma
1. Daengsvang, S. A monograph on the genus Gnathostoma & gnathostomiasis in Thailand, p. 1, SEAMIC/IMFJ, Tokyo, 1980.
2. Waikagul, J., and Diaz Chamacho, S.P. Gnathostomiasis. In: Darwin, K.D., and Fried, B. (eds), Food-Borne Parasitic Zoonoses, Fish and Plant-Borne Parasites, World Class Parasites, Chapter 11, p. 235, Springer, 2007.
3. Magana, M. et al. Gnathostomiasis: Clinicopathologic study. Am. J. Dermatopathol. 26, 91, 2004.
4. Ollaque, W., Ollaque, J., and Guevara de Veliz, A. Eosinophilic migratory nodular paniculitis in (human gnathostomiasis in Ecuador). 1st �nding of parasites in South America. Med. Cutan. Ibero. Lat. Am. 10, 73, 1982.
5. Villar de Ciprini, E. Paniculitis migratoria eosino�lica en el Peru, Gnathostoma como agente causal. Rev. Peru Med. Exp. Salud Publica 20, 220, 2003.
6. Kaminsky, C.A. et al. Eosinophilic migratory nodular panniculitis (human gnathostomiasis). Med. Cutan. Ibero. Lat. Am. 17, 158, 1989.
7. Nawa, Y., Ogata, K., and Imai, J. Gnathostomiasis by Gnathostoma doloresi. Saishin Igaku 44, 807, 1989. (In Japanese).
8. Ando, H. et al. Two human cases of gnathostomiasis and discovery of a second intermediate host of Gnathostoma nipponicum in Japan. J. Parasitol. 74, 626, 1988.
9. Nomura, Y. et al. Gnathostomiasis possibly caused by Gnathostoma malaysiae. Tokai J. Exp. Clin. Med. 25, 1, 2000.
10. Chai, J.Y. et al. An outbreak of gnathostomiasis among Korean emigrants in Myanmar. Am. J. Trop. Med. Hyg. 69, 67, 2003.
11. Akahane, H., Iwata, K., and Miyazaki, I. Studies on Gnathostoma hispidum Fedtchenko, 1872 parasitic loaches imported from China. Jpn. J. Parasitol. 31, 507, 1982. (In Japanese with English summary). 12. Montero, E. et al. Human gnathostomiasis in Spain: First report in humans. Acta Trop. 78, 59, 2001. 13. Herman, J.S. et al. Gnathostomiasis acquired by British tourists in Botswana. Emerg. Infect. Dis. 15, 594, 2009. 14. Hale, D.C., Blumberg, L., and Frean, J. Case report: Gnathostomiasis in two travelers to Zambia. Am. J. Trop. Med. Hyg. 68, 707, 2003. 15. LeRoux, P.L. Gnathostoma spinigerum in a lion in Northern Rhodesia. Trans. R. Soc. Trop. Med. Hyg. 52, 14, 1958. 16. Almeyda-Artigas, R.J., Bargues, M.D., and Mas-Coma, S. ITS-2 rDNA sequencing of Gnathostoma species (Nematoda) and elucidating of the species causing gnathostomiasis in the Americas. J. Parasitol. 86, 537, 2000. 17. Ngarmamonpirat, C. et al. Analysis of sequence variation in Gnathostoma spinigerum mitochondrial DNA by single- strand conformation polymorphism analysis and DNA sequence. Parasitol. Int. 54, 65, 2005. 18. Rojekittikhun, W. On the biology of Gnathostoma spinigerum. J. Trop. Med. Parasitol. 25, 91, 2002. 19. Rojekittikhun, W. et al. Record numbers of Gnathostoma larvae in swamp eels. J. Trop. Med. Parasitol. 24, 79, 2001. 20. Leon-Regagnon, V. et al. Gnathostomiasis in �sh from Tres Palos Lagoon, Guerrero, Mexico. Emerg. Infect. Dis. Letter to the editor, 2000. 21. Komalamisra, C., Nuamtanong, S., and Dekumyoy, P. Pila ampullacea and Pomacea canaliculata, as new paratenic hosts of Gnathostoma spinigerum. Southeast Asian J. Trop. Med. Public Health 40, 243, 2009. 22. Rojekittikhun, R., and Buchachart, K. The infectivitity of frozen Gnathostoma spinigerum encysted larvae in mice. J. Trop. Med. Parasitol. 25, 79, 2002. 23. Miyazaki, I. On the genus Gnathostoma and human gnathostomiasis, with special reference to Japan. Exp. Parasitol. 9, 338, 1960. 24. Nontasut, P. et al. Comparison of ivermectin and albendazole treatment for gnathostomiasis. Southeast Asian J. Trop. Med. Public Health 31, 374, 2000. 25. Chappuis, F., Farineli, T., and Loutan, L. Ivermectin treatment of a traveler who returned from Peru with cutaneous gnathostomiasis. Clin. Infect. Dis. 33, 17, 2001. 26. Parola, P. et al. Clinical picture: Gnathostomiasis. Lancet 357, 1011, 2001. 27. Dekumyoy, P. et al. A seven-year retrospective evaluation of gnathostomiasis and diagnostic speci�city by immunoblot. Proceedings of the �rst International Meeting on Gnatho stomiasis, p. 45, Culiacan, Sinaloa, Mexico, March 2002. 28. Suntharasamai, P. et al. ELISA for immunodiagnosis of human gnathostomiasis. Southeast Asian J. Trop. Med. Public Health 16, 274, 1985. 29. Dhamkrong-at, A. et al. Enzyme-linked immunosorbent assay for detection of antibody to Gnathostoma antigen in patients with intermittent cutaneous migratory swelling. J. Clin. Microbiol. 23, 847, 1986. 30. Nuchprayoon, S. et al. Study of speci�c IgG subclass antibodies for diagnosis of Gnathostoma spinigerum. Parasitol. Res. 91,137, 2003. 31. Nopparatana, C. et al. Puri�cation of Gnathostoma spinigerum speci�c antigen and immunodiagnosis of human gnathostomiasis. Int. J. Parasitol. 21, 677, 1991. 32. Chaicumpa, W. et al. Monoclonal antibody to a diagnostic Mr 24,000 antigen of Gnathostoma spinigerum. Int. J. Parasitol. 21, 677, 1991.
33. Rojekittikhun, W., Saito, S., and Sendo, F. Crude and monoclonal antibody af�nity-puri�ed larval Gnathostoma spinigerum larval antigens for immunodiagnosis of human gnathostomiasis. Southeast Asian J. Trop. Med. Public Health 26, 439, 1995.
34. Diaz-Camacho, S.P. et al. Clinical manifestations and immunodiagnosis of gnathostomiasis in Culiacan, Mexico. Am. J. Trop. Med. Hyg. 59, 908, 1998.
35. Ishiwata, K. et al. Evaluation of the antigenic similarities of adult-worm extracts from three Gnathostoma species, using sera from Mexican and Japanese patients with Gnathostoma infections. Ann. Trop. Med. Parasitol. 97, 629, 2003.
36. Anantaphuti, M.T., Nuamtanong, S., and Dekumyoy, P. Diagnostic value of IgG4 in gnathostomiasis. Trop. Med. Int. Health 10, 1013, 2005.
37. Laummaunwai, P. et al. Evaluation of human IgG class and subclass antibodies to a 24 kDa antigenic component of Gnathostoma spinigerum for the serodiagnosis of gnathostomiasis. Parasitol. Res. 101, 703, 2007.
38. Tapchaisri, P. et al. Gnathostoma spinigerum speci�c recombinant antigen for immunodiagnosis of human gnathostomiasis. 6th Seminar on Food-and Water-borne Parasitic Zoonoses (FBPZ6) incorporating the 3rd International Meeting on Gnathostomiasis (abstract). Bangkok, Thailand p. 52, 2–4 December 2009.
39. Uparananukraw, P. et al. Molecular cloning of a gene encoding matrix metalloproteinase-like protein from Gnathostoma spinigerum. Parasitol. Res. 87, 751, 2001.
40. Laummaunwai, P., Intapan, P.M., Wongkham, C., Lulitanond, V., and Maleewong, W. Identi�cation of antigenic components of Gnathostoma spinigerum advanced-third stage larvae by two-dimensional gel electrophoresis and mass spectro metry. Southeast Asian J. Trop. Med. Public Health, 39 (suppl. 1), 19, 2008.
41. Tapchaisri, P. et al. Speci�c antigen of Gnathostoma spinigerum for immunodiagnosis of human gnathostomiasis. Int. J. Parasitol. 21, 315, 1991.
42. Maleewong, W. et al. Serodiagnosis of human gnathostomiasis. Southeast Asian J. Trop. Med. Public Health 19, 201, 1988.
43. Tuntipopipat, S., Chawengkirttikul, R., and Sirisinha, S. A simpli�ed method for the fractionation of Gnathostomaspeci�c antigens for serodiagnosis of human gnathostomosis. J. Helminthol. 67, 297, 1993.
44. Soesatyo, M.H. et al. IgE responses in human gnathostomiasis. Trans. R. Soc. Trop. Med. Hyg. 81, 799, 1987. 45. Saksirisampant, W. et al. Speci�c IgE antibody responses to somatic and excretory–secretory antigens of third stage G. spinigerum larvae in human gnathostomiasis. J. Med. Assoc. Thai. 84, S173, 2001. 46. Boongird, P. et al. Neurological manifestation of gnathostomiasis. J. Neurol. Sci. 31, 279, 1977. 47. Rusnak, R.M., and Lucey, D.R. Clinical Gnathostomiasis. Case report and review of the English language literature. Clin. Infect. Dis. 16, 33, 1993. 48. Chandenier, J. et al. Medullary gnathostomiasis in a white patient: Use of immunodiagnosis and magnetic resonance imaging. Clin. Infect. Dis. 32, 154, 2001. 49. Sawanyawisuth, K. et al. MR imaging �ndings in cerebrospinal gnathostomiasis. Am. J. Neuroradiol. 25, 446, 2004. 50. Sawanyawisuth, K. et al. MR imaging �ndings in cauda equina gnathostomiasis. AJNR Am. J. Neuroradiol. 26, 39, 2005. 51. Hennies, F. et al. Gnathostomiasis: Import from Laos. JDDG 4, 414, 2006. 52. Ando, K. et al. Comparative study on DNA sequences of ribosomal DNA and cytochrome c oxidase subunit 1 of mitochondrial DNA among �ve species of gnathostomes. J. Helminthol. 80, 7, 2006. 53. Kifune, T. et al. Gnathostoma binucleatum (Spirurida: Gnathostomatidae) from the freshwater �sh in Tabasco, Mexico. Rev. Biol. Trop. 52, 371, 2004. 54. Martı´nez-Salazar, E.A., and León-Règagnon, V. Con�rmation of Gnathostoma binucleatum AlmeydaArtigas, 1991, advanced third-stage larvae in Tres Palos Lagoon, Mexico, by morphological and molecular data. J. Parasitol. 91, 962, 2005. 55. Bertoni-Ruiz, F. et al. A new species of Gnathostoma (Nematoda: Gnathostomatidae) in Procyon lotor hernandezii from Mexico. J. Parasitol. 91, 1143, 2005. 56. Hernández-Gómez, R.E. et al. Molecular identi�cation of the advanced third-stage larvae (ADV L3) of Gnathostoma lamothei in Tabasco, Mexico. Parasitol. Int. 59, 97, 2010. 57. Sieu, T.P.M. et al. Prevalence of Gnathostoma spinigerum infection in wild and cultured swamp eels in Vietnam. J. Parasitol. 98, 246, 2009. 58. Sambrook, J., Fritsch, E.F., and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory, Cold Spring Harbor, New York 1989. 55 Chapter 55 - Haemonchus
2. Hoberg, E.P., and Lichtenfels, J.R. Phylogenetic systematic analysis of the Trichostrongylidae (Nematoda), with an initial assessment of coevolution and biogeography. J. Parasitol. 80, 976, 1994.
3. Blaxter, M.L. Nematoda: Genes, genomes and the evolution of parasitism. Adv. Parasitol. 54, 101, 2003.
4. De Ley, P.B. Systematic position and phylogeny. In: Lee, D. (ed.), The Biology of Nematodes, Taylor & Francis, London, pp. 1–30, 2002.
5. Chan, M.S. The global burden of intestinal nematode infections—Fifty years on. Parasitol. Today 13, 438, 1997.
6. Gibbons, L.M.K. A key for the identi�cation of genera of the nematode family Trichostrongylidae Leiper, 1912. J. Helminthol. 56, 185, 1982.
7. Lichtenfels, J.R., and Pilitt, P.A. Synlophe patterns of the Haemonchinae of ruminants (Nematoda: Trichostrongyloidea). J. Parasitol. 86, 1093, 2000.
8. Ghadirian, E., and Arfaa, F. First report of human infection with Haemonchus contortus, Ostertagia ostertagi, and Marshallagia marshalli (family Trichostrongylidae) in Iran. J. Parasitol. 59, 1144, 1973.
9. Faust E.C., Russell P.F., and Rodney, C. J., Craig and Faust’s Clinical Parasitology, Lea & Febiger, Philadelphia, pp. 890, 1970.
10. Magalhães, P.S. Notes d’helminthologie bresilienne— L’Haemonchus contortus comme parasite accidentel de l’homme. Arch. Parasitol. 12, 283, 1908.
11. Fleury, G.C., Correa, M.O.A., and Neto, V.A. Identi�cação do Trichostrongylus colubriformis como parasita do homem. Rev. Inst. Meã. trop. São Paulo 12, 288, 1970.
12. Travassos, L., Ensaio monogra�co da familia Trichostrongilidae. Mem. Inst. Oswaldo Cruz 13, 1, 1921.
13. Sweet, W.C. The intestinal parasites of man in Australia and its dependencies as found by the Australian hookworm campaign. Med. J. Aust. 1, 353, 1924. 14. Heydon, G.M., and Green, A.K. Some worm infectations of man in Australia. Med. J. Aust. 1, 619, 1931.
15. Soulsby, E.J.L. Helminths, Arthropods and Protozoa of domesticated animals. Bailliere Tindall, London, 809pp., 1982.
16. Georgi, J.R., and McCulloch, C.E., Diagnositic morphometry: Identi�cation of helminth eggs by discriminant analysis of morphometric data. Proc. Helminthol. Soc. Washington 56, 44, 1989.
17. Veglia, F. The anatomy and life-history of Haemonchus contortus (Rud.). Third and Fourth Reports of the Director of Veterinary Research 1915, 347, 1916.
18. van Wyk, J.A., Cabaret, J., and Michael, L.M. Morphological identi�cation of nematode larvae of small ruminants and cattle simpli�ed. Vet. Parasitol. 119, 277, 2004.
19. Bowman, D.D. (ed.), In: Georgis’ Parasitology for Veterinarians, Saunders Elsevier, St. Louis, 464pp., 2009.
20. Gibbs, H.C. Hypobiosis in parasitic nematodes—An update. Adv. Parasitol. 25, 129, 1986.
21. Lindqvist, A. et al. The dynamics, prevalence and impact of nematode infections in organically raised sheep in Sweden, Acta Vet. Scand. 42, 377, 2001.
22. Boreham, R.E. et al. Human trichostrongyliasis in Queensland. Pathology 27, 182, 1995.
23. Crab, A. et al. The parasitic hematophagous worm Haemonchus contortus inhibits human platelet aggregation and adhesion: Partial puri�cation of a platelet inhibitor. Thromb. Haemost. 87, 899, 2002. 24. Konˇáková, I. Use of modi�ed McMaster method for the diagnosis of intestinal helminth infections and estimating parasitic egg load in human faecal samples. Helminthologia 46, 62, 2009. 25. Egwang, T.G., and Slocombe, J.O.D. Ef�ciency and sensitivity of techniques for recovering nematode eggs from bovine feces. Can. J. Comp. Med. 45, 243, 1981. 26. Cox, D.D., and Todd, A.C. Survey of gastrointestinal parasitism in Wisconsin dairy cattle. JAVMA 141, 706, 1962. 27. Goodman, D. et al. A comparison of methods for detecting the eggs of Ascaris, Trichuris, and hookworm in infant stool, and the epidemiology of infection in Zanzibari infants. Am. J. Trop. Med. Hyg. 76, 725, 2007. 28. Harmon, A.F. et al. Comparison of three different preservatives for morphological and real-time PCR analyses of Haemonchus contortus eggs. Vet. Parasitol. 145, 361, 2007. 29. Pawlowski, Z.S., and Stott, G.J. Hookworm Infection and Anemia: Approaches to Prevention and Control, Vol. 96. World Health Organization, Geneva, 1991. 30. Garcia, L.S. Practical Guide to Diagnostic Parasitology, ASM Press, Washington, DC, 448pp., 2009. 31. Hubert, J., and Kerboeuf, D. A new method for culture of larvae used in diagnosis of ruminant gastrointestinal strongylosis: Comparison with fecal cultures. Can. J. Comp. Med. 48, 63, 1984. 32. Gasser, R.B., Cantacessi, C., and Loukas, A. DNA technological progress toward advanced diagnostic tools to support human hookworm control. Biotechnol. Adv. 26, 35, 2008. 33. Gasser, R.B. et al. Toward practical, DNA-based diagnostic methods for parasitic nematodes of livestock—Bionomic and biotechnological implications. Biotechnol. Adv. 26, 325, 2008. 34. Mitreva, M. et al. Parasitic nematodes—From genomes to control. Vet. Parasitol. 148, 31, 2007. 35. Beh, K.J., Foley, R.C., and Goodwin, E.J. Restriction fragment length patterns of DNA from parasitic nematodes of sheep, Res. Vet. Sci. 46, 127, 1989. 36. Christensen, C.M., Zarlenga, D.S., and Gasbarre, L.C. Ostertagia, Haemonchus, Cooperia, and Oesophagostomum: Construction and characterization of genus-speci�c DNA probes to differentiate important parasites of cattle, Exp. Parasitol. 78, 93, 1994. 37. Mes, T.H., and Cornelissen, A.W. Non-homogenized ITS regions in the parasitic nematode Cooperia oncophora, Parasitology 129, 213, 2004. 38. Hillis, D.M., and Dixon, M.T. Ribosomal DNA: Molecular evolution and phylogenetic inference. Q. Rev. Biol. 66, 411, 1991. 39. Heise, M., Epe, C., and Schnieder, T. Differences in the second internal transcribed spacer (ITS-2) of eight species of gastrointestinal nematodes of ruminants. J. Parasitol. 85, 431, 1999. 40. Yong, T.S. et al. Differential diagnosis of Trichostrongylus and hookworm eggs via PCR using ITS-1 sequence, Korean J. Parasitol. 45, 69, 2007. 41. Garretson, P.D. et al., Anthelmintic resistant Haemonchus contortus in a giraffe (Giraffa camelopardalis) in Florida, J. Zoo Wildl. Med. 40, 131, 2009. 42. Traub, R.J. et al. PCR-based coprodiagnostic tools reveal dogs as reservoirs of zoonotic ancylostomiasis caused by Ancylostoma ceylanicum in temple communities in Bangkok. Vet. Parasitol. 155, 67, 2008.
43. Traub, R.J., Monis, P.T., and Robertson, I.D. Molecular epidemiology: A multidisciplinary approach to understanding parasitic zoonoses. Int. J. Parasitol. 35, 1295, 2005.
44. Zarlenga, D.S. et al. A multiplex PCR assay for differentiating economically important gastrointestinal nematodes of cattle. Vet. Parasitol. 97, 199, 2001.
45. von Samson-Himmelstjerna, G., Harder, A., and Schnieder, T. Quantitative analysis of ITS2 sequences in trichostrongyle parasites. Int. J. Parasitol. 32, 1529, 2002.
46. Verweij, J.J. et al. Simultaneous detection and quanti�cation of Ancylostoma duodenale, Necator americanus, and Oesophagostomum bifurcum in fecal samples using multiplex real-time PCR. Am. J. Trop. Med. Hyg. 77, 685, 2007.
47. de Gruijter, J.M. et al. Polymerase chain reaction-based differential diagnosis of Ancyclostoma duodenale and Nacator americanus infections in humans in northern Ghana. Trop. Med. Int. Health 10, 574, 2005.
48. Harmon, A.F., Zarlenga, D.S., and Hildreth, M.B., Improved methods for isolating DNA from Ostertagia ostertagi eggs in cattle feces. Vet. Parasitol. 135, 297, 2006.
49. Monteiro, L. et al. Complex polysaccharides as PCR inhibitors in feces: Helicobacter pylori model. J. Clin. Microbiol. 35, 995, 1997.
50. Thornton, C.G., and Passen, S. Inhibition of PCR ampli�cation by phytic acid, and treatment of bovine fecal specimens with phytase to reduce inhibition. J. Microbiol. Methods 59, 43, 2004.
51. Kreader, C.A. Relief of ampli�cation inhibition in PCR with bovine serum albumin or T4 gene 32 protein, Appl. Environ. Microbiol. 62, 1102, 1996.
52. Bott, N.J. et al. A combined microscopic-molecular method for the diagnosis of strongylid infections in sheep. Int. J. Parasitol. 39, 1277, 2009.
53. Quach, N., Goodman, M.F., and Shibata, D. In vitro mutation artifacts after formalin �xation and error prone translesion synthesis during PCR. BMC Clin. Pathol. 4, 1, 2004.
54. Gasser, R.B., Stewart, L.E., and Speare, R. Genetic markers in ribosomal DNA for hookworm identi�cation. Acta Trop. 62, 15, 1996.
55. Stevenson, L.A., Chilton, N.B., and Gasser, R.B. Differentiation of Haemonchus placei from H. contortus (Nematoda: Trichostrongylidae) by the ribosomal DNA second internal transcribed spacer. Int. J. Parasitol. 25, 483, 1995.
56. Harmon, A.F. et al. Real-time PCR for quantifying Haemonchus contortus eggs and potential limiting factors, Parasitol. Res. 101, 71, 2007. 57. Learmount, J. et al. Development and validation of real-time PCR methods for diagnosis of Teladorsagia circumcincta and Haemonchus contortus in sheep. Vet. Parasitol. 166, 268, 2009. 58. Wolstenholme, A.J. et al. Drug resistance in veterinary helminths. Trends Parasitol. 20, 469, 2004. 59. Coles, G.C. et al. The detection of anthelmintic resistance in nematodes of veterinary importance. Vet. Parasitol. 136, 167, 2006. 60. von Samson-Himmelstjerna, G. Molecular diagnosis of anthelmintic resistance. Vet. Parasitol. 136, 99, 2006. 61. Zheng, Q. et al. The control of hookworm infection in China. Parasit. Vectors 2, 44, 2009. 62. Keiser, J., and Utzinger, J. Ef�cacy of current drugs against soil-transmitted helminth infections: Systematic review and meta-analysis. JAMA. 299, 1937, 2008. 63. Palmer, D.G., and McCombe, I.L. Lectin staining of trichostrongylid nematode eggs of sheep: Rapid identi�cation of Haemonchus contortus eggs with peanut agglutinin. Int. J. Parasitol. 26, 447, 1996. 64. Jurasek, M.E. et al. Modi�cation and further evaluation of a �uorescein-labeled peanut agglutinin test for identi�cation of Haemonchus contortus eggs. Vet. Parasitol. 169, 209, 2010. 65. Gomez-Munoz, M.T., Cuquerella, M., and Alunda, J.M. Identi�cation and partial puri�cation of a 26 kilodalton antigen of adult Haemonchus contortus. Int. J. Parasitol. 26, 311, 1996. 66. Gomez-Munoz, M.T. et al. Serodiagnosis of haemonchosis with a somatic antigen (Hc26) in several breeds of sheep. J. Vet. Diagn. Invest. 12, 354, 2000. 67. Mahmoud, M.S., Abou Gamra, M.M., and Elkhayat, M.M. Ancylostoma duodenale infection: A study of serum immunoglobulin G4 response to the excretory secretory antigen of adult worm. J. Egypt Soc. Parasitol. 35, 1, 2005. 68. Palmer, D.R., Bradley, M., and Bundy, D.A. IgG4 responses to antigens of adult Necator americanus: Potential for use in large-scale epidemiological studies. Bull. World Health Organ. 74, 381, 1996. 69. Zarlenga, D.S. et al. Cloning and characterization of ribosomal RNA genes from three species of Haemonchus (Nematoda: Trichostrongyloidea) and identi�cation of PCR primers for rapid differentiation. Exp. Parasitol. 78, 28, 1994. 70. Chilton, N.B. et al. The evolutionary origins of nematodes within the order Strongylida are related to predilection sites within hosts. Mol. Phylogenet. Evol. 40, 118, 2006. 56 Chapter 56 - Loa
1. Gentillini, M., and Du�o, B. Filarioses. Médecine Tropicale, Flammarion, Paris, pp. 173–195, 1982.
2. World Health Organization. WHO Expert Commitee on Filariasis. Fourth Report, WHO Technical Series, 702, 1984.
3. World Health Organization. Lymphatic Filariasis: The Disease and Its Control. Fifth Report of the WHO Expert Commitee on Filariasis. WHO Technical Report Series, 821, 1992.
4. Muller, R. Worms and Disease. Heinemann Medical Books, London, UK, 1975.
5. Mongin. Observations sur un ver trouvé dans la conjonctive à maribou. île Saint Domingue. J. Med. Chir. Pharm. etc (Paris) 32, 338–339, 1770.
6. Guyon, J. Sur un nouveau cas de �laire, sous conjonctive, ou Filaria occuli des auteurs, observe au Gabon (Côte occidentale d’’frique). C. R. Séances Acad. Sci. 59, 743–748, 1964.
7. Brumpt, E. Les �larioses humaines en Afrique. C. R. Soc. Biol. 56, 758–760, 1904.
8. Connor, D.H., Nea�e, R.C., and Meyers, W.M. Loiasis: Pathology of Tropical and Extra-Ordinary Diseases. Armed Forces Institute of Pathology, Washington, pp. 356–359, 1976.
30. Hulin, C. et al. Atteinte pulmonaire d’évolution favorable au cours d’une �lariose de type Loa loa. Bull. Soc. Path. Exp. 87, 248–250, 1994.
31. Klion, A.D. et al. Loiasis in endemic and non endemic populations: Immunologically mediated differences in clinical presentation. J. Infect. Dis. 163, 1318–1325, 1991b.
32. Nutman, T.B. et al. Immunologic correlates of the hyperresponsive syndrome of loiasis. J. Infect. Dis. 157, 544– 550, 1988.
33. Boulestiex, G., and Carme, B. Encephalite au cours du traitement de la �lariose à loase par diethylcarbamazine. A propos de 6 observations. Bull. Soc. Pathol. Exot. 79, 649– 654, 1986.
34. Egwang, T.G. et al. The indenti�cation and partial characterization of an immunodominant 29–31 kilodalton surface antigen expressed by adult worms of the human �laria Loa loa. Mol. Biochem. Parasitol. 31, 263–277, 1988.
35. Pinder, M., Dupont, A., and Egwang, T.G. Identi�cation of a surface antigen on Loa loa micro�lariae the recognition of which correlates with the amicro�laremic state in man. J. Immunol. 141, 2480–2486, 1988.
36. Akué, J.P., Egwang, T.G., and Devaney, E. High levels of parasite- speci�c IgG4 in the absence of micro�laremia in Loa loa infection. Trop. Med. Parasitol. 45, 246–248, 1994.
37. Akué, J.P., Hommel, M., and Devaney, E. High levels of parasite- speci�c IgG1 correlate with the amicro�laremic state in Loa loa infection. J. Infect. Dis. 175, 158–163, 1997.
38. Pinder, M., Leclaire, A., and Everaere, S. Antibody-dependent cell-mediated immune reactions to Loa loa micro�lariae in amicro�laremic subjects. Parasite Immunol. 14, 541–556, 1992.
39. Baize, S. et al. T helper responsiveness in human Loa loa infection: Defective speci�c proliferation and cytokine production by CD4+ T cells from micro�laremic subjects compared with amicro�laremics. Clin. Exp. Immunol. 108, 272–278, 1997.
40. Leroy, E. et al. Experimental infection of a nonhuman primate with Loa loa induces transient strong immune activation followed by peripheral unresponsiveness of helper T cells. Infect. Immun. 65, 1876–1882, 1997.
41. Goussard, B. et al. Age of appearance of IgG, IgGM and IgE antibodies speci�c for Loa loa in Gabonese children. Microbiol. Immunol. 28, 787–792, 1984.
42. Van Hoegaerden, M. et al. Filariasis due to Loa loa and Mansonella perstans: Distribution in the region of Okondja, Haut Ogooué Province, Gabon, with parasitological and serological follow-up over one year. Trans. R. Soc. Trop. Med. Hyg. 81, 441–446, 1987.
43. Lal, R.B., and Ottesen, E.A. Enhanced diagnostic speci�city in human �lariasis by IgG4 antibody assessment. J. Infect. Dis. 158,1034, 1983.
44. Lal, R.B. et al. Lack of IgG4 antibody response to carbohydrate antigens in patients with lymphatic �lariasis. Immunology 74, 333–337, 1991.
45. Ottesen, E.A. et al. Prominence of IgG4 in the IgG antibody response to human �lariasis. J. Immunol. 134, 2707–2712, 1985.
46. Lucius, R. et al. Speci�c and sensitive IgG4 immunodiagnosis of onchocerciasis with a recombinant 33Kd Onchocerca volvulus protein (OV33). Trop. Med. Parasitol. 43, 139–145, 1992. 47. Weil, G.J. et al. IgG4 subclass antibody serology for onchocerciasis. J. Infect. Dis. 161, 549–554, 1990. 48. Akué, J.P. et al. Analysis of speci�c IgG subclasses in a population naturally exposed to Loa loa. Am. J. Trop. Med. Hyg. 49(suppl), 209, 1993. 49. Garraud, O., and Nutman, T.B. The role of cytokines in human B-cell differentiation into immunoglobulin-secreting cells. Bull. Inst. Pasteur 94, 285–309, 1996. 50. Dimock, K.A., Eberhard, M.L., and Lamma, P.J. Th1-Like anti�larial immune responses predominate in antigen-negative persons. Infect. Immun. 64, 2962–2967, 1996. 51. Eveland, L.K., Yermakov, V., and Kennedy, M. Loa loa infection without micro�lariae. Trans. R. Soc. Trop. Med. Hyg. 69, 354–355, 1975. 52. Dupont, A., Zué-Ndong, J., and Pinder, M. Common occurence of amicro�laremic Loa loa �lariasis within the endemic region. Trans. R. Soc. Trop. Med. Hyg. 82, 730, 1988. 53. Touré, F.S. et al. IgG4 serology of loiasis in three endemic villages in south-eastern Gabon. Trop. Med. Int. Health 3, 313–317, 1998. 54. Richard-Lenoble, D. et al. Prévalence des �larioses humaines sanguicoles au Gabon. Bull. Soc. Pathol. Exot. 73, 192–199, 1980. 55. Languillat, G. et al. Enquête sur l’étiologie de l’hypofécondité au Gabon Oriental: Endémie �larienne (Loa loa et D. perstans). Fréquences des micro�laires dans les hydrocèles. Rev. Epidemiol. Santé Publique 26, 273–283, 1978. 56. Burbelo, P.D. et al. Rapid antibody quanti�cation and generation of whole proteome antibody response pro�les using LIPS (luciferase immunoprecipitation systems). Biochem. Biophys. Res. Commun. 352, 889–895, 2007. 57. McReynolds, L.A., DeSimone, S.M., and Williams, S.A. Cloning and comparison of repeated DNA sequences from the human �larial parasite Brugia malayi and the animal parasite Brugia pahangi. Proc. Natl Acad. Sci. USA 83, 797–801, 1986. 58. Williams, S.A., DeSimone, S.M., and McReynolds, L.A. Species-speci�c oligonucleotide probes for the identi�cation of human �larial parasites. Mol. Biochem. Parasitol. 28, 163– 169, 1988. 59. Williams, S.A. et al. A polymerase chain reaction assay for the detection of Wuchereria bancrofti in blood samples from French Polynesia. Trans. R. Soc. Trop. Med. Hyg. 90, 384–387, 1996. 60. Erttmann, K.D. et al. A DNA sequence specific for forest form Onchocerca volvulus. Nature 327, 415–417, 1987. 61. Klion, A.D. et al. Cloning and characterization of a species-speci�c repetitive DNA sequence from Loa loa. Mol. Biochem. Parasitol. 45, 297–305, 1991. Erratum in: Mol. Biochem. Parasitol. 47, 265. 62. Egwang, T.G., Ajuh, P.M., and Akue, J.P. Cloning and characterization of a Loa loa-speci�c repetitive DNA. Mol. Biochem. Parasitol. 56, 189–196, 1992. 63. McReynolds, L.A., Kennedy, M.W., and Selkirk, M.E. The polyprotein allergens of nematodes. Parasitol. Today 9, 403– 406, 1993. 64. Ajuh, P.M. et al. Loa loa: Structural diversity of a 15-kDa repetitive antigen. Exp. Parasitol. 81, 145–153, 1995. 65. Touré, F.S. et al. Species-speci�c sequence in the repeat 3 region of the gene encoding a putative Loa loa allergen: A diagnostic tool for occult loiasis. Am. J. Trop. Med. Hyg. 56, 57–60, 1997a.
66. Touré, F.S. et al. Detection of Loa loa-speci�c DNA in blood from occult-infected individuals. Exp. Parasitol. 86, 163–170, 1997b.
67. Lizotte, M.R. et al. A polymerase chain reaction assay for the detection of Brugia malayi in blood. Am. J. Trop. Med. Hyg. 51, 314–321, 1994.
68. Zimmermann, P.A. et al. Polymerase chain reaction-based diagnosis of Onchocerca volvulus infection: Improved detection of patients with Onchocerciasis. J. Infect. Dis. 169, 686– 689, 1994.
69. Touré, F.S. et al. Human occult loiasis: Field evaluation of a nested polymerase chain reaction assay for the detection of occult infection. Trop. Med. Int. Health 3, 505–511, 1998. 70. Touré, F.S. et al. Relationship between the intensity of Loa loa �lariasis transmission and prevalence of infections. Med. Trop. 59, 249–252, 1998. [In French]. 71. Pinder, M., Everaere, S., and Roelants, G.E. Loa loa: Immunological responses during experimental infections in mandrills (Mandrillus sphinx). Exp. Parasitol. 79, 126–136, 1994. 72. Touré, F.S. et al. Use of polymerase chain reaction for accurate follow-up of Loa loa experimental infection in Mandrillus sphinx. Am. J. Trop. Med. Hyg. 61, 956–959, 1999. 57 Chapter 57 - Mansonella
19. Godoy, G.A. et al. Mansonella ozzardi infections in Indians of the Southwestern part of the state of Bolivar, Venezuela. Am. J. Trop. Med. Hyg. 29, 373, 1980.
20. Marinkelle, C.J., and German, E. Mansonelliasis in the Comisaria del Vaupes of Colombia. Trop. Geogr. Med. 22, 101, 1970.
21. Fischer, P., Bamuhiiga, J., and Buttner, D.W. Occurrence and diagnosis of Mansonella streptocerca in Uganda. Acta Trop. 63, 43, 1997.
22. Bartoloni, A. et al. Mansonella ozzardi infection in Bolivia: Prevalence and clinical associations in the Chaco region. Am. J. Trop. Med. Hyg. 61, 830, 1999.
23. Orihel, T.C., Eberhard, M.L., and Lowrie, R.C. Jr. Mansonella ozzardi: The course of patency in experimentally-infected patas monkeys. Trop. Med. Parasitol. 44, 49, 1993.
24. Sasa, M. Human �lariasis: A Global Survey of Epidemiology and Control. University Park Press, Baltimore, 1976.
25. Duke, B.O. A case of streptocerciasis in a European. Ann. Trop. Med. Parasitol. 51, 364, 1957.
26. Meyers, W.M. et al. Human streptocerciasis. A clinico- pathologic study of 40 Africans (Zairians) including identi�cation of the adult �laria. Am. J. Trop. Med. Hyg. 21, 528, 1972.
27. Nutman, T.B., Nash, T.E., and Ottesen, E.A. Ivermectin in the successful treatment of a patient with Mansonella ozzardi infection. J. Infect. Dis. 156, 662, 1987.
28. Branco, B.C. et al. Achados oculares entre habitantes do municipio de Pauini e possível associação entre lesões corneanas e mansonelose na Amazônia. Arq. Bras. Oftalmol. 61, 674, 1998.
29. Cohen, J.M., Ribeiro, J.A., and Martins, M. Ocular manifestations in mansonelliasis. Arq. Bras. Oftalmol. 71, 167, 2008.
30. Garrido, C., and Campos, M. First report of presumed parasitic keratitis in Indians from the Brazilian Amazon. Cornea 19, 817, 2000.
31. Bregani, E.R. et al. Case report: Intraocular localization of Mansonella perstans in a patient from south Chad. Trans. R. Soc. Trop. Med. Hyg. 96, 654, 2002.
32. Sasa, M. Micro�laria survey methods and analysis of survey data in �lariasis control programmes. Bull. World Health Organ. 37, 629, 1967.
33. WHO. Bench Aids for the Diagnosis of Filarial Infections. WHO, Geneva, 1997.
34. Medeiros, J.F. et al. Current pro�le of Mansonella ozzardi (Nematoda: Onchocercidae) in communities along the Ituxi river, Labrea municipality, Amazonas, Brazil. Mem. Inst. Oswaldo Cruz 103, 409, 2008.
35. Post, R.J. et al. The morphological discrimination of micro�lariae of Onchocerca volvulus from Mansonella ozzardi. Parasitology 127, 21, 2003.
36. Wanji, S. et al. Epidemiology of concomitant infections due to Loa loa, Mansonella perstans, and Onchocerca volvulus in rain forest villages of Cameroon. Med. Microbiol. Immunol. 192, 15, 2003. 37. Crewe, W. Mansonelliasis. In: Service, M.W. (ed), The Encyclopedia of Arthropod-Transmitted Infections. CABI Publishing, Wallingford, p. 327, 2001. 38. Fischer, P. et al. Detection of the �larial parasite Mansonella streptocerca in skin biopsies by a nested polymerase chain reaction-based assay. Am. J. Trop. Med. Hyg. 58, 816, 1998. 39. Xie, H., Bain, O., and Williams, S.A. Molecular phylogenetic studies on �larial parasites based on 5S ribosomal spacer sequences. Parasite 1, 141, 1994. 40. Hojas, R.M., and Post, R.J. Regional genetic variation in the major sperm protein genes of Onchocerca volvulus and Mansonella ozzardi (Nematoda: Filarioidea). Int. J. Parasitol. 30, 1459, 2000. 41. Scott, A.L. et al. Major sperm protein and actin genes in free-living and parasitic nematodes. Parasitology 98, 471, 1989. 42. Setterquist, R.A. et al. Diagnostic probes targeting the major sperm protein gene that may be useful in the molecular identi�cation of nematodes. J. Nematol. 28, 414, 1996. 43. Morales-Hojas, R. et al. Characterisation of nuclear ribosomal DNA sequences from Onchocerca volvulus and Mansonella ozzardi (Nematoda: Filarioidea) and development of a PCRbased method for their detection in skin biopsies. Int. J. Parasitol. 31, 169, 2001. 44. Newton, L.A. et al. Systematic relationships of some members of the genera Oesophagostomum and Chabertia (Nematoda: Chabertiidae) based on ribosomal DNA sequence data. Int. J. Parasitol. 28, 1781, 1998. 45. Tang, T.H. et al. Nested PCR to detect and distinguish the sympatric �larial species Onchocerca volvulus, Mansonella ozzardi and Mansonella perstans in the Amazon region. Mem. Inst. Oswaldo Cruz 105, 823, 2010. 46. Moraes, M.A. et al. The concentration of Mansonella ozzardi micro�lariae in skin capillaries. Trans. R. Soc. Trop. Med. Hyg. 77, 463, 1983. 47. Moraes, M.A.P. Mansonella ozzardi micro�lariae in skin snips. Trans. R. Soc. Trop. Med. Hyg. 70, 16, 1976. 48. Ewert, A., Smith, J.H., and Corredor, A. Micro�lariae of Mansonella ozzardi in human skin biopsies. Am. J. Trop. Med. Hyg. 30, 988, 1981. 49. Fuglsang, H., and Anderson, J. Micro�lariae of Onchocerca volvulus in blood and urine before, during, and after treatment with diethylcarbamazine. J. Helminthol. 48, 93, 1974. 50. Anderson, R.I., Fazen, L.E., and Buck, A.A. Onchocerciasis in Guatemala. II. Micro�lariae in urine, blood, and sputum after diethylcarbamazine. Am. J. Trop. Med. Hyg. 24, 58, 1975. 51. Duke, B.O., Moore, P.J., and Vincelette, J. Factors in�uencing the passage of Onchocerca volvulus micro�lariae into the urine. Tropenmed. Parasitol. 26, 449, 1975. 52. Morales-Hojas, R. Molecular systematics of �larial parasites, with an emphasis on groups of medical and veterinary importance, and its relevance for epidemiology. Infect. Genet. Evol. 9, 748, 2009. 58 Chapter 58 - Marshallagia
TABLE 58.2
Molecular Markers for M. marshalli/M. occidentalis
Marker Product Length (bp) GenBank TM Accession Number Author/s and/or Reference
Marshallagia marshalli
ITS-1 384 AY013242 [76]
ITS-2 274 AJ577469 Leignel and Cabaret a
26S rRNA 50 AJ400715 [79]
5.8S rRNA 133 AY013242 [76]
Marshallagia occidentalis
ITS-1 384 AY013243 [76]
ITS-2 235 AY013244 [76]
26S rRNA 50 AY013244 [76]
5.8S rRNA 133 AY013243 [76] a Unpublished data.
16. De Ley, P., and Blaxter, M. Systematic position and phylogeny. In: Lee, D.L. (ed.), The Biology of Nematodes. Chapter 1. Taylor & Francis, London and New York, 2002.
17. Drózdz, J. Polymorphism in the Ostertagiinae Lopez-Neyra, 1947 and comments on the systematics of these nematodes. Syst. Parasitol. 32, 91, 1995.
18. Suarez, V.H., and Cabaret, J. Similarities between species of the Ostertagiinae (Nematoda: Trichostrongyloidea) in relation to host-speci�city and climatic environment. Syst. Parasitol. 20, 179, 1991.
19. Halvorsen, O., and Bye, K. Parasites, biodiversity, and population dynamics in an ecosystem in the high arctic, Vet. Parasitol. 84, 205, 1999. 20. Poulin, R. Evolutionary Ecology of Parasites: From Individuals to Communities. Chapman and Hall, London, 1998.
21. Zaffaroni, E. et al. Host speci�city of abomasal nematodes in free ranging alpine ruminants. Vet. Parasitol. 90, 221, 2000.
22. Bergstrom, R.C. Prevalence of Marshallagia marshalli Orloff, 1933 in wild ruminants in Wyoming. Proc. Oklahoma Acad. Sci. 55, 101, 1975.
23. Kistner, T.P. et al. Helminth parasites of bighorn sheep in Oregon, J. Wildl. Dis. 13, 125, 1977.
24. Nielsen, C.A., and Neiland, K.A. Sheep Disease Report, vol. 14, Alaska Department of Fish and Game, Juneau, 1974.
25. Lichtenfels, J.R., and Pilitt, P.A. Cuticular ridge patterns of Marshallagia marshalli and Ostertagia occidentalis (Nematoda: Trichostrongyloidea) parasitic in ruminants of North America. Proc. Helminthol. Soc. Wash. 56, 173, 1989.
26. Kerr, G.R., and Holmes, J.C. Parasites of mountain goats in west-central Alberta. J. Wildl. Manag. 30, 786, 1966.
27. Jacquiet, P. et al. Dry areas: An example of seasonal evolution of helminth infection on sheep and goats in southern Mauritania. Vet. Parasitol. 56, 137, 1995.
28. Valcárcel, F., and García-Romero, C. Prevalence and seasonal pattern of caprinae trichostrongyles in a dry area of central Spain. J. Vet. Med. B 46, 673, 1999.
29. Garijo, M.M., Ortiz, J.M., and De Ibáñez, M.R.R. Helminths in a giraffe (Giraffa camelopardalis giraffa) from a zoo in Spain. Onderst. J. Vet. Res. 71, 153, 2004.
30. Taylor, M.A. et al. Veterinary Parasitology. 3rd ed. Blackwell Publishing, Oxford, 2007.
31. Skrjabin, K.I., Shikhobalova, N.P., and Shultz, R.S. Essentials of Nematology. Vol. IV. Trichostrongylids of Animals and Man. Academy of Sciences of the URSS, Moscow, 1954. [Translated by the Israel Program for Scienti�c Translations, 1960].
32. Rickard-Ballweber, L. Ecto- and endoparasites of new world Camelids. Vet. Clin. North Am. Food Anim. Prac. 25, 2959, 2009.
33. Bergstrom, R.C. Incidence of Marshallagia marshalli Orloff, 1933 in Wyoming sheep, Ovis aries, and pronghorn antelope. Antilocapra Americana, Proc. Helminthol. Soc. Wash. 42, 61, 1975.
34. Manfredi, M.T. et al. Abomasal nematodes of the red deer Cervus elaphus in north-eastern Italy. J. Helminthol. 81, 247, 2007.
35. Rossi, L., Eckel, B., and Ferroglio, E. A survey of the gastrointestinal nematodes of roe deer (Capreolus capreolus) in a mountain habitat. Parasitologia 39, 303, 1997.
36. Morgan, E.R. et al. Helminths of saiga antelope in Kazakhstan: Implications for conservation and livestock production. J. Wildl. Dis. 41, 149, 2005. 37. Webster, W.A., and Rowell, J. Some helminths parasites from the small intestine of free-ranging muskoxen Ovibos moschatus (Zimmermann) of Devon and Ellsemere Islands, Northwest Territories. Can. J. Zool. 58, 304, 1980. 38. Pisanu, B. et al. Inventaire préliminaire des helminthes gastrointestinaux du mou�on de Corse (Ovis amon musimon) de Lile Haute, Archipel de Kerguelen. Gibier Faun. Sauv. 11, 327, 1994. 39. Rossi, L. et al. La simpatria mu�one/camoscio modi�ca sensibilmente le rispettive comunità elmintiche? Suppl. Ric. Biol. Selvaggina 24, 79, 1996. 40. Eslami, A. et al. Gastrointestinal nematodes of wild sheep (Ovis orientalis) from Iran. J. Wildl. Dis. 15, 263, 1979. 41. Rossi, L. et al. Elmintofauna del camoscio (Rupicapra rupicapra) nel Parco Naturale Argentera. Ann. Fac. Med. Vet. Torino. 33, 1 (1988–1989). 42. Cancrini, G. et al. Occurrence of pulmonary and gastrointestinal nematodes in the Abruzzo chamois. In: Lovari, S. (ed.), The Biology and Management of Mountain Ungulates. Chapter 29. Croom Helm, London, 1985. 43. Lanfranchi, P. et al. A three-year evaluation of abomasal helminths of Piz Albris ibex colony. Z. Jagdwiss. 41, 24 (1995) [in German]. 44. Pérez, J.M. et al. A survey of the gastrointestinal nematodes of Spanish ibex (Capra pyrenaica) in a high mountain habitat. J. Parasitol. 89, 315, 2003. 45. Gibbons, L.M., and Khalil, L.F. A key for the identi�cation of genera of the nematode family Trichostrongylidae Leiper, 1912. J. Helminthol. 56, 185, 1982. 46. Durette-Desset, M.C. Keys to genera of the superfamily Trichostrongyloidea. In:. Anderson, R.C., and Chabaud, A.G. (eds), CIH Keys to the Nematode Parasites of Vertebrates. CAB, Farnham Royal, 1983. 47. Irvine, R.J. et al. Life-history strategies and population dynamics of abomasal nematodes in Svalbard reindeer (Rangifer tarandus platyrhynchus). Parasitology 120, 297, 2000. 48. Lee, D.L. Life cycles. In: Lee, D.L. (ed.), The Biology of Nematodes. Chapter 3. Taylor & Francis, London, 2002. 49. Smith, G., and Grenfell, B.T. The population biology of Ostertagia ostertagi. Parasitol. Today 1, 76, 1985. 50. Irvine, R.J. et al. Contrasting regulation of fecundity in two abomasal nematodes of Svalbard reindeer (Rangifer tarandus platyrhynchus). Parasitology 122, 673, 2001. 51. Côté, S.D. et al. Resistance to abomasal nematodes and individual genetic variability in reindeer. Mol. Ecol. 14, 4159, 2005. 52. Decristophoris, P.M.A., von Hardenberg, A., and McElligott, A.G., Testosterone is positively related to the output of nematode eggs in male Alpine ibex (Capra ibex) faeces. Evol. Ecol. Res. 9, 1, 2007. 53. Ezenwa, V.O. Interactions among host diet, nutritional status and gastrointestinal parasite infection in wild bovids. Int. J. Parasitol. 34, 535, 2004. 54. Ghadirian, E., and Arfaa, F. First report of human infection with Haemonchus contortus, Ostertagia ostertagi, and Marshallagia marshalli (Family Trichostrongylidae) in Iran. J. Parasitol. 59, 1144, 1973. 55. Faust, E.C., Russell, P.F., and Jung, R.C. Craig and Faust’s Clinical Parasitology. Lea & Febiger, Philadelphia, 1970. 56. Stear, M.J. et al. The processes in�uencing the distribution of parasitic nematodes among naturally infected lambs. Parasitology 117, 165, 1998. 57. Sykes, A.R. Parasitism and production in farm-animals. Anim. Prod. 59, 155, 1994.
58. Al-Khalidi, N.W. et al. Pathogenicity of Marshallagia marshalli (Ransom) in sheep, Ind. J. Anim. Sci. 59, 917, 1989.
59. MAFF. Manual of Veterinary Parasitology Laboratory Techniques. HMSO, London, 1986.
60. Eysker, M., and Kooyman, F.N.J. Notes on necropsy and herbage processing techniques for gastrointestinal nematodes of ruminants. Vet. Parasitol. 46, 205–213, 1993.
61. Gaba, S. et al. Estimation of abomasum strongyle nematode infections in sheep at necropsy: Tentative proposals for a simpli�ed technique. Vet. Parasitol. 140, 105, 2006.
62. Ash, L.R., and Orihel, T.C. Parasites: A Guide to Laboratory Procedures and Identi�cation. ASCP Press, Chicago, 1987.
63. Thienpont, D., Rochette, F., and Vanparijs, O.F.J. Diagnosing Helminthiasis by Coprological Examination. Jansen Research Foundation, Beerse, 1986.
64. Zarlenga, D.S. et al. A multiplex PCR assay for differentiating economically important gastrointestinal nematodes of cattle. Vet. Parasitol. 97, 199, 2001.
65. Harmon, A.F., Zarlenga, D.S., and Hildreth, M.B. Improved methods for isolating DNA from Ostertagia ostertagi eggs in cattle feces. Vet. Parasitol. 135, 297, 2006.
66. Callaghan, M.J., and Beh, K.J. A tandemly repetitive DNA sequence is present at diverse locations in the genome of Ostertagia circumcincta. Gene 174, 273, 1996.
67. Zarlenga, D.S. et al. Identi�cation and semi-quanti�cation of Ostertagia ostertagi eggs by enzymatic ampli�cation of ITS 1 sequences. Vet. Parasitol. 77, 245, 1998.
68. Otsen, M. et al. Microsatellite diversity of isolates of the parasitic nematode Haemonchus contortus. Mol. Biochem. Parasitol. 110, 69, 2000.
69. Prichard, R. Genetic variability following selection of Haemonchus contortus with antihelmintics. Trends Parasitol. 17, 445, 2001.
70. Grillo, V., Jackson, F., and Gilleard, J.S. Characterisation of Teladorsagia circumcincta microsatellites and their development as population genetic markers. Mol. Biochem. Parasitol. 148, 181, 2006. 71. Álvarez-Sánchez, M.A. et al. Real time PCR for the diagnosis of benzimidazole resistance in trichostrongylids of sheep. Vet. Parasitol. 129, 291, 2005. 72. Wimmer, B. et al. Non-invasive assessment of parasitic nematode species diversity in wild Soay sheep using molecular markers. Int. J. Parasitol. 34, 625, 2004. 73. von Samson-Himmelstjerna, G., Harder, A., and Schnieder, T. Quantitative analysis of ITS2 sequences in trichostrongyle parasites. Int. J. Parasitol. 32, 1529, 2002. 74. Stevenson, L.A., Gasser, R.B., and Chilton, N.B. The ITS-2 rDNA of Teladorsagia circumcincta, T. trifurcata and T. davtiani (Nematoda: Trichostrongylidae) indicates that these taxa are one species. Int. J. Parasitol. 26, 1123, 1996. 75. Dallas, J.F., Irvine, R.J., and Halvorsen, O. DNA evidence that Ostertagia gruehneri and Ostertagia arctica (Nematoda: Ostertagiinae) in reindeer from Norway and Svalbard are conspeci�c. Int. J. Parasitol. 30, 655, 2000. 76. Dallas, J.F., Irvine, R.J., and Halvorsen, O. DNA evidence that Marshallagia marshalli Ransom, 1907 and M. occidentalis Ransom, 1907 (Nematoda: Ostertagiinae) from Svalbard reindeer are cospeci�c. Syst. Parasitol. 50, 101, 2001. 77. Hoste, H. et al. A comparison of the �rst internal transcribed spacer of ribosomal DNA in seven species of Trichostrongylus (Nematoda: Trichostrongylidae). Int. J. Parasitol. 28, 1251, 1998. 78. Hoste, H. et al. Differences in the second internal transcribed spacer (ribosomal DNA) between �ve species of Trichostrongylus (Nematoda: Trichostrongylidae). Int. J. Parasitol. 25, 75, 1995. 79. Dallas, J.F. et al. Identi�cation by polymerase chain reaction (PCR) of Marshallagia marshalli and Ostertagia guehneri from Svalbard reindeer. Int. J. Parasitol. 30, 863, 2000. 80. Sambrook, J., MacCallum, P., and Russell, D. Molecular Cloning: A Laboratory Manual. 3rd ed. Cold Spring Harbor Laboratory Press, New York, 2001. 81. Brooks, D.R., and Hoberg, E.P. Triage for the biosphere: The need and rationale for taxonomic inventories and phylogenetic studies of parasites. Comp. Parasitol. 67, 1, 2000. 59 Chapter 59 - Necator
1. Lichtenfels, J.R. Keys to the Genera of the Superfamilies Ancylostomatoidea and Diaphanocephaliodea. Farnham Royal, Commonwealth Agricultural Bureaux, 1980.
2. Bethony, J. et al. Soil-transmitted helminth infections: Ascariasis, trichuriasis, and hookworm. Lancet 367, 1521, 2006. 3. John, D.T., and Petri, W.A., Jr. Markell and Voge’s Medical Parasitology, 9th edn. Saunders Elsevier, St. Louis, 2006. 4. Hotez, P.J. et al. Hookworm: The great infection of mankind. PLoS Med. 2, e67, 2005. 5. CDC PDx-Lab Assistance: Hookworm Infection. www.dpd. cdc.gov/dpdx/HTML/Hookworm.htm 6. Hotez, P.J. et al. Hookworm infection. N. Engl. J. Med. 351, 799, 2004. 7. de Silva, N.R. Impact of mass chemotherapy for the control of �lariasis on geohelminth infections in Sri Lanka. Ann. Trop. Med. Parasitol. 97, 421, 2003. 8. Liu, C.H. et al. Epidemiology of human hookworm infections among adult villagers in Hejiang and Santai Counties, Sichuan Province, China. Acta Trop. 73, 243, 1999. 9. Olsen, A. et al. Cross-sectional parasitological survey for helminth infections among �sh farmers in Nghe An province, Vietnam. Acta Trop. 100, 199, 2006. 10. Bethony, J. et al. Emerging patterns of hookworm infections: In�uence of aging on the intensity of Necator infection in Hainan Province, People’s Republic of China. Clin. Inf. Dis. 35, 1336, 2002. 11. Hotez, P.J. et al. Current concepts: Hookworm infection. N. Engl. J. Med. 351, 799, 2004. 12. Brooker, S. et al. Epidemiology of Plasmodium-helminth co-infection in Africa: Populations at risk, potential impact on anemia and prospects for combining control. Am. J. Trop. Med. Hyg. 77(6 suppl.), 88, 2007. 13. McDevitt, M.A. et al. The anemia of malaria infection: Role of in�ammatory cytokines. Curr. Hematol. Rep. 3, 97, 2004. 14. Williamson, A.L., Brindley, P.J., and Loukas, A. Hookworm cathepsin D aspartic proteases: Contributing roles in the host-speci�c degradation of serum proteins and skin macromolecules. Parasitology 126, 179, 2003. 15. Stoltzfus, R.J. et al. Malaria, hookworms and recent fever are related to anemia and iron status indicators in 0- to 5-y old Zanzibari children and these relationships change with age. J. Nutr. 130, 1724, 2000. 16. Bundy, D.A., Chan, M.S., and Savioli, L. Hookworm infection in pregnancy. Trans. R. Soc. Trop. Med. Hyg. 89, 521, 1995. 17. Sakti, H. et al. Evidence for an association between hookworm infection and cognitive function in Indonesian school children. Trop. Med. Int. Health 4, 322, 1999. 18. Christian, P., Khatry, S.K., and West, J.P., Jr. Antenatal anthelminthic treatment, birthweight, and infant survival in rural Nepal. Lancet 364, 981, 2004. 19. Brooker, S., Bethony, J., and Hotez, P.J. Hookworm infection in the 21st century. Adv. Parasitol. 58, 197, 2004. 20. Albonico, M. et al. Controlling soil-transmitted helminthiasis in pre-school-age children through preventive chemotherapy. PLoS Negl. Trop. Dis. 2, e126, 2008. 21. Albonico, M. et al. Intervention for the control of soiltransmitted helminthiasis in the community. Adv. Parasitol. 61, 311, 2006. 22. De Rochars, M.B. et al. Community-wide reduction in prevalence and intensity of intestinal helminths as a collateral bene�t of lymphatic �lariasis elimination programs. Am. J. Trop. Med. Hyg. 71, 466, 2004. 23. Fox, L.M. et al. Ultrasonographic examination of Haitian children with lymphatic �lariasis: A longitudinal assessment in the context of anti�larial drug treatment. Am. J. Trop. Med. Hyg. 72, 642, 2005. 24. Kaplan, R.M. Drug resistance in nematodes of veterinary importance: A status report. Trends Parasitol. 20, 477, 2004.
25. Prichard, R.K. et al. The problem of anthelmintic resistance in nematodes. Aust. Vet. J. 56, 239, 1980.
26. Silvestre, A., and Cabaret, J. Mutation in position 167 of isotype 1 β-tubulin gene of Trichostrongylid nematodes: Role in benzimidazole resistance? Mol. Biochem. Parasitol. 120, 297, 2002.
27. Hodgkinson, J.E. et al. The role of polymorphisms at β-tubulin isotype 1 codons 167 and 200 in benzimidazole resistance in cyathostomins. Int. J. Parasitol. 38, 1149, 2008.
28. Kwa, M.S., Veenstra, J.G., and Roos, M.H. Benzimidazole resistance in Haemonchus contoruts is correlated with a conserved mutation at amino acid 200 in β-tubulin isotype 1. Mol. Biochem. Parasitol. 63, 299, 1994.
29. Ghisi, M., Kaminsky, R., and Maser, P. Phenotyping and genotyping of Haemonchus contortus isolates reveals a new putative candidate mutation for benzimidazole resistance in nematodes. Vet. Parasitol. 144, 313, 2007.
30. Albonico, M. et al. Development of the egg hatch assay for detection of anthelminthic resistance in human hookworms. Int. J. Parasitol. 35, 803, 2005.
31. Kotze, A.C. et al. Field evaluation of anthelmintic drug sensitivity using in vitro egg hatch and larval motility assays with Necator americanus recovered from human clinical isolates. Int. J. Parasitol. 35, 445, 2005.
32. Kotze, A.C. An in vitro larval motility assay to determine anthelmintic sensitivity for human hookworm and Strongyloides species. Am. J. Trop. Med. Hyg. 71, 608, 2004.
33 De Clercq, D. et al. Failure of mebendazole in treatment of human hookworm infections in the southern region of Mali. Am. J. Trop. Med. Hyg. 57, 25, 1997.
34. Geerts S., and Gryseels B. Drug resistance in human helminths: Current situation and lessons from livestock. Clin. Microbiol. Rev. 13, 207, 2000.
35. Albonico, M., Wright, V., and Bickle, Q. Molecular analysis of the β-tubulin gene of human hookworms as a basis for possible benzimidazole resistance on Pemba Island. Mol. Biochem. Parasitol. 134, 281, 2004.
36. Schwenkenbecher, J.M. et al. Characterization of beta-tubulin genes in hookworms and investigation of resistance-associated mutations using real-time PCR. Mol. Biochem. Parasitol. 156, 167–174, 2007.
37. Churcher, T.S. et al. Mass treatment of parasitic diseases: Implications for the spread of anthelmintic resistance. In: Weber, J.T. (ed.), Antimicrobial Resistance—Beyond the Breakpoint. Issues Infect. Dis., vol. 6, pp. 120–137, Karger, Basel, 2010.
38. Albonico, M. et al. Rate of re-infection with intestinal nematodes after treatment of children with mebendazole or albendazole in a highly endemic area. Trans. R. Soc. Trop. Med. Hyg. 89, 538, 1995.
39. Bethony, J.M. et al. Randomized, placebo-controlled, doubleblind trial of the Na-ASP-2 hookworm vaccine in unexposed adults. Vaccine 26, 2408, 2008.
40. Diemert, D.J., Bethony, J.M., and Hotez, P.J. Hookworm vaccines. Vaccines 46, 282, 2008.
41. Pearson, M.S. An enzymatically inactivated hemoglobinase from Necator americanus induces neutralizing antibodies against multiple hookworm species and protects dogs against heterologous hookworm infection. FASEB J. 23, 3007, 2009.
42. Loukas, A. et al. Vaccination with recombinant aspartic hemoglobinase reduces parasite load and blood loss after hookworm infection in dogs. PLoS Med. 2, e295, 2005. 43. Zhan, B. et al. Biochemical characterization and vaccine potential of a heme-binding glutathione transferase from the adult hookworm Ancylostoma caninum. Infect. Immun. 73, 6903, 2005. 44. Brown, A. et al. An initial characterization of the proteolytic enzymes secreted by the adult stage of the human hookworm Necator americanus. Parasitology 110, 555, 1995. 45. Brown, A., and Pritchard, D.I. The imunogenicity of hookworm (Necator americanus) acetylcholinesterase (AChE) in man. Parasite Immunol. 15, 195, 1993. 46 Pritchard, D.I. et al. Epidemiology and immunology of Necator americanus infection in a community in Papua New Guinea: Humoral responses to excretory-secretory and cuticular collagen antigens. Parasitology 100, 317, 1990. 47. Pritchard, D.I. et al. Isotypic variation in antibody responses in a community in Papua New Guinea to larval and adult antigens during infection, and following reinfection, with the hookworm Necator americanus. Parasite Immunol. 14, 617, 1992. 48. Palmer, D.R., Bradley, M., and Bundy, D.A. IgG4 responses to antigens of adult Necator americanus: Potential for use in large-scale epidemiological studies. Bull. World Health Org. 74, 381, 1996. 49. Quinnell, R.J., Bethony, J., and Pritchard, D.I. The immunoepidemiology of human hookworm infection. Parasite Immunol. 26, 443, 2004. 50. Loukas, A., Constant, S.L., and Bethony, J.M. Immunobiology of hookworm infection. FEMS Immunol. Med. Microbiol. 43, 115, 2005. 51. Chow, S.C. The human hookworm pathogen Necator americanus induces apoptosis in T lymphocytes. Parasitology 22, 21, 2000. 52. Daub, J. et al. A survey of genes expressed in adults of the human hookworm, Necator americanus. Parasitology 120, 171, 2000. 53. Brophy, P.M. et al. Secretory nematode SOD-offensive or defensive? Int. J. Parasitol. 25, 865, 1995. 54. Taiwo, F.A. et al. Cu/Zn superoxide dismutase in excretorysecretory products of the human hookworm Necator americanus: An electron paramagnetic study. Eur. J. Biochem. 264, 434, 1999. 55. Pritchard, D.I. Do haematophagous parasites secrete SOD and promote blood �ow? Int. J. Parasitol. 26, 1339, 1996. 56. Culley, F.J. et al. Eotaxin is speci�cally cleaved by hookworm metalloproteases preventing its action in vitro and in vivo. J. Immunol. 165, 6447, 2000. 57. Pritchard, D.I. et al. A hookworm allergen which strongly resembles calreticulin. Parasite Immunol. 21, 439, 1999. 58. Pritchard, D.I., and Brown, A. Is Necator americanus approaching a mutualistic relationship with humans? Trends Parasitol. 17, 169, 2001. 59. Pritchard, D.I. et al. Immunity in humans to N. americanus: IgE, parasite weight and fecundity. Parasite Immunol. 71, 71, 1995. 60. Croese, J. et al. A proof of concept study establishing Necator americanus in Crohn’s patients and reservoir donors. Gut 55, 136, 2006. 61. Lynch, N.R. et al. Clinical improvement of asthma after anthelminthic treatment in a tropical situation. Am. J. Respir. Crit. Care Med. 156, 50, 1997. 62. Pritchard, D.I. The pro-allergic in�uences of helminth parasites. Mem. Inst. Oswaldo Cruz 92, 15, 1997. 63. Oppenheimer, S.J. Iron and infection: A clinical review. J. Singap. Paediatr. Soc. 36, 1, 1994.
64 Nacher, M. et al. Short report: Hookworm infection is associated with decreased body temperature during mild Plasmodium falciparum malaria. Am. J. Trop. Med. Hyg. 65, 136, 2001.
65 Cooper, P.J. et al. Human infection with Ascaris lumbricoides is associated with a polarized cytokine response. J. Infect. Dis. 182, 1207, 2000.
66. Harada, Y., and Mori, O. A new method for culturing hookworm. Yonago Acta Med. 1, 177, 1955.
67. Polderman, A.M. et al. Oesophagostomiasis, a common infection of man in northern Togo and Ghana. Am. J. Trop. Med. Hyg. 44, 336, 1991.
68. Blotkamp, J. et al. Observations on the morphology of adults and larval stages of Oesophagostomum sp. isolated from man in northern Togo and Ghana. J. Helminthol. 67, 49, 1993.
69. Krepel, H.P. et al. Quantitative interpretation of coprocultures in a population infected with Oesophagostomum bifurcum. Trop. Geogr. Med. 47, 157, 1995.
70. Hebert, P.D. et al. Biological identi�cation though DNA barcodes. Proc. Biol. Sci. 270, 313, 2003.
71. Blaxter, M.L. The promise of a DNA taxonomy. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 359, 669, 2004.
72. Schwenkenbecher, J.M., Mordue, A.J., and Piertney, S.B. Phylogenetic analysis indicates that Culicoides dewul� should not be considered part of the Culicoides obsoletus complex. Bull. Entomol. Res. 99, 371, 2009.
73. Schwenkenbecher, J.M. et al. Discrimination of Culicoides midge larvae using multiplex polymerase chain reaction assays based on DNA sequence variation at the mitochondrial cytochrome C oxidase I gene. J. Med. Entomol., 46, 610, 2009.
74. Zhan, B. et al. Species-speci�c identi�cation of human hookworms by PCR of the mitochondrial cytochrome oxidase I gene. J. Parasitol. 87, 1227, 2001.
75. Hawdon, J.M. et al. Genetic structure of populations of the human hookworm, Necator americanus, in China. Mol. Ecol. 10, 1433, 2001.
76. Yong, T.S. et al. Differential diagnosis of Trichostrongylus and hookworm eggs via PCR using ITS-1 sequences. Korean J. Parasitol. 45, 69, 2007. 77. Verweij, J.J. et al. Simultaneous detection and quanti�cation of Ancylostoma duodenale, Necator americanus, and Oesophagostomum bifurcum in fecal samples using multiplex real-time PCR. Am. J. Trop. Med. Hyg. 77, 685, 2007. 78. Orita, M. et al. Detection of polymorphisms of human DNA by gel electrophoresis as single-strand conformation polymorphisms. Proc. Natl. Acad. Sci. USA 86, 2766, 1989. 79. Gasser, R.B., and Monti, J.R. Identi�cation of parasitic nematodes by PCR-SSCP of ITS-2 rDNA. Mol. Cell. Probes 11, 201, 1997. 80. Gasser, R.B. et al. A mutation scanning approach for the identi�cation of hookworm species and analysis of population variation, Mol. Biochem. Parasitol. 92, 303, 1998. 81. Hu, M. et al. Single-strand conformation polymorphismbased analysis of mitochondrial cytochrome c oxidase subunit 1 reveals signi�cant substructuring in hookworm populations. Electrophoresis 23, 27, 2002. 82. Saiki, R.K. et al. Enzymatic ampli�cation of beta-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230, 1350, 1985. 83. Hawdon, J.M. Differentiation between the human hookworms Ancylostoma duodenale and Necator americanus using PCRRFLP. J. Parasitol. 82, 642, 1996. 84. Kato, K., and Miura, M. Comparative examinations. Jpn. J. Parasitol. 3, 35, 1954. 85. Katz, N., Chavez, A., and Pellegrino, J. A simple device for quantitative stool thick-smear technique in Schistosomiasis mansoni. Rev. Inst. Med. Trop. Sao Paulo 14, 397, 1972. 86. Engels, D., Nahimana, S., and Gryseels, B. Comparison of the direct faecal smear and two thick smear techniques for the diagnosis of intestinal parasitic infections, Trans. R. Soc. Trop. Med. Hyg. 90, 523, 1996. 87. Santos, F.L., Cerqueira, E.J., and Soares, N.M., Comparison of the thick smear and Kato-Katz techniques for diagnosis of intestinal helminth infections. Rev. Soc. Bras. Med. Trop. 38, 196, 2005. 88. Rivera, W.L., Tachibana, H., and Kanbara, H. Application of the polymerase chain reaction (PCR) in the epidemiology of Entamoeba histolytica and Entamoeba dispar infections, Tokaj J. Exp. Clin. Med. 23, 413, 1998. 60 Chapter 60 - Oesophagostomum
1. Blotkamp, J. et al. Observations on the morphology of adults and larval stages of Oesophagostomum sp. isolated from man in northern Togo and Ghana. J. Helminthol. 67, 49, 1993.
2. Railliet, A., and Henry, A. Encore un nouveau sclerostomien (Oesophagostomum brumpti nov. sp.) parasite de l’homme. Comtes Rendus De Seances De La Societe De Biologie 58, 643, 1905.
3. Haaf, E., and van Soest, A.H. Oesophagostomiasis in man in North Ghana. Trop. Geogr. Med. 16, 49, 1964.
4. Gigase, P. et al. Frequency of symptomatic human oesophagostomiasis (helminthoma) in Northern Togo. In: Geerts, S., Kumar, V., and Brandt, J. (eds), Helminth Zoonosis, p. 223, Martinus Nijhoff Publ., Dordrecht, 1987.
5. Polderman, A.M. et al. Oesophagostomiasis, a common infection of man in northern Togo and Ghana. Am. J. Trop. Med. Hyg. 44, 336, 1991.
6. Krepel, H.P. et al. Human Oesophagostomum infection in northern Togo and Ghana: Epidemiological aspects. Ann. Trop. Med. Parasitol. 86, 289, 1992.
7. Krepel, H.P., and Polderman, A.M. Egg production of Oesophagostomum bifurcum, a locally common parasite of humans in Togo. Am. J. Trop. Med. Hyg. 46, 469, 1992.
8. Krepel, H.P. et al. Treatment of mixed Oesophagostomum and hookworm infection: Effect of albendazole, pyrantel pamoate, levamisole and thiabendazole. Trans. R. Soc. Trop. Med. Hyg. 87, 87, 1993.
9. Krepel, H.P. et al. Reinfection patterns of Oesophagostomum bifurcum after anthelmintic treatment. Trop. Geogr. Med. 47, 160, 1995.
10. Krepel, H.P. et al. Quantitative interpretation of coprocultures in a population infected with Oesophagostomum bifurcum. Trop. Geogr. Med. 47, 157, 1995.
11. de Gruijter, J.M. et al. Screening for haplotypic variability within Oesophagostomum bifurcum (Nematoda) employing a single-strand conformation polymorphism approach. Mol. Cell Probes 16, 185, 2002. 12. Gasser, R.B. et al. Screening for nucleotide variations in ribosomal DNA arrays of Oesophagostomum bifurcum by polymerase chain reaction-coupled single-strand conformation polymorphism. Electrophoresis 20, 1486, 1999.
13. Pit, D.S. et al. The capacity of the third-stage larvae of Oesophagostomum bifurcum to survive adverse conditions. Ann. Trop. Med. Parasitol. 94, 165, 2000. 14. Polderman, A.M., and Blotkamp, J. Oesophagostomum infections in humans. Parasitol. Today 11, 451, 1995. 15. Gasser, R.B. et al. Insights into the epidemiology and genetic make-up of Oesophagostomum bifurcum from human and non-human primates using molecular tools. Parasitology 132, 453, 2006. 16. de Gruijter, J.M. et al., Morphological variability within Oesophagostomum bifurcum among different primate species from Ghana. J. Helminthol., 80, 357, 2006. 17. Eberhard, M.L. et al. Experimental Oesophagostomum bifurcum in monkeys. J. Helminthol. 75, 51, 2001. 18. de Gruijter, J.M. et al. High resolution DNA �ngerprinting by AFLP to study the genetic variation among Oesophagostomum bifurcum (Nematoda) from human and non-human primates from Ghana. Parasitology 130, 229, 2005. 19. de Gruijter, J.M. et al. Genetic substructuring within Oesophagostomum bifurcum (Nematoda) from human and nonhuman primates from Ghana based on random ampli�ed polymorphic DNA analysis. Am. J. Trop. Med. Hyg. 71, 227, 2004. 20. Storey, P.A. et al. Clinical epidemiology and classi�cation of human oesophagostomiasis. Trans. Roy. Soc. Trop. Med. Hyg. 94, 177, 2000. 21. Storey, P.A. et al. Ultrasound diagnosis of oesophagostomiasis. Br. J. Radiol. 73, 328, 2000. 22. Storey, P.A. et al. Ultrasound appearance of preclinical Oesophagostomum bifurcum induced colonic pathology. GUT 48, 565, 2001. 23. Storey, P.A. et al. Ultrasonographic detection and assessment of preclinical Oesophagostomum bifurcum—Induced colonic pathology. Clin. Inf. Dis. 33, 166, 2001. 24. Storey, P.A. et al. Intraobserver and interobserver variation of ultrasound diagnosis of Oesophagostomum bifurcum colon lesions. Am. J. Trop. Med. Hyg. 67, 680, 2002. 25. Storey, P.A. et al. The effect of albendazole on Oesophagostomum bifurcum infection and pathology in children from rural northern Ghana. Ann. Trop. Med. Parasitol. 95, 87, 2001. 26. Katz, N. et al. A simple device for quantitative stool thicksmear technique in Schistosomiasis mansoni. Rev. Inst. Med. Trop. Sao Paulo 14, 397, 1972. 27. Verweij, J.J. et al. PCR assay for the speci�c ampli�cation of Oesophagostomum bifurcum DNA from human faeces. Int. J. Parasitol. 30, 137, 2000. 28. Verweij, J.J. et al. Determining the prevalence of Oesophagostomum bifurcum and Necator americanus infections using speci�c PCR ampli�cation of DNA from faecal samples. Trop. Med. Int. Health 6, 726, 2001. 29. Verweij, J.J. et al. Simultaneous detection and quanti�cation of Ancylostoma duodenale, Necator americanus, and Oesophagostomum bifurcum in fecal samples using multiplex real-time PCR. Am. J. Trop. Med. Hyg. 77, 685, 2007. 30. de Gruijter, J.M. et al. Polymerase chain reaction-based differential diagnosis of Ancylostoma duodenale and Necator americanus infections in humans in northern Ghana. Trop. Med. Int. Health 10, 574, 2005. 31. Pit, D.S.S. et al. Parasite-speci�c antibody and cellular immune responses in human infected with Necator americanus and Oesophagostomum bifurcum. Parasitol. Res. 87, 722, 2001. 32. Polderman, A.M. et al. Serological diagnosis of Oesophagostomum infections. Trans. R. Soc. Trop. Med. Hyg. 87, 433, 1993. 33. Romstad, A. et al. Differentiation of Oesophagostomum bifurcum from Necator americanus by PCR using genetic markers in spacer ribosomal DNA. Mol. Cell Probes 11, 169, 1997.
34. de Gruijter, J.M. et al. AFLP �ngerprinting for the analysis of genetic diversity within Necator americanus. Mol. Cell Probes 20, 317, 2006.
35. Gasser, R.B. et al. DNA technological progress toward advanced diagnostic tools to support human hookworm control. Biotechnol. Adv. 26, 35, 2008.
36. Verweij, J.J. et al. Prevalence of Entamoeba histolytica and Entamoeba dispar in northern Ghana. Trop. Med. Int. Health 8, 1153, 2003.
37. Gasser, R.B. et al. Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res. 21, 2525, 1993. 38. Niesters, H.G. Clinical virology in real time. J. Clin. Virol. 25(Suppl 3), S3, 2002. 39. Ziem, J.B. et al. Annual mass treatment with albendazole might eliminate human oesophagostomiasis from the endemic focus in northern Ghana. Trop. Med. Int. Health 11, 1759, 2006. 40. Ziem, J.B. et al. Impact of repeated mass treatment on human Oesophagostomum and hookworm infections in northern Ghana. Trop. Med. Int. Health 11, 1764, 2006. 41. Ziem, J.B. et al. Mass treatment with albendazole reduces the prevalence and severity of Oesophagostomum-induced nodular pathology in northern Ghana. Trans. R. Soc. Trop. Med. Hyg. 100, 760, 2006. 61 Chapter 61 - Onchocerca
1. Bain, O. Evolutionary relationships among �larial nematodes. In: Klei, T.R., and Rajan, T.V. (eds) World Class Parasites. The Filaria, Vol. 5, Kluwer Academic Publishers, Boston, p. 21, 2002.
2. Egyed, Z. et al. Morphologic and genetic characterization of Onchocerca lupi infecting dogs. Vet. Parasitol. 102, 309, 2001.
3. Egyed, Z. et al. Molecular phylogenetic analysis of Onchocerca lupi and its Wolbachia endosymbiont. Vet. Parasitol. 108, 153, 2002.
4. Sreter-Lancz, Z., Szell, Z., and Sreter, T. Molecular genetic comparison of Onchocerca sp. infecting dogs in Europe with other spirurid nematodes including Onchocerca lienalis. Vet. Parasitol. 148, 365, 2007.
5. Sreter, T., and Szell, Z. Onchocercosis: A newly recognized disease in dogs. Vet. Parasitol. 151, 1, 2008.
5a. Faísca, P. et al. A case of canine ocular onchocercosis in Portugal. Vet. Ophthalmol. 13, 117, 2010. 6. Eberhard, M.L. et al. Ocular Onchocerca infections in two dogs in western United States. Vet. Parasitol. 90, 333, 2000. 7. Orihel, T.C. et al. Onchocerciasis in a California dog. Am. J. Trop. Med. Hyg. 44, 513, 1991. 8. Anderson, R.C. Nematode Parasites of Vertebrates: Their Development and Transmission, 2nd. CABI Publishing, Cambridge, MA, 2000. 9. Morales-Hojas, R. Molecular systematics of �larial parasites, with an emphasis on groups of medical and veterinary importance, and its relevance for epidemiology. Infect. Genet. Evol. 9, 748, 2009. 10. Bain, O. Le genre Onchocerca: Hypothèses sur son évolution et clé dichotomique des espèces. Ann. Parasitol. Hum. Comp. 56, 503, 1981. 11. Chabaud, A.G., and Bain, O. The evolutionary expansion of the Spirurida. Int. J. Parasitol. 24, 1179, 1994. 12. Krueger, A., Fischer, P., and Morales-Hojas, R. Molecular phylogeny of the �laria genus Onchocerca with special emphasis on Afrotropical human and bovine parasites. Acta Trop. 101, 1, 2007. 13. Morales-Hojas, R., Cheke, R.A., and Post, R.J. Molecular systematics of �ve Onchocerca species (Nematoda: Filarioidea) including the human parasite, O. volvulus, suggest sympatric speciation. J. Helminthol. 80, 281, 2006. 14. Muller, R. Identi�cation of Onchocerca. In: Proceedings 17th Symposium of the British Society for Parasitology, Problems in the Identi�cation ofParasites and their Vectors, Blackwell, Oxford, p. 175, 1979. 15. Fain, A., Herin, V., and Thienpont, D. Filariases of cattle in Ruanda-Urundi. III. Parasitological study. B. Filariae of the genera Setaria and Onchocerca, and micro�lariae in blood and skin. Ann. Soc. Belg. Med. Trop. 35, 555, 1955. 16. Duke, B.O. The population dynamics of Onchocerca volvulus in the human host. Trop. Med. Parasitol. 44, 61, 1993. 17. Patnaik, B. Onchocerciasis due to O. armillata in cattle in Orissa. J. Helminthol. 36, 313, 1962. 18. Bain, O. Redescription of �ve species of Onchocerca (author’s translation). Ann. Parasitol. Hum. Comp. 50, 763, 1975. 19. ElBihari, S., and Hussein, H.S. Location of the micro�lariae of Onchocerca armillata. J. Parasitol. 61, 656, 1975. 20. WHO. Onchocerciasis and its control. World Health Organ. Tech. Rep. Ser. 852, 1, 1995. 21. Burnham, G. Onchocerciasis. Lancet 351, 1341, 1998. 22. Plaisier, A.P. et al. The reproductive lifespan of Onchocerca volvulus in West African savanna. Acta Trop. 48, 271, 1991. 23. Schulz-Key, H., and Soboslay, P.T. Reproductive biology and population dynamics of Onchocerca volvulus in the vertebrate host. Parasite, 1, S53, 1994. 24. Schulz-Key, H. Observations on the reproductive biology of Onchocerca volvulus. Acta Leiden 59, 27, 1990. 25. Dalmat, H.T. The black�ies (Diptera, Simuliidae) of Guatemala and their role as vectors of onchocerciasis. Smithson. Misc. Collect. 125, 1425, 1955. 26. Duke, B.O. Human onchocerciasis—An overview of the disease. Acta Leiden 59, 9, 1990. 27. Zimmerman, P.A. et al. Recent evolutionary history of American Onchocerca volvulus, based on analysis of a tandemly repeated DNA sequence family. Mol. Biol. Evol. 11, 384, 1994. 28. Morales-Hojas, R. et al. Characterisation of nuclear ribosomal DNA sequences from Onchocerca volvulus and Mansonella ozzardi (Nematoda: Filarioidea) and development of a PCRbased method for their detection in skin biopsies. Int. J. Parasitol. 31, 169, 2001.
29. Morales-Hojas, R., Cheke, R.A., and Post, R.J. A preli minary analysis of the population genetics and molecular phylogenetics of Onchocerca volvulus (Nematoda: Filarioidea) using nuclear ribosomal second internal trans cribed spacer sequences. Mem. Inst. Oswaldo Cruz 102, 879, 2007.
30. Hoerauf, A. et al. Onchocerciasis. BMJ 326, 207, 2003.
31. Duke, B.O., and Anderson, J. A comparison of the lesions produced in the cornea of the rabbit eye by micro�lariae of the forest and Sudan-savanna strains of Onchocerca volvulus from Cameroon. I. The clinical picture. Z. Tropenmed. Parasitol. 23, 354, 1972.
32. Anderson, J. et al. Studies on onchocerciasis in the United Cameroon Republic. II. Comparison of onchocerciasis in rain-forest and Sudan-savanna. Trans. R. Soc. Trop. Med. Hyg. 68, 209, 1974.
33. WHO. WHO expert committee on onchocerciasis. World Health Organ. Tech. Rep. Ser. 752, 1, 1987.
34. Anosike, J.C. et al. The prevalence, intensity and clinical manifestations of Onchocerca volvulus infection in Toro local government area of Bauchi State, Nigeria. Int. J. Hyg. Environ. Health 203, 459, 2001.
35. De Leon, J.R., and Duke, B.O. Experimental studies on the transmission of Guatemalan and West African strains of Onchocerca volvulus by Simulium ochraceum, S. metallicum and S. callidum. Trans. R. Soc. Trop. Med. Hyg. 60, 735, 1966.
36. Duke, B.O., Moore, P.J., and De Leon, J.R. Onchocerca– Simulium complexes. V. The intake and subsequent fate of micro�lariae of a Guatemalan strain of Onchocerca volvulus in forest and Sudan-savanna forms of West African Simulium damnosum. Ann. Trop. Med. Parasitol. 61, 332, 1967.
37. Duke, B.O., Lewis, D.J., and Moore, P.J. Onchocerca– Simulium complexes. I. Transmission of forest and Sudansavanna strains of Onchocerca volvulus, from Cameroon, by Simulium damnosum from various West African bioclimatic zones. Ann. Trop. Med. Parasitol. 60, 318, 1966.
38. Duke, B.O. Onchocerca-Simulium complexes. VI. Experimental studies on the transmission of Venezuelan and West African strains of Onchocerca volvulus by Simulium metallicum and S. exiguum in Venezuela. Ann. Trop. Med. Parasitol. 64, 421, 1970.
39. Basáñez, M.G. et al. Onchocerca-Simulium complexes in Venezuela: Can human onchocerciasis spread outside its present endemic areas? Parasitology 120, 143, 2000.
40. Basáñez, M.G., Churcher, T.S., and Grillet, M.E. Onchocerca– Simulium interactions and the population and evolutionary biology of Onchocerca volvulus. Adv. Parasitol. 68, 263, 2009.
41. Walsh, J.F. et al., Standardization of criteria for assessing the effect of Simulium control in onchocerciasis control programmes. Trans. R. Soc. Trop. Med. Hyg. 72, 675, 1978.
42. Molyneux, D.H. Onchocerciasis control and elimination: Coming of age in resource-constrained health systems. Trends Parasitol. 21, 525, 2005.
43. Ottesen, E.A. Immune responsiveness and the pathogenesis of human onchocerciasis. J. Infect. Dis. 171, 659, 1995.
44. Turaga, P.S. et al. Immunity to onchocerciasis: Cells from putatively immune individuals produce enhanced levels of interleukin-5, gamma interferon, and granulocyte– macrophage colony-stimulating factor in response to Onchocerca volvulus larval and male worm antigens. Infect. Immun. 68, 1905, 2000.
45. Abraham, D., Lucius, R., and Trees, A.J. Immunity to Onchocerca spp. in animal hosts. Trends Parasitol. 18, 164, 2002. 46. Brattig, N.W. et al. Onchocerca volvulus-exposed persons fail to produce interferon-gamma in response to O. volvulus antigen but mount proliferative responses with interleukin-5 and IL-13 production that decrease with increasing micro�larial density. J. Infect. Dis. 185, 1148, 2002. 47. Saint Andre, A. et al. The role of endosymbiotic Wolbachia bacteria in the pathogenesis of river blindness. Science 295, 1892, 2002. 48. Brattig, N.W., Buttner, D.W., and Hoerauf, A. Neutrophil accumulation around Onchocerca worms and chemotaxis of neutrophils are dependent on Wolbachia endobacteria. Microbes Infect. 3, 439, 2001. 49. Brattig, N.W. et al. Lipopolysaccharide-like molecules derived from Wolbachia endobacteria of the �laria Onchocerca volvulus are candidate mediators in the sequence of in�ammatory and anti-in�ammatory responses of human monocytes. Microbes Infect. 2, 1147, 2000. 50. Cross, H.F. et al. Severe reactions to �larial chemotherapy and release of Wolbachia endosymbionts into blood. Lancet 358, 1873, 2001. 51. Taylor, M.J. et al. Wolbachia bacteria in �larial immunity and disease. Parasite Immunol. 23, 401, 2001. 52. Fischer, P. et al. An aspartate aminotransferase of Wolbachia endobacteria from Onchocerca volvulus is recognized by IgG1 antibodies from residents of endemic areas. Parasitol. Res. 90, 38, 2003. 53. Taylor, M.J., Bandi, C., and Hoerauf, A. Wolbachia bacterial endosymbionts of �larial nematodes. Adv. Parasitol. 60, 245, 2005. 54. Johnston, K.L., and Taylor, M.J. Wolbachia in �larial parasites: Targets for �larial infection and disease control. Curr. Infect. Dis. Rep. 9, 55, 2007. 55. WHO. Bench Aids for the Diagnosis of Filarial Infections. WHO, Geneva, 1997. 56. Prost, A., and Prod’hon, J. Parasitological diagnosis of onchocerciasis. A critical review of present methods (author’s transl). Med. Trop. (Mars) 38, 519, 1978. 57. Schulz-Key, H., and Karam, M. Quantitative assessment of micro�lariae and adults of Onchocerca volvulus in ethanol�xed biopsies and nodules. Trans. R. Soc. Trop. Med. Hyg. 78, 157, 1984. 58. Fischer, P. et al. Parasitological and clinical characterization of Simulium neavei-transmitted onchocerciasis in western Uganda. Trop. Med. Parasitol. 44, 311, 1993. 59. Schulz-Key, H., Albiez, E.J., and Buttner, D.W. Isolation of living adult Onchocerca volvulus from nodules. Tropenmed. Parasitol. 28, 428, 1977. 60. Fischer, P. et al. Strain differentiation of Onchocerca volvulus from Uganda using DNA probes. Parasitology 112, 401, 1996. 61. Post, R.J. et al. The morphological discrimination of micro�lariae of Onchocerca volvulus from Mansonella ozzardi. Parasitology 127, 21, 2003. 62. Ewert, A., Smith, J.H., and Corredor, A. Micro�lariae of Mansonella ozzardi in human skin biopsies. Am. J. Trop. Med. Hyg. 30, 988, 1981. 63. Moraes, M.A. et al. The concentration of Mansonella ozzardi micro�lariae in skin capillaries. Trans. R. Soc. Trop. Med. Hyg. 77, 463, 1983. 64. Moraes, M.A.P. Mansonella ozzardi micro�lariae in skin snips. Trans. R. Soc. Trop. Med. Hyg. 70, 16, 1976. 65. Meredith, S.E. et al. Cloning and characterization of an Onchocerca volvulus speci�c DNA sequence. Mol. Biochem. Parasitol. 36, 1, 1989.
66. Erttmann, K.D. et al. Isolation and characterization of form speci�c DNA sequences of O. volvulus. Acta Leiden 59, 253, 1990.
67. Erttmann, K.D. et al. A DNA sequence speci�c for forest form Onchocerca volvulus. Nature 327, 415, 1987.
68. Harnett, W. et al. An oligonucleotide probe speci�c for Onchocerca volvulus. Mol. Biochem. Parasitol. 35, 119, 1989.
69. Meredith, S.E. et al. Onchocerca volvulus: Application of the polymerase chain reaction to identi�cation and strain differentiation of the parasite. Exp. Parasitol. 73, 335, 1991.
70. Shah, J.S. et al. Characterization of an Onchocerca-speci�c DNA clone from Onchocerca volvulus. Am. J. Trop. Med. Hyg. 37, 376, 1987.
71. Zimmerman, P.A. et al. Polymerase chain reaction-based diagnosis of Onchocerca volvulus infection: Improved detection of patients with onchocerciasis. J. Infect. Dis. 169, 686, 1994.
72. Zimmerman, P.A., Toe, L., and Unnasch, T.R. Design of Onchocerca DNA probes based upon analysis of a repeated sequence family. Mol. Biochem. Parasitol. 58, 259, 1993. 73. Unnasch, T.R., and Meredith, S.E. The use of degenerate primers in conjunction with strain and species oligonucleotides to classify Onchocerca volvulus. Methods Mol. Biol. 50, 293, 1996. 74. Hojas, R.M., and Post, R.J. Regional genetic variation in the major sperm protein genes of Onchocerca volvulus and Mansonella ozzardi (Nematoda: Filarioidea). Int. J. Parasitol. 30, 1459, 2000. 75. Tang, T.H. et al. Nested PCR to detect and distinguish the sympatric �larial species Onchocerca volvulus, Mansonella ozzardi and Mansonella perstans in the Amazon Region. Mem. Inst. Oswaldo Cruz 105, 823, 2010. 76. Oskam, L. et al. Polymerase chain reaction for detecting Onchocerca volvulus in pools of black�ies. Trop. Med. Int. Health 1, 522, 1996. 77. Nutman, T.B. et al. A universally applicable diagnostic approach to �larial and other infections. Parasitol. Today 10, 239, 1994. 78. Newton, L.A. et al. Systematic relationships of some members of the Oesophagostomum and Chabertia (Nematoda: Chabertiidae) based on ribosomal DNA sequence data. Int. J. Parasitol. 28, 1781, 1998. 62 Chapter 62 - Pseudoterranova and Contracaecum
7. Koh, M.S. et al. A case of gastric pseudoterranoviasis in a 43-year old man in Korea. Korean J. Parasitol. 37, 47, 1999. [In Korean].
8. Yu, J.R. et al. A human case of gastric infection by Pseudoterranova decipiens larva. Korean J. Parasitol. 39, 193, 2001.
9. Vidal-Martinez, V.M. et al. Experimental infection of Contracaecum multipapillatum (Nematoda, Anisakidae) from México in the domestic cat. J. Parasitol. 80, 576, 1994.
10. Mattiucci, S., and Nascetti, G. Advances and trends in the molecular systematics of Anisakid nematodes, with implications for their evolutionary ecology and host-parasite co- evolutionary processes. Adv. Parasitol. 66, 47, 2008.
11. Baylis, H.A. Some ascarids in the British Museum (Nat. Hist.). Parasitology 8, 360, 1916.
12. Johnston, T.H., and Mawson, P.M. Parasitic nematodes. Rep. Br. N. Z. Aust. Antartic Res. Exp. 5, 74, 1945.
13. Karokhin, V.I. Two new species of Porrocacum from Siberian birds of prey. In: Pod’yanolskaya, V.P. (ed.), Helminthological Collection. Nauka, Moscow, pp. 132–141, 1946. [In Russian].
14. Myers, B.J. Phocanema, a new genus for the anisakid nematode of seals. Can. J. Zool. 37, 459, 1959.
15. Gibson, D.I. The systematics of ascaridoid nematodes-a current assessment. In: Stone, A.R., Platt, H.M., and Khalil, L.F. (eds), Concepts in Nematode Systematics. Academic Press, New York, pp. 321–338, 1983.
16. Mozgovoi, A.A., Ascaridata. In: Skrjabin, K.I., Shikhobalova, N.P., and Mozgovoi, A.A., (eds), Descriptive Catalogue of Parasitic Nematodes, Vol. II. Press of Academy of Sciences of USSR, Moscow, Russia, pp. 537–542, 1951. [In Russian]
17. Gibson, D.I., and Colin, J.A. The Terranova enigma. Parasitology 85, 36, 1982.
18. Deardorff, T.L., and Overstreet, R.M. Terranova ceticola n. sp. (Nematoda: Anisakidae) from the dwarf sperm whale; Kogia simus (Owen), in the Gulf of Mexico. Syst. Parasitol. 3, 25, 1981.
19. Paggi, L. et al. Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the North Atlantic and Norwegian and Barents Seas. Int. J. Parasitol. 21, 195, 1991.
20. Paggi, L. et al. Pseudoterranova decipiens species A and B (Nematoda: Ascaridoidea): Nomenclatural designation, morphological diagnostic characters and genetic markers. Syst. Parasitol. 45, 185, 2000.
21. Brattey, J., and Stenson, G.B. Host speci�city and abundance of parasitic nematodes (Ascaridoidea) from the stomachs of �ve phocid species from Newfoundland and Labrador. Can. J. Zool. 71, 2156, 1993.
22. Mattiucci, S. et al. Allozyme and morphological identi�cation of Anisakis, Contracaecum and Pseudoterranova from Japanese waters (Nematoda: Ascaridoidea). Syst. Parasitol. 40, 81, 1998.
23. Paggi, L. et al. A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whale (Ziphiidae): Allozyme and morphological evidence. Syst. Parasitol. 40, 161, 1998.
24. Nadler, S.A. et al. Molecular phylogenetics and diagnosis of Anisakis, Pseudoterranova, and Contracaecum from northern paci�c marine mammals. J. Parasitol. 91, 1413, 2005.
42. Nagano, K. Gastric terranovasis. In: Ishikura, H.M., and Namiki, M., (eds), Gastric anisakiasis in Japan, SpringerVerlag, Tokyo, pp. 133–140, 1989.
43. Mercado, R. et al. Human case of gastric infection by a fourth larval stage of Pseudoterranova decipiens (Nematoda, Anisakidae). Rev. Saude Publica 31, 178, 1997.
44. Mercado, R. et al. Human infection by Pseudoterranova decipiens (Nematoda, Anisakidae) in Chile: Report of seven cases. Mem. Inst. Oswaldo Cruz 96, 653, 2001.
45. Pinel, C. et al. Gastric anisakidosis due to Pseudoterranova decipiens larva. Lancet 347, 1829, 1996.
46. Fioravanti, M.L. et al. Pseudoterranova decipiens (Nematoda: Anisakidae), human infection in Italy, Parassitologia 46, S112, 2004.
47. Oshima, T. Anisakiasis—Is the sushi bar guilty? Parasitology 3, 44, 1987.
48. Adams, A.M. et al. Parasites of risks to public health. Rev. Sci. Tech. Off. Int. Epiz. 16, 652, 1997.
49. Deardorff, T.L. et al. Histopathology induced by larval Terranova (Type HA) (Nematoda: Anisakinae) in experimentally infected rats. J. Parasitol. 69, 191, 1983.
50. Smith, J.W. Ascaridoid nematodes and pathology of the alimentary tract and its associated organs in vertebrates, including man: A literature review. Helminthol. Abstr. 68, 49, 1999.
51. Little, M.D., and Most, H. Anisakid larva from the throat of a woman in New York. Am. J. Trop. Med. Hyg. 22, 609, 1973.
52. Amin, O.M. et al. An unusual case of anisakiasis in California, U.S.A. Comp. Parasitol. 67, 71, 2000.
53. Asaishi, K. et al. Studies on the pathogenic mechanisms of anisakiasis: II Histological changes of stomach wall due to the injection of worm extracts. Jpn. J. Parasitol. 23, 34, 1974. (in Japanese)
54. Berland, B. Nematodes from some Norwegian marine �shes. Sarsia 2, 1, 1961.
55. Mattiucci, S. et al. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: Morphological description, genetic relationships between congeners and ecological data. Syst. Parasitol. 74, 199, 2009.
56. Mattiucci, S. et al. Contracaecum gibsoni n. sp. and C. overstreeti n. sp. (Nematoda: Anisakidae) from Dalmatian pelican Pelecanus crispus (L.) in Greek waters: Genetic and morphological evidence. Syst. Parasitol. 75, 207, 2010.
57. Zhu, X.Q. et al. Assessing sequence variation in the internal transcribed spacers of ribosomal DNA within and among members of the Contracaecum osculatum complex (Nematoda: Ascaridoidea: Anisakidae). Parasitol. Res. 86, 677, 2000. 58. Hu, M. et al. Mutation scanning for sequence variation in three mitochondrial DNA regions for members of the Contracaecum osculatum (Nematoda:Ascaridoidea) complex. Electrophoresis 22, 1069, 2001. 59. McClelland, G. The trouble with sealworms (Pseudoterranova decipiens species complex, Nematoda): A Review. Parasitology 124, 183, 2002. 60. Levsen, A. et al. Low detection ef�ciency of candling as a commonly recommended inspection method for nematode larvae in the �esh of pelagic �sh. J. Food Prot. 68, 828, 2005. 61. Murray, M.G., and Thompson, W.F. Rapid isolation of highmolecular-weight plant DNA. Nucleic Acids Res. 8, 4321, 1980. 62. Valentini, A. et al. Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial cox-2 sequences, and comparison with allozyme data. J. Parasitol. 92, 156, 2006. 63. Hall, T.A. BioEdit: A user-friendly biological sequence alignment and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser. 41, 95, 1999. 64. Swofford, D.L. PAUP*. Phylogenetic Analysis using Parsimony (*and Other Methods). Sinauer Associates, Sunderland, Massachusetts, 2003. 65. Paltridge, G.P. et al. Intestinal anisakiasis : A New Zealand disease. N. Z. Med. J. 97, 558, 1984. 66. Palm, H.W. et al. Molecular genotyping of Anisakis Dujardin, 1845 (Nematoda: Ascaridoidea: Anisakidae) larvae from marine �sh of Balinese and Javanese waters, Indonesia. Helminthologia 45, 3, 2008. 67. Mattiucci, S. et al. Human Anisakidosis in Italy: Molecular and histological identi�cation of two new cases. Parassitologia 49, S226, 2007. 68. Cattan, P.E., and Carvajal, J. A study of the migration of larval Anisakis simplex (Nematoda: Ascaridida) in the Chilean hake, Merluccius gayi (Guichenot). J. Fish Biol. 24, 649, 1984. 69. Rodriguez-Mahillo, A.I. et al. Cloning and characterization of the Anisakis simplex allergen Ani s4 as a cysteine-protease inhibitor. Int. J. Parasitol. 37, 907, 2007. 70. Moneo, I. et al. Isolation of a heat-resistant allergen from �sh parasite Anisakis simplex. Parasitol. Res. 96, 285, 2005. 71. Stormo, S.K. et al. Cold tolerance in sealworm (Pseudoterranova decipiens) due to heat-shock adaptations. Parasitology 136, 1317, 2009. 72. Arizono, N. et al., Human infection with Pseudoterranova azarasi roundworm. Emerg. Infect. Dis., 17, 3, March 2011. 63 Chapter 63 - Strongyloides
6. King, S.E., and Mascie-Taylor, C.G., Strongyloides fuelleborni kellyi and other intestinal helminths in children from Papua New Guinea: Associations with nutritional status and socioeconomic factors. P. N. G. Med. J., 47, 181, 2004.
7. Little, M.D., Comparative morphology of six species of Strongyloides (Nematoda) and rede�nition of the genus. J. Parasitol., 52, 69, 1966.
8. Yamada, M. et al., Species-speci�c differences in heterogonic development of serially transferred free-living generations of Strongyloides planiceps and Strongyloides stercoralis. J. Parasitol., 77, 592, 1991.
9. Gill, G.V. et al., A British Second World War veteran with disseminated strongyloidiasis. Trans. R. Soc. Trop. Med. Hyg., 98, 382, 2004.
10. Prendki, V. et al., Strongyloidiasis in man 75 years after initial exposure. Emerg. Infect. Dis., 17, 5, 2011.
11. Olsen, A. et al., Strongyloidiasis—the most neglected of the neglected tropical diseases? Trans. R. Soc. Trop. Med. Hyg., 103, 967, 2009.
12. Montes, M., Sawhney, C., and Barros, N., Strongyloides stercoralis: There but not seen. Curr. Opin. Infect. Dis., 23, 500, 2010.
13. Safdar, A. et al., Strongyloidiasis in patients at a comprehensive cancer center in the United States: A retrospective study covering the years 1971–2003. Cancer, 100, 1531, 2004.
14. Steinmann, P. et al., Occurrence of Strongyloides stercoralis in Yunnan Province, China, and comparison of diagnostic methods. PLoS Negl. Trop. Dis., 1, e75, 2007.
15. DeVault, G.A. et al., Opportunistic infections with Strongyloides stercoralis in renal transplantation. Rev. Infect. Dis., 12, 653, 1990.
16. Roman-Sanchez, P. et al., High prevalence of Strongyloides stercoralis among farm workers on the Mediterranean coast of Spain: Analysis of the predictive factors of infection in developed countries. Am. J. Trop. Med. Hyg., 69, 336, 2003.
17. Czachor, J.S., and Jonas, A.P., Transmission of Strongyloides steracolis person to person. J. Travel Med., 7, 211, 2000.
18. Liu, L.X., and Weller, P.F., Strongyloidiasis and other intestinal nematode infections. Infect. Dis. Clin. North. Am., 7, 655, 1993.
19. Mahmoud, A.A., Strongyloidiasis. Clin. Infect. Dis., 23, 949, 1996.
20. Sudarshi, S. et al., Clinical presentation and diagnostic sensitivity of laboratory tests for Strongyloides stercoralis in travellers compared with immigrants in a non-endemic country. Trop. Med. Int. Health, 8, 728, 2003.
21. Keiser, P.B., and Nutman, T.B., Strongyloides stercoralis in the immunocompromised population. Clin. Microbiol. Rev., 17, 208, 2004.
22. Lim, S. et al., Complicated and fatal Strongyloides infection in Canadians: Risk factors, diagnosis and management. CMAJ, 171, 479, 2004.
23. Segarra-Newnham, M., Manifestations, diagnosis, and treatment of Strongyloides stercoralis infection. Ann. Pharmacother., 41, 1992, 2007.
24. Greiner, K., Bettencourt, J., and Semolic, C., Strongyloidiasis: A review and update by case example. Clin. Lab. Sci., 21, 82, 2008.
25. von Kuster, L.C., and Genta, R.M., Cutaneous manifestations of strongyloidiasis. Arch. Dermatol., 124, 1826, 1988.
26. Livneh, A. et al., Strongyloides stercoralis hyperinfection mimicking systemic lupus erythematosus �are. Arthritis Rheum., 31, 930, 1988.
27. Chu, E., Whitlock, W.L., and Dietrich, R.A., Pulmonary hyperinfection syndrome with Strongyloides stercoralis. Chest, 97, 1475, 1990. 28. Ly, M.N. et al., Cutaneous Strongyloides stercoralis infection: An unusual presentation. J. Am. Acad. Dermatol., 49, S157, 2003. 29. Gutierrez, Y. et al., Strongyloides stercoralis eosinophilic granulomatous enterocolitis. Am. J. Surg. Pathol., 20, 603, 1996. 30. Cannavo, S.P., Guarneri, F., and Guarneri, C., Strongyloidiasis histologically mimicking eosinophilic folliculitis. Eur. J. Dermatol., 14, 264, 2004. 31. Currie, B.J., and McCarthy, J.S., Strongyloides stercoralis infection as a manifestation of immune restoration syndrome. Clin. Infect. Dis., 40, 635, 2005. 32. Vithayasai, N., and Jennuvat, S., Gastrointestinal manifestations in severe strongyloidiasis: Report of 3 cases and literature review. Southeast Asian J. Trop. Med. Public Health, 41, 22, 2010. 33. Mans�eld, L.S. et al., Strongyloides stercoralis: Maintenance of exceedingly chronic infections. Am. J. Trop. Med. Hyg., 55, 617, 1996. 34. Sandlund, J.T., Kauffman, W., and Flynn, P.M., Strongyloides stercoralis infection mimicking relapse in a child with small noncleaved cell lymphoma. Am. J. Clin. Oncol., 20, 215, 1997. 35. Thompson, B.F. et al., The spectrum of GI strongyloidiasis: An endoscopic-pathologic study. Gastrointest. Endosc., 59, 906, 2004. 36. Concha, R., Harrington, W.J., and Rogers, A.I., Intestinal strongyloidiasis: Recognition, management, and determinants of outcome. J. Clin. Gastroenterol., 39, 203, 2005. 37. Csermely, L. et al., Strongyloides hyper-infection causing lifethreatening gastrointestinal bleeding. World J. Gastroenterol., 12, 6401, 2006. 38. Ghoshal, U.C. et al., Strongyloides stercoralis infestation in a patient with severe ulcerative colitis. Indian J. Med. Sci., 60, 106, 2006. 39. Yoshida, H. et al., Recurrent paralytic ileus associated with strongyloidiasis in a patient with systemic lupus erythematosus. Mod. Rheumatol., 16, 44, 2006. 40. Qu, Z. et al., Strongyloides colitis is a lethal mimic of ulcerative colitis: The key morphologic differential diagnosis. Hum. Pathol., 40, 572, 2009. 41. Bannon, J.P., Fater, M., and Solit, R., Intestinal ileus secondary to Strongyloides stercoralis infection: Case report and review of the literature. Am. Surg., 61, 377, 1995. 42. Al-Bahrani, Z.R., Al-Saleem, T., and Al-Gailani, M.A., Subacute intestinal obstruction by Strongyloides stercolaris. J. Infect., 30, 47, 1995. 43. Friedenberg, F. et al., Duodenal obstruction caused by Strongyloides stercoralis enteritis in an HTLV-1-infected host. Dig. Dis. Sci., 44, 1184, 1999. 44. Suvarna, D. et al., In�ltrating Strongyloides stercoralis presenting as duodenal obstruction. Indian J. Gastroenterol., 24, 173, 2005. 45. Harish, K. et al., Strongyloidiasis presenting as duodenal obstruction. Trop. Gastroenterol., 26, 201, 2005. 46. Juchems, M.S. et al., Strongyloides stercoralis: A rare cause of obstructive duodenal stenosis. Digestion, 77, 141, 2008. 47. Cruz, R.J. Jr., Vincenzi, R., and Ketzer, B.M., Duodenal obstruction—an unusual presentation of Strongyloides stercoralis enteritis: A case report. World J. Emerg. Surg., 5, 23, 2010. 48. Marcos, L.A. et al., Strongyloides hyperinfection syndrome: An emerging global infectious disease. Trans. R. Soc. Trop. Med. Hyg., 102, 314, 2008.
49. Igra-Siegman, Y. et al., Syndrome of hyperinfection with Strongyloides stercoralis. Rev. Infect. Dis., 3, 397, 1981.
50. Nonaka, D. et al., Paralytic ileus due to strongyloidiasis: Case report and review of the literature. Am. J. Trop. Med. Hyg., 59, 535, 1998.
51. Lemos, L.B. et al., Hyperinfection syndrome in strongyloidiasis: Report of two cases. Ann. Diagn. Pathol., 7, 87, 2003.
52. Potter, A., Stephens, D., and De Keulenaer, B., Strongyloides hyper-infection: A case for awareness. Ann. Trop. Med. Parasitol., 97, 855, 2003.
53. Namisato, S. et al., Pulmonary strongyloidiasis in a patient receiving prednisolone therapy. Intern. Med., 43, 731, 2004.
54. Schaeffer, M.W. et al., Strongyloides hyperinfection syndrome after heart transplantation: Case report and review of the literature. J. Heart Lung Transplant., 23, 905, 2004.
55. Mayayo, E. et al., Strongyloides stercolaris infection mimicking a malignant tumour in a non-immunocompromised patient. Diagnosis by bronchoalveolar cytology. J. Clin. Pathol., 58, 420, 2005.
56. Kaslow, J.E. et al., Disseminated strongyloidiasis: An unheralded risk of corticosteroid therapy. J. Allergy Clin. Immunol., 86, 138, 1990.
57. Newton, R.C. et al., Strongyloides stercoralis hyperinfection in a carrier of HTLV-I virus with evidence of selective immunosuppression. Am. J. Med., 92, 202, 1992.
58. Morgello, S. et al., Central nervous system Strongyloides stercoralis in acquired immunode�ciency syndrome: A report of two cases and review of the literature. Acta Neuropathol. (Berl), 86, 285, 1993.
59. Nandy, A. et al., Fulminating strongyloidiasis complicating Indian kala-azar. Trop. Geogr. Med., 47, 139, 1995.
60. Nucci, M. et al., Strongyloidiasis in patients with hematologic malignancies. Clin. Infect. Dis., 21, 675, 1995.
61. Vadlamudi, R.S., Chi, D.S., and Krishnaswamy, G., Intestinal strongyloidiasis and hyperinfection syndrome. Clin. Mol. Allergy, 4, 8, 2006.
62. Palau, L.A., and Pankey, G.A., Strongyloides hyperinfection in a renal transplant recipient receiving cyclosporine: Possible Strongyloides stercoralis transmission by kidney transplant. Am. J. Trop. Med. Hyg., 57, 413, 1997.
63. Gotuzzo, E. et al., Strongyloides stercoralis hyperinfection associated with human T cell lymphotropic virus type-1 infection in Peru. Am. J. Trop. Med. Hyg., 60, 146, 1999.
64. Adedayo, C.O., Grell, G.A., and Bellot, P., Case study: Fatal strongyloides associated with human T-cell lymphotropic virus type 1 infection. Am. J. Trop. Med. Hyg., 65, 650, 2001.
65. Adedayo, O., Grell, G., and Bellot, P., Hyperinfective strongyloidiasis in the medical ward: A review of 27 cases in 5 years. South Med. J., 95, 711, 2002.
66. Feitosa, G. et al., High prevalence of giardiasis and stronglyloidiasis among HIV-infected patients in Bahia, Brazil. Braz. J. Infect. Dis., 5, 339, 2001.
67. Rodrigues, M.A. et al., Invasive enteritis by Strongyloides stercoralis presenting as acute abdominal distress under corticosteroid therapy. Rev. Hosp. Clin. Fac. Med. Sao Paulo, 56, 103, 2001.
68. Singh, S., Human strongyloidiasis in AIDS era: Its zoonotic importance. J. Assoc. Physicians India, 50, 415, 2002.
69. Orlent, H. et al., Strongyloidiasis pre and post autologous peripheral blood stem cell transplantation. Bone Marrow Transplant., 32, 115, 2003.
70. Carvalho, E.M., and Da Fonseca Porto, A., Epidemiological and clinical interaction between HTLV-1 and Strongyloides stercoralis. Parasite Immunol., 26, 487, 2004. 71. Ohnishi, K. et al., Strongyloidiasis in a patient with acquired immunode�ciency syndrome. J. Infect. Chemother., 10, 178, 2004. 72. Viney, M.E. et al., Why does HIV infection not lead to disseminated strongyloidiasis? J. Infect. Dis., 190, 2175, 2004. 73. Arsic´-Arsenijevic´, V. et al., Fatal Strongyloides sterocoralis infection in a young woman with lupus glomerulonephritis. J. Nephrol., 18, 787, 2005. 74. Ben-Youssef, R. et al., Strongyloides stercoralis infection from pancreas allograft: Case report. Transplantation, 80, 997, 2005. 75. El Masry, H.Z., and O’Donnell, J., Fatal Strongyloides hyperinfection in heart transplantation. J. Heart Lung Transplant, 24, 1980, 2005. 76. Hauber, H.P. et al., Fatal outcome of a hyperinfection syndrome despite successful eradication of Strongyloides with subcutaneous ivermectin. Infection, 33, 383, 2005. 77. Lanzafame, M. et al., Strongyloidiasis in an HIV-1–infected patient after highly active antiretroviral therapy–induced immune restoration (letter). J. Infect. Dis., 191, 1027, 2005. 78. Reddy, I.S., and Swarnalata, G., Fatal disseminated strongyloidiasis in patients on immunosuppressive therapy. Indian J. Dermatol. Venerol. Leprol., 71, 38, 2005. 79. Satyanarayana, S. et al., Disseminated Strongyloides stercoralis in AIDS: A report from India. Indian J. Pathol. Microbiol., 48, 472, 2005. 80. Seet, R.C.S., Lau, L.G., and Tambyah, P.A., Strongyloides hyperinfection and hypogammaglobulinemia. Clin. Diagn. Lab. Immunol., 12, 680, 2005. 81. Brown, M., Cartledge, J.D., and Miller, R.F., Dissemination of Strongyloides stercoralis as an immune restoration phenomenon in an HIV-1–infected man on antiretroviral therapy. Int. J. STD AIDS, 17, 560, 2006. 82. Mora, C.S., Segami, M.I., and Hidalgo, J.A., Strongyloides stercoralis hyperinfection in systemic lupus erythematosus and the antiphospholipid syndrome. Semin. Arthritis Rheum., 36, 135, 2006. 83. Qazilbash, M.H. et al., Strongyloidiasis after unrelated nonmyeloablative allogeneic stem cell transplantation. Bone Marrow Transplant., 38, 393, 2006. 84. Ramdial, P.K., Hlatshwayo, N.H., and Singh, B., Strongyloides stercoralis mesenteric lymphadenopathy: Clue to the etiopathogenesis of intestinal pseudo-obstruction in HIV-infected patients. Ann. Diagn. Pathol., 10, 209, 2006. 85. Fardet, L. et al., Severe strongyloidiasis in corticosteroidtreated patients: Case series and literature review. J. Infect., 54, 18, 2007. 86. Said, T. et al., Hyperinfection strongyloidiasis: An anticipated outbreak in kidney transplant recipients in Kuwait. Transplant. Proc., 39, 1014, 2007. 87. Taylor, C.L. et al., Immune reconstitution syndrome to Strongyloides stercoralis infection (letter). AIDS, 21, 649, 2007. 88. Kia, E.B. et al., A case of fatal strongyloidiasis in a patient with chronic lymphocytic leukemia and molecular characterization of the isolate. Korean J. Parasitol., 46, 261, 2008. 89. Patel, G. et al., Strongyloides hyperinfection syndrome after intestinal transplantation. Transpl. Infect. Dis., 10, 137, 2008. 90. Dulley, F.L. et al., Strongyloides stercoralis hyperinfection after allogeneic stem cell transplantation. Bone Marrow Transplant., 43, 741, 2009. 91. Mizuno, S. et al., Strongyloides hyperinfection syndrome following simultaneous heart and kidney transplantation. Transpl. Int., 22, 251, 2009.
92. Roxby, A.C., Gottlieb, G.S., and Limaye, A.P., Strongyloidiasis in transplant patients. Clin. Infect. Dis., 49, 1411, 2009.
93. Vilela, E.G. et al., Strongyloides stercoralis hyperinfection syndrome after liver transplantation: Case report and literature review. Transpl. Infect. Dis., 11, 132, 2009.
94. Wirk, B., and Wingard, J.R., Strongyloides stercoralis hyperinfection in hematopoietic stem cell transplantation. Transpl. Infect. Dis., 11, 143, 2009.
95. Iriemenam, N.C. et al., Strongyloides stercoralis and the immune response. Parasitol. Int., 59, 9, 2010.
96. Thomas, M.C., and Costello, S.A., Disseminated strongyloidiasis arising from a single dose of dexamethasone before stereotactic radiosurgery. Int. J. Clin. Pract., 52, 520, 1998.
97. Wong, T.Y.H. et al., Nephrotic syndrome in strongyloidiasis: Remission after eradication with antihelminthic agents. Nephron, 79, 333, 1998.
98. Hughes, R., and McGuire, G., Delayed diagnosis of disseminated strongyloidiasis. Intensive Care Med., 27, 310, 2001.
99. Kim, A.C., and Lupatkin, H.C., Strongyloides stercoralis infection as a manifestation of immune restoration syndrome (letter). Clin. Infect. Dis., 39, 439, 2004.
100. Giavina-Bianchi, P. et al., A rare intestinal manifestation in a patient with common variable immunode�ciency and strongyloidiasis. Int. Arch. Allergy Immunol., 140, 199, 2006.
101. Lam, C.S. et al., Disseminated strongyloidiasis: A retrospective study of clinical course and outcome. Eur. J. Clin. Microbiol. Infect. Dis., 25, 14, 2006.
102. Galimberti, R. et al., Disseminated strongyloidiasis in immunocompromised patients–report of three cases. Int. J. Dermatol., 48, 975, 2009.
103. Basile, A. et al., Disseminated Strongyloides stercoralis: hyperinfection during medical immunosuppression. J. Am. Acad. Dermatol., 63, 896, 2010.
104. Hsieh, Y.P., Won, Y.K., and Chen, M.L., Minimal change nephrotic syndrome in association with strongyloidiasis. Clin. Nephrol., 66, 459, 2006.
105. Jain, A.K., Agarwal, S.K., and el Sadr, W., Streptococcus bovis bacteremia and meningitis associated with Strongyloides stercoralis colitis in a patient infected with human immunode�ciency virus. Clin. Infect. Dis., 18, 253, 1994.
106. Link, K., and Orenstein, R., Bacterial complications of strongyloidiasis: Streptococcus bovis meningitis. South Med. J., 92, 728, 1999.
107. Linder, J.D. et al., Streptococcus bovis bacteremia associated with Strongyloides stercoralis colitis. Gastrointest. Endosc., 52, 796, 2000.
108. de Silva, T., Raychaudhuri, M., and Poulton, M., HIV infection associated with Strongyloides stercoralis colitis resulting in Streptococcus bovis bacteraemia and meningitis. Sex. Transm. Infect., 81, 276, 2005.
109. Newberry, A.M. et al., Strongyloides hyperinfection presented as acute respiratory failure and Gram-negative sepsis. Chest, 128, 3681, 2005.
110. Shorman, M., and Al-Taw�q, J.A., Strongyloides stercoralis hyperinfection presenting as acute respiratory failure and Gram-negative sepsis in a patient with astrocytoma. Int. J. Infect. Dis., 13, e288, 2009.
111. Pitisuttithum, P., Supanaranond, W., and Chindanond, D., A randomized comparative study of albendazole and thiabendazole in chronic strongyloidiasis. Southeast Asian J. Trop. Med. Public Health, 26, 735, 1995. 112. Ashraf, M., Gue, C.L., and Baddour, L.M., Case report: Strongyloidiasis refractory to treatment with ivermectin. Am. J. Med. Sci., 311, 178, 1996. 113. Toma, H. et al.. Comparative studies on the ef�cacy of three antihelminthics on treatment of human strongyloidiasis in Okinawa, Japan. Southeast Asian J. Trop. Med. Public Health, 31, 147, 2000. 114. Zaha, O. et al., Strongyloidiasis—progress in diagnosis and treatment. Intern. Med., 39, 695, 2000. 115. Zaha, O. et al., Ef�cacy of ivermectin for chronic strongyloidiasis: Two single doses given 2 weeks apart. J. Infect. Chemother., 8, 94, 2002. 116. Marty, F.M. et al., Treatment of human disseminated strongyloidiasis with a parenteral veterinary formulation of ivermectin. Clin. Infect. Dis., 41, e5, 2005. 117. Pacanowski, J. et al., Subcutaneous ivermectin as a safe salvage therapy in Strongyloides stercoralis hyperinfection syndrome: A case report. Am. J. Trop. Med. Hyg., 73, 122, 2005. 118. Boulware, D.R. et al., Maltreatment of Strongyloides infection: Case series and worldwide physicians-in-training survey. Am. J. Med., 120, e1, 2007. 119. Balagopal, A. et al., Detection and treatment of Strongyloides hyperinfection syndrome following lung transplantation. Transpl. Infect. Dis., 11, 149, 2009. 120. Huston, J.M. et al., Treatment of Strongyloides stercoralis hyperinfection-associated septic shock and acute respiratory distress syndrome with drotrecogin alfa (activated) in a renal transplant recipient. Transpl. Infect. Dis., 11, 277, 2009. 121. Datry, A. et al., Treatment of Strongyloides stercoralis infection with ivermectin compared with albendazole: Results of an open study of 60 cases. Trans. R. Soc. Trop. Med. Hyg., 88, 344, 1994. 122. Chiodini, P.L. et al., Parenteral ivermectin in Strongyloides hyperinfection. Lancet, 355, 43, 2000. 123. Orem, J. et al., Strongyloides stercoralis hyperinfection in a patient with AIDS in Uganda successfully treated with ivermectin. Clin. Infect. Dis., 37, 152, 2003. 124. Gulbas, Z. et al., Successful ivermectin treatment of hepatic strongyloidiasis presenting with severe eosinophilia. South Med. J., 97, 907, 2004. 125. Gann, P.H., Neva, F.A., and Gam, A.A., A randomized trial of single- and two-dose ivermectin versus thiabendazole for treatment of strongyloidiasis. J. Infect. Dis., 169, 1076, 1994. 126. Tarr, P. et al., Case report: Rectal administration of ivermectin to a patient with Strongyloides hyperinfection syndrome. Am. J. Trop. Med. Hyg., 68, 453, 2003. 127. Muennig, P. et al., The cost-effectiveness of ivermectin vs. albendazole in the presumptive treatment of strongyloidiasis in immigrants to the United States. Epidemiol. Infect., 132, 1055, 2004. 128. Pornsuriyasak, P., Niticharoenpong, K., and Sakapibunnan, A., Disseminated strongyloidiasis successfully treated with extended duration ivermectin combined with albendazole: A case report of intractable strongyloidiasis. Southeast Asian J. Trop. Med. Public Health, 35, 531, 2004. 129. Turner, S.A. et al., Parenteral administration of ivermectin in a patient with disseminated strongyloidiasis. Am. J. Trop. Med. Hyg., 73, 911, 2005. 130. Lichtenberger, P. et al., Hyperinfection strongyloidiasis in a liver transplant recipient treated with parenteral ivermectin. Transpl. Infect. Dis., 11, 137, 2009. 131. Grein, J.D. et al., Serum ivermectin levels after enteral and subcutaneous administration for Strongyloides hyperinfection: A case report. Scand. J. Infect. Dis., 42, 234, 2010. 132. Conway, D.J. et al., Towards effective control of Strongyloides stercoralis. Trends Parasitol., 11, 420, 1995.
133. Takayanagui, O.M. et al., Detection of Strongyloides stercoralis in the cerebrospinal �uid of a patient with acquired immunode�ciency syndrome. Neurology, 45, 193, 1995.
134. Marchi, B.J., and Cantos, G.A., Evaluation of techniques for the diagnosis of Strongyloides stercoralis in human immunode�ciency virus (HIV) positive and HIV negative individuals in the city of Itajai, Brazil. Braz. J. Infect. Dis., 7, 402, 2003.
135. Siddiqui, A.A., and Berk, S.L., Diagnosis of Strongyloides stercoralis infection. Clin. Infect. Dis., 33, 1040, 2001.
136. Kishimoto, K. et al., Endoscopic and histopathological study on the duodenum of Strongyloides stercoralis hyperinfection. World J. Gastroenterol., 14, 1768, 2008.
137. Uparanukraw, P., Phongsri, S., and Morakote, N., Fluctuations of larval excretion in Strongyloides stercoralis infection. Am. J. Trop. Med. Hyg., 60, 967, 1999.
138. Steiner, B. et al., Strongyloides stercoralis eggs in a urethral smear after bone marrow transplantation (letter). Clin. Infect. Dis., 34, 1280, 2002.
139. Sato, Y. et al., Ef�cacy of stool examination for detection of Strongyloides infection. Am. J. Trop. Med. Hyg., 53, 248, 1995.
140. de Paula, F.M. et al., Parasitological and immunological diagnoses of strongyloidiasis in immunocompromised and nonimmunocompromised children at Uberlandia, State of Minas Gerais, Brazil. Rev. Inst. Med. Trop. Sao Paulo, 42, 51, 2000.
141. Koga, K. et al., An evaluation of the agar plate method for the detection of Strongyloides stercoralis in northern Thailand. J. Trop. Med. Hyg., 93, 183, 1990.
142. Hirata, T. et al., Increased detection rate of Strongyloides stercoralis by repeated stool examinations using the agar plate culture method. Am. J. Trop. Med. Hyg., 77, 683, 2007.
143. Grapsa, D. et al., Strongyloides stercoralis in a bronchial washing specimen processed as conventional and Thin-Prep smears: report of a case and a review of the literature. Diagn. Cytopathol., 37, 903, 2009.
144. Loutfy, M.R. et al., Serology and eosinophil count in the diagnosis and management of strongyloidiasis in a nonendemic area. Am. J. Trop. Med. Hyg., 66, 749, 2002. 145. Genta, R.M., Predictive value of an enzyme-linked immunosorbent assay (ELISA) for the serodiagnosis of strongyloidiasis. Am. J. Clin. Pathol., 89, 391, 1988. 146. Huaman, M.C. et al., Gelatin particle indirect agglutination and enzyme-linked immunosorbent assay for diagnosis of strongyloidiasis using Strongyloides venezuelensis antigen. Trans. R. Soc. Trop. Med. Hyg., 97, 535, 2003. 147. Lindo, J.F. et al., Prospective evaluation of enzyme-linked immunosorbent assay and immunoblot methods for the diagnosis of endemic Strongyloides stercoralis infection. Am. J. Trop. Med. Hyg., 51, 175, 1994. 148. Melville, L.A., Sykes, A.M., and McCarthy, J.S., The betatubulin genes of two Strongyloides species. Exp. Parasitol., 112, 144, 2006. 149. ten Hove, R.J. et al., Molecular diagnostics of intestinal parasites in returning travellers. Eur. J. Clin. Microbiol. Infect. Dis., 28, 1045, 2009. 150. Hasegawa, H. et al., Hyper-variable regions in 18S rDNA of Strongyloides spp. as markers for species-speci�c diagnosis. Parasitol. Res., 104, 869, 2009. 151. Hasegawa, H. et al., Molecular identi�cation of the causative agent of human strongyloidiasis acquired in Tanzania: Dispersal and diversity of Strongyloides spp. and their hosts. Parasitol. Int., 59, 407, 2010. 152. Taniuchi, M. et al., High throughput multiplex PCR and probe-based detection with Luminex beads for seven intestinal parasites. Am. J. Trop. Med. Hyg., 84, 332, 2011. 153. Basuni, M. et al., A pentaplex real-time polymerase chain reaction assay for detection of four species of soil-transmitted helminths. Am. J. Trop. Med. Hyg., 84, 338, 2011. 154. Kramme, S. et al., Novel real-time PCR for the universal detection of Strongyloides spp. J. Med. Microbiol., 60, 454, 2011. 155. Verweij, J.J. et al., Molecular diagnosis of Strongyloides stercoralis in faecal samples using real-time PCR. Trans. R. Soc. Trop. Med. Hyg., 103, 342, 2009; Erratum: 104, 378, 2010. 64 Chapter 64 - Syphacia
1. Riley, W.A. A mouse oxyurid, Syphacia obvelata, as a parasite of man. J. Parasitol. 6, 89, 1919.
2. Stone, W.B., and Manwell, R.D. Potential helminth infections in humans from pet or laboratory mice and hamsters. Public Health Rep. 81, 647, 1966.
3. Seurat, L.G. Sur les Oxyures de Mammifères, Comptes Rendus de la Société Biologique (Paris) 79, 64, 1916.
4. Committee on Infectious Diseases of Rats and Mice, National Research Council, Infectious Disease of Mice and Rats, National Academy Press, Washington, DC, 156p, 1991.
5. Grice, R.L., and Prociv, P. In vitro embryonation of Syphacia obvelata eggs. Int. J. Parasitol. 23, 257, 1993.
6. Petter, A.J., and Quentin, J-D. Keys to genera of the Oxyuroidea. In: Anderson, R.C., Chabaud, A.G., and Willmott, S. (eds), CIH Keys to the Nematode Parasites of Vertebrates. Commonwealth Agricultural Bureaux, No. 4, 1976.
7. Flynn, R.J. Parasites of Laboratory Animals. Iowa State Press, Ames, IA, p. 236, 1973.
8. Taffs, L.F. Pinworm infections in laboratory rodents: A review. Lab. Anim. 10, 1, 1976.
9. Blaxter, M.L. et al. A molecular evolutionary framework for the phylum Nematoda. Nature 392, 71, 1998.
10. Nadler, S.A. et al. Molecular phylogeny of clade III nematodes reveals multiple origins of tissue parasitism. Parasitology 134, 1421, 2007.
11. Feldman, S.H., and Bowman, S.G. Molecular phylogeny of the pinworms of mice, rats and rabbits, and its use to develop molecular beacon assays for the detection of pinworms in mice. Lab. Anim. 36, 1, 2007.
12. Okamakoto, M. et al. Phylogenetic relationships of rodent pinworms (genus Syphacia) in Japan inferred from mitochondrial CO1 gene sequences. J. Vet. Med. Sci. 69, 545, 2007. 13. Wescott, R.B., Malczewski, A., and Van Hoosier, G.L. The in�uence of �lter top caging on transmission of pinworm infections in mice. Lab. Anim. Sci. 26, 742, 1976. 14. Marillier, R.G. et al. IL-4/IL-13 independent goblet cell hyperplasia in experimental helminth infections. BMC Immunol. 9, 11, 2008. 15. Bugarski, D. et al. Hematopoietic changes and altered reactivity to IL-17 in Syphacia obvelata-infected mice. Parasitol. Int. 55, 91, 2006. 16. Stewart, P.W., and Chapes, S.K. Role of major histo compatibility complex class II in resistance of mice to naturally acquired infection with Syphacia obvelata. Comp. Med. 53, 70, 2003. 17. Michels, C. et al. Infection with Syphacia obvelata (pinworm) induces protective Th2 immune responses and in�uences ovalbumin-induced allergic reactions. Infect. Immun. 74, 5926, 2006. 18. Fallon, P.G., and Mangan, N.E. Suppression of the T h 2-type allergic reactions by helminth infection. Nat. Rev. Microbiol. 7, 220, 2007. 19. Tung, K.S.K. et al. Regulatory T-cell, endogenous antigen and neonatal environment in the prevent and induction of autoimmune disease. Immunol. Rev. 182, 135, 2001. 20. Clarke, C.L., and Perdue, K.A. Detection and clearance of Syphacia obvelata infection in Swiss Webster and athymic nude mice. Contemp. Top. Lab. Anim. Sci. 34, 9, 2004. 21. Hill, W.A., Randolph, M.M., and Mandrell, T.D. Sensitivity of perianal tape impressions to diagnose pinworm (Syphacia spp.) infections in rats (Rattus norvegicus) and mice (Mus musculus). J. Am Assoc. Lab. Anim. Sci. 48, 378, 2009. 22. Goncalves, L. et al. Helminth parasites of conventionally maintained laboratory mice—II. Inbred strains with an adaptation of the anal swab technique. Mem. Inst. Oswaldo Cruz, Rio De Janieiro, 93, 121, 1998. 23. Parel, J.D.C., Galula, J.U., and Ooi, H.-K. Characterization of rDNA sequences from Syphacia obvelata, Syphacia muris, and Aspiculuris tetraptera and development of a PCR-based method for identi�cation. Vet. Parasitol. 153, 379, 2008. 24. Jobet, E. et al. Use of random ampli�ed polymorphic DNA (RAPD) for generating speci�c DNA probes for oxyuroid species (Nematoda). Parasite 5, 47, 1998. 25. Traub, T.K.J. et al. Application of a species-speci�c PCRRFLP to identify Ancylostoma eggs directly from canine faeces. Vet. Parasitol. 123, 245, 2004. 26. de Gruijter, J.M. et al. Polymerase chain reaction-based differential diagnosis of Ancylostoma duodenale and Necator americanus infections in humans in northern Ghana. Trop. Med. Int. Health 10, 574, 2005. 27. Verweij, J.J. et al. PCR assay for the speci�c ampli�cation of Oesophagostomum bifurcum DNA from human faeces. Int. J. Parasitol. 30, 137, 2000. 28. Rinaldi, L., Russo, T., and Schioppi, M. Passalurus ambiguus: New insights into the copromicroscopic diagnosis and circadian rhythm of egg excretion. Parasitol. Res. 101, 557, 2007. 29. Ellis, R.E., Sulston, J.E., and Coulson, A.R. The rDNA of E. elegans: Sequence and structure. Nucleic Acids Res. 14, 2345, 1986. 65 Chapter 65 - Thelazia
12. Anderson, R.C. A study of Filaria martis Gmelin 1790, from Martes foina and Pedetes caffer. Can. J. Zool. 38, 157–167, 1960.
13. Chabaud, A.G., and Bain, O. The evolutionary expansion of the Spirurida. Int. J. Parasitol. 24, 1179–1201, 1994.
14. Otranto, D. et al. Thelazia callipaeda (Spirurida, Thelaziidae) of carnivores and humans: Morphological study by light and scanning electron microscopy. Parassitologia 45, 125–133, 2003.
15. Otranto, D. et al. Nematode biology and larval development of Thelazia callipaeda (Spirurida, Thelaziidae) in the drosophilid intermediate host in Europe and China. Parasitology 131, 847–855, 2005.
16. Shi, Y.E. et al. Thelazia callipaeda (Nematoda: Spirurida): Transmission by �ies from dogs to children in Hubei, China. Trans. R. Soc. Trop. Med. Hyg. 82, 627, 1988.
17. Otranto, D. et al. Musca domestica is not a vector of Thelazia callipaeda in experimental or natural conditions. Med. Vet. Entomol. 19, 1–5, 2005.
18. Kozlov, D.P. Study on the biology of Thelazia callipaeda. Papers of Helminthologic Laboratory. Academy of Sciences USSR 142, 732–733, 1963.
19. Wang, Z.X., Shen, J.L., and Wang, K.C. Further studies on the life cycle if Thelazia callipaeda in China. Chin. J. Parasitol. Paras. Dis. 20, 328–331, 2002.
20. Otranto, D. et al. Phortica variegata is an intermediate host of Thelazia callipaeda under natural conditions: Evidence for pathogen transmission by a male arthropod vector. Int. J. Parasitol. 36, 1167–1173, 2006.
21. Otranto, D. et al. Biology of Thelazia callipaeda (Spirurida, Thelaziidae) eyeworms in naturally infected de�nitive hosts. Parasitology 129, 627–633, 2004.
22. Rossi, L. and Bertaglia, P. Presence of Thelazia callipaeda Raillet & Henry, 1910 in Piedmont, Italy. Parassitologia 31, 167–172, 1989.
23. Di Sacco, B., Ciocca, A., and Sirtori, G. Thelazia callipaeda (Railliet and Henry, 1910) nel sacco congiuntivale di un gatto di Milano. Veterinaria 4, 81–84, 1995.
24. Lia, R. et al. First report of Thelazia callipaeda in southern Italy- Basilicata region. Proceedings of the Eight Conference of the European Federation of Parasitologist. Acta Parasitol. 45, 178, 2000.
25. Otranto, D. et al. Current status and epidemiological observations of Thelazia callipaeda (Spirurida, Thelaziidae) in dogs, cats and foxes in Italy: A “coincidence” or a parasitic disease of the Old Continent? Vet. Parasitol. 116, 315–325, 2003.
26. Rossi, L. et al. Thelazia callipaeda in foxes from North-West Italy. Parassitologia 44, 159, 2002.
27. Otranto, D., Cantacessi, C., and Mallia, E. First report of Thelazia callipaeda (Spirurida, Thelaziidae) in wolves (Canis lupus) in Italy. J. Wildl. Dis. 43, 508–511, 2007.
28. Chermette, R., Guillot, J., and Bussiéras, J. Canine ocular thelaziosis in Europe. Vet. Rec. 154, 248, 2004.
29. Hermosilla, C., Herrmann, B., and Bauer, C. First case of Thelazia callipaeda infection in a dog in Germany. Vet. Rec. 154, 568–569, 2004.
30. Dorchies, Ph. et al. First reports of autochthonous eyeworm infection by Thelazia callipaeda (Spirurida, Thelaziidae) in dogs and cat from France. Vet. Parasitol. 149, 294–297, 2007.
31. Malacrida, F. et al. Emergence of canine ocular Thelaziosis caused by Thelazia callipaeda in southern Switzerland. Vet. Parasitol. 157(3–4), 321–327, 2008. 32. Otranto, D. et al. Thelazia callipaeda (Spirurida, Thelaziidae) in wild animals: Report of new host species and ecological implications. Vet. Parasitol. 166, 262–267, 2009. 33. Herr, J., Schley, L., and Roper, T.J. Socio-spatial organization of urban stone martens. J. Zool. 277, 54–62, 2009. 34. Shen, J.L. et al. Human thelaziosis—A neglected parasitic disease of the eye. J. Parasitol. 92, 186–190, 2006. 35. Stuckey, E.J. Circumocular �lariasis. Chin. Med. J. 31, 24, 1917. 36. Miroshnichenko, V.A. et al. A case of ocular thelaziasis in a 3-year-old child. Vestnik Oftalmologii 104, 64, 1988 [In Russian]. 37. Chen, C.C., and Zhang, Y.M. A case report of human thelaziosis in the eye anterior chamber. Chin. J. Ophthalm. 4, 466, 1954. 38. Min, S., Jae, R.Y., and Hyun Y.P. Enzooticity of the dogs, the reservoir host of Thelazia callipaeda, in Korea. Kor. J. Parasitol. 40, 101–103, 2002. 39. Koyama, Y. et al. Five cases of thelaziasis. Br. J. Ophthalmol. 84, 441, 2000. 40. Yospaiboon, Y. et al. Ocular thelaziasis in Thailand: A case report. J. Med. Assoc. Thail. 72, 469–473, 1989. 41. Wang, Z.X., Shen, J.L., and Du, J.S. Studies on the relationship between alternative generation of house dogs and control of human thelaziosis. J. Trop. Dis. Parasitol. 1, 204–207, 2003. 42. Shen, Y.P. et al. Human infections with Thelazia callipaeda in China. Jiangsu Med. 4, 41–42, 1979 [In Chinese]. 43. Zhonghua, Y.Z. Analysis of 83 cases of human thelaziosis in China. Zhonghua Yan Ke Za Zhi (Chin. J. Ophthalmol.) 16, 64–67, 1980 [In Chinese]. 44. Jiang, Z.X., Xu, L.Q., and Yu, L.Q. Human thelaziasis in China. Zhongguo Ji Sheng Chong Bing Fang Zhi Za Zhi (Chin. J. Parasitic Dis. Control) 4, 48–51, 1991 [In Chinese]. 45. Otranto, D., and Dutto, M. Human thelaziasis, Europe. Emerg. Infect. Dis. 4, 647–649, 2008. 46. Wang, Z.X., and Yang, Z.X. Studies on the biology of Thelazia callipaeda and its experimental pathogenic observations on rabbits. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi (Chinese J. Parasitol. Parasitic Dis.) 3, 128–130, 1985 [In Chinese]. 47. Wang, K.C., Wang, Z.X., and Shen, J.L. Canine infection with Thelazia callipaeda and human thelaziosis. J. Trop. Dis. Parasitol. 28, 216–218, 1999 [In Chinese]. 48. Zakir, R. et al. Intraocular infestation with the worm, Thelazia callipaeda. Br. J. Ophthalmol. 83, 1994, 1999. 49. Ohira, A. Five cases of thelaziasis. Br. J. Ophthalmol. 84, 439, 2000. 50. Smith, J. Bacterial conjunctivitis. Clin. Evid. 12, 926–932, 2004. 51. Rossi, L., and Peruccio, C. Thelaziosi oculare nel cane: Aspetti clinici e terapeutici. Veterinaria, 2, 47–50, 1989 [In Italian, with abstract in English]. 52. Lia, R.P. et al. Field ef�cacy of moxidectin 1 per cent against Thelazia callipaeda in naturally infected dogs. Vet. Rec. 154(5), 143–145, 2004. 53. Bianciardi, P., and Otranto, D. Treatment of dog thelaziosis caused by Thelazia callipaeda (Spirurida, Thelaziidae) using a topical formulation of imidacloprid 10% and moxidectin 2.5%. Vet. Parasitol. 129(1–2), 89–93, 2005. 54. Rossi, L. et al. Use of sustained-release moxidectin to prevent eyeworm (Thelazia callipaeda) infection in dogs. Vet. Rec. 161, 820–821, 2007. 55. Otranto, D., and Traversa, D. Molecular characterization of the �rst internal transcribed spacer of ribosomal DNA of the most common species of eyeworms (Thelazioidea: Thelazia). J. Parasitol. 90, 185–188, 2004. 56. Otranto, D. et al. Analysis of genetic variability within Thelazia callipaeda (Nematoda: Thelazioidea) from Europe and Asia by sequencing and mutation scanning of mitochondrial cytochrome c oxidase subunit 1. Mol. Cell. Probes 19, 306–313, 2005.
57. Yamaguti, S. The Nematodes of Vertebrates Part I. Systema Helminthum. Vol. III. Interscience Publisher, Inc., New York, USA, pp. 634–636, 1961.
58. Price, E.W. A new nematode parasitic in the eyes of dogs in the United States. J. Parasitol. 17, 112–113, 1930.
59. Price, E.W. A note on the occurrences of eye worms in dogs in the United States. North. Am. Vet. 41, 49–58, 1931.
60. Gasser, R.B. et al. Single-strand conformation polymorphism (SSCP) for the analysis of genetic variation. Nat. Prot. 1, 3121–3128, 2006.
61. Okada, T. Bleeding sap preference of the Drosophilid �ies. Jpn. J. Appl. Entomol. Zool. 3, 216–229, 1962. 62. Kimura, M.T. et al. Breeding sites of Drosophilid �ies in and near Sapporo, Northern Japan, with supplementary notes on adult feeding habits. Kontyû 45, 571–582, 1977. 63. Máca, J. Revision of Palearctic species of Amiota subg. Phortica (Diptera, Drosophilidae). Acta Entomol. Bohemoslovaca 74, 115–130, 1977. 64. Jin, D. Parasites of human eyes in China. Zhonghua Yan Ke Za Zhi (Chin. J. Ophthalmol.) 7, 227, 1957 [In Chinese]. 65. Otranto, D., Brianti, E., Cantacessi, C., Lia, R. P., and Máca, J. The zoophilic fruit�y Phortica variegata: Morphology, ecology and biological niche. Medical and Veterinary Entomology 20, 358–364, 2006. 66 Chapter 66 - Toxocara
3. Iddawela, D.R. et al. A seroepidemiological study of toxocariasis and risk factors for infection in children in Sri Lanka. Southeast Asian J. Trop. Med. Public Health 34, 7, 2003.
4. Prociv, P. Observations on the transmission and development of Toxocara pteropodis (Ascaridoidea: Nematoda) in the Australian grey-headed �ying-fox, Pteropus poliocephalus (Pteropodidae: Megachiroptera). Z. Parasitenkd. 69, 773, 1983.
5. Glickman, L.T., Schantz, P.M., and Cypess, R.H. Canine and human toxocariasis: Review of transmission, pathogenesis and clinical disease. J. Am. Vet. Med. Ass. 175, 1265, 1979.
6. Brunanska, M., Dubinsky, P., and Reiterova, K. Toxocara canis: Ultrastructural aspects of larval moulting in the maturing eggs. Int. J. Parasitol. 25, 683, 1995.
7. Borgsteede, F.H.M. Heminth parasites of wild foxes (Vulpes vulpes) in the Netherlands. Z. Parasitenkd. 70, 527, 1984.
8. Smith, G.C. et al. Prevalence of zoonotic important parasites in the red fox (Vulpes vulpes) in Great Britain. Vet. Parasitol. 118, 133, 2003.
9. Dubná, S. et al. Contamination of soil with Toxocara eggs in urban (Praque) and rural areas in the Czech Republic. Vet. Parasitol. 144, 81, 2007.
10. Düwel, D. The prevalence of Toxocara eggs in the sand in children’s playgrounds in Frankfurt/M. Ann. Trop. Med. Parasitol. 78, 633, 1984.
11. Emehelu, C.O., and Fakae, B.B. Prevalence of Toxocara canis ova on playgrounds of nursery schools in Nsukka, Nigeria. Int. J. Zoonoses 13, 158, 1986.
12. Gawor, J. et al. Environmental and personal risk factors for toxocariasis in children diagnosed disease in urban and rural areas of central Poland. Vet. Parasitol. 155, 217, 2008.
13. Holland, C. et al. Families, parks, gardens and toxocariasis. Scand. J. Infect. Dis. 23, 225, 1991. 14. Ruiz de Ybáñez, M.R., Garijo, M.M., and Alonso, F.D. Prevalence and viability of eggs of Toxocara spp. and Toxascaris leonina in public parks in eastern Spain. J. Helminthol. 75, 169, 2001.
15. Uga, S. Prevalence of Toxocara eggs and number of faecal deposits from dogs and cats in sandpits of public parks in Japan. J. Helminthol. 67, 78, 1993.
16. Fillaux, J. et al. Epidemiology of toxocariasis in a steppe environment: The Patagonia study. Am. J. Trop. Med. Hyg. 76, 1144, 2007.
17. Magnaval, J.-F. et al. Highlights of human toxocariasis. Kor. J. Parasitol. 39, 1, 2001.
18. Kozan, E. et al. Prevalence of helminth eggs on raw vegetables used for salads. Food Contr. 16, 239, 2005.
19. Gaspard, P., Wiart, J., and Schwartzbrod, J. Parasitological contamination of urban sludge used for agricultural purposes. Waste Manag. Res. 15, 429, 1997.
20. Moorhouse, D.E. Toxocariasis—A possible cause of the Palm Island mystery disease. Med. J. Aust. 1, 172, 1982.
21. Wolfe, A., and Wright, I.P. Human toxocariasis and direct contact with dogs. Vet. Rec. 152, 419, 2003.
22. Hoffmeister, B. et al. Cerebral toxocariasis after consumption of raw duck liver. Am. J. Trop. Med. Hyg. 76, 600, 2007.
23. Lee, K.-T. et al. Studies on the including possibility of human visceral larva migrans associated with eating habit of raw liver of domestic animals. Kor. J. Parasitol. 14, 51, 1976.
24. Stürchler, D. Weiss, N., and Gassner, M. Transmission of toxocariasis. J. Infect. Dis. 162, 571, 1990.
25. Umeche, N., and Mandah, L.E. Musca domestica as carrier of intestinal helminths in Calabor, Nigeria. East Afr. Med. J. 66, 349, 1989. 26. Magnaval, J.-F., Glickman, L.T., and Dorchies, Ph. La toxocarose, une zoonose helminthique majeure. Rev. Med. Vet. 145, 611, 1994). 27. Chomel, B.B. et al. Serosurvey of some major zoonotic infections in children and teenagers in Bali, Indonesia. Southeast Asian J. Trop. Med. Public Health 24, 321, 1993. 28. Magnaval, J.F. et al. Epidemiology of human toxocariasis in La Réunion. Trans. R. Soc. Trop. Med. Hyg. 88, 531, 1994. 29. Pawłowski, Z. Toxocariasis in humans: Clinical expression and treatment dilemma. J. Helminthol. 75, 299, 2001. 30. Wilder, H.C. Nematode endophthalmitis. Trans. Am. Acad. Ophthalmol. Otolaryngol. 55, 99, 1950. 31. Nichols, R.L. The etiology of visceral larva migrans. J. Parasitol. 42, 349, 1956. 32. Magnaval, J.F. et al. Human Toxocara infection of the central nervous system and neurological disorders: A case–control study. Parasitology 115, 537, 1997. 33. Bass, J.L. et al. Asymptomatic toxocariasis in children. Clin. Pediatr. 26, 441, 1987. 34. Baldisserotto, M. et al. Ultrasound �ndings in children with toxocariasis: Report on 18 cases. Pediatr. Radiol. 29, 316, 1999. 35. Vidal, J., Sztajnbok, J., and Seguro, A.C. Meningoencephalitis due to Toxocara canis: Case report and review of the literature. Am. J. Trop. Med. Hyg. 69, 341, 2003. 36. Kennedy, J.J., and Defeo, E. Ocular toxocariasis demonstrated by ultrasound. Ann. Ophthalmol. 13, 1357, 1981. 37. Hossack, J. et al. A case of adult hepatic toxocariasis. Natl. Clin. Pract. Gastroenterol. Hepatol. 5, 344, 2008. 38. Magnaval, J.-F. et al. Eosinophil cationic protein as a possible marker of active human Toxocara infection. Allergy 56, 1096, 2001. 39. Smith, H. et al. How common is human toxocariasis? Towards standardizing our knowledge. Trends Parasitol. 25, 182, 2009. 40. De Savigny, D.H. In vitro maintenance of Toxocara canis larvae and a simple method for the production of Toxocara ES antigen for use in serodiagnostic tests for visceral larva migrans. J. Parasitol. 61, 781, 1979. 41. Badley, J.E. et al. Analysis of Toxocara canis larval excretory– secretory antigens: Physicochemical characterization ant antibody recognition. J. Parasitol. 73, 593, 1987. 42. Magnaval, J.F. et al. Application of the western blotting procedure for immunodiagnosis of human toxocariasis. Parasitol. Res. 77, 697, 1991. 43. Fong, M.-Y., and Lau, Y.L. Recombinant expression of the larval excretory-secretory antigen TES-120 of Toxocara canis in the methylotrophic yeast Pichia pastoris. Parasitol. Res. 92, 173, 2004. 44. Yamasaki, H. et al. Development of a highly speci�c recombinant Toxocara canis second-stage larva excretory– secretory antigen for immunodiagnosis of human toxocariasis. J. Clin. Microbiol. 38, 1409, 2000. 45. Obwaller, A. et al. Toxocara infestations in humans: Symptomatic course of toxocarosis correlates signi�cantly with levels of IgE/anti-IgE immune complexes. Parasite Immunol. 20, 311, 1998. 46. Hübner, J., Uhlikova, M., and Leissova, M. Diagnosis of the early phase of larval toxocariasis using IgG avidity. Epidemiol. Mikrobiol. Immunol. 50, 67, 2001. 47. Park, S.-P. et al. Five cases of ocular toxocariasis con�rmed by serology. Kor. J. Parasitol. 38, 267, 2000. 48. Tinneli, M. et al. RAST and ELISA in three autochthonous parasitic infections in Italy: Toxocariasis, hydatidosis and fascioliasis. Boll. Ist. Sieroter Milan 66, 38, 1987.
49. Smith, H.V. et al. A paper radioimmunosorbent test (PRIST) for the detection of larva-speci�c antibodies to Toxocara canis in human sera. J. Immunol. Methods 37, 47, 1980.
50. Khalil, H.M. et al. Immunodiagnostic of visceral toxocarias. J. Egypt. Soc. Parasitol. 19, 381, 1989.
51. Glickaman, L.T. et al. Evaluation of serodiagnostic tests for visceral larva migrans. Am. J. Trop. Med. Hyg. 27, 492, 1978.
52. Nagakura, K. et al. Serologic differentiation between Toxocara canis and Toxocara cati. J. Infect. Dis. 162, 1227, 1980.
53. Huntley, C.C., Costas, M.C., and Lyerly, A. Visceral larva migrans syndrome: Clinical characteristics and immunologic studies in 51 patients. Pediatrics 36, 523, 1965.
54. Krupp, I.M. Hemagglutination test for the detection of antibodies speci�c for Ascaris and Toxocara antigens in patients with suspected visceral larva migrans. Am. J. Trop. Med. Hyg. 23, 378, 1974.
55. Bisseru, B., and Woodruff, A.W. The detection of circulating antibody in human toxocara infections using the indirect �uorescent antibody test. J. Clin. Pathol. 21, 449, 1968.
56. Matthes, H.F., and Buchwalder, R. The larva precipitation test with Ascaris suum and Toxocara canis larvae. Angew. Parasitol. 23, 1, 1982.
57. Woodruff, A.W. Toxocariasis. Br. Med. J. 3, 663, 1970.
58. Nadler, S.A. Advantages and disadvantages of molecular phylogenetics: A case study of ascaridoid nematodes. J. Nematol. 27, 423, 1995.
59. Nadler, S.A. Biochemical polymorphism in Parascaris equorum, Toxocara canis and Toxocara cati. Mol. Biochem. Parasitol. 18, 45, 1986.
60. Turcˇeková, L’., and Dubinský, P. Differentiation between Toxocara canis and T. cati using restriction pro�les and ribosomal gene probe. Helminthologia 33, 223, 1996.
61. Wu, Z. et al. Primers for polymerase chain reaction to detect genomic DNA of Toxocara canis and T. cati. J. Helminthol. 71, 77, 1997.
62. Jacobs, D.E. et al. PCR-based methods for identi�cation of potentially zoonotic ascaridoid parasites of the dog, fox and cat. Acta Trop. 68, 191, 1997.
64. Epe, C. et al. Transmission trials, ITS2-PCR and PAPR-PCR show identity of Toxocara canis isolates from red fox and dog. Vet. Parasitol. 84, 101, 1999. 65. Li, M.-W. et al. The occurence of Toxocara malaysiensis in cats in China, con�rmed by sequence-based analyses of ribosomal DNA. Parasitol. Res. 99, 554, 2006. 65. Zhu, X.Q. et al. Molecular characterization of a Toxocara variant from cats in Kuala Lumpur, Malaysia. Parasitology 117, 155, 1998. 66. Jex, A.R. et al. The mitochondrial genome of Toxocara canis. Plos Negl. Trop. Dis. 2, e273, 2008. 67. Li, M.-W. et al. The complete mitochodrial genomes for three Toxocara species of human and animal health signi�cance. BMC Genomics 9, 224, 2005. 68. Mizgajska, H., and Rejmenciak, A. Differentiation of eggs of Toxocara canis and Toxocara cati—Parasites of dog and cat. Wiad. Parazyt. 43, 435, 1997. 69. Borecka, A. Differentiation of Toxocara spp eggs isolated from the soil by the PCR-linked RFLP method. Helminthologia 41, 185, 2004. 70. Fogt-Wyrwas, R., Jarosz, W., and Mizgajska-Wiktor, H. Utilizing a polymerase reaction method for the detection of Toxocara canis and T. cati eggs in soil. J. Helminthol. 81, 75, 2007. 71. Borecka, A., and Gawor, J. Modi�cation of gDNA extraction from soil for PCR designed for the routine examination of soil contaminated with Toxocara spp. eggs. J. Helminthol. 82, 119, 2008. 72. Rai, S.K. et al. Use of polymerase chain reaction in the diagnosis of toxocariasis: An experimental study. Southeast Asian J. Trop. Med. Public Health 28, 541, 1997. 73. Borecka, A. et al. Detection of Toxocara canis larvae by PCR in the liver of experimentally infected Mongolian gerbils (Meriones unguiculatus). Helminthologia 45, 147, 2008. 74. Ishiwata, K. et al. Identi�cation of tissue-embedded ascarid larvae by ribosomal DNA sequencing. Parasitol. Res. 92, 50, 2004. 75. Moreira, D. Ef�cient removal of PCR inhibitors using agarose-embedded DNA preparations. Nucleic Acids Res. 26, 3309, 1998. 76. Yeates, C. et al. Methods for microbial DNA extraction from soil for PCR ampli�cation. Biol. Proced. Online 1, 40, 1998. 67 Chapter 67 - Trichinella
1. Pozio, E., and Murrell, K.D. Systematics and epidemiology of Trichinella. Adv. Parasitol. 63, 367, 2006.
2. Pozio, E. Taxonomy, biology and epidemiology of Trichinella parasites. In: Dupouy-Camet, J., and Murrell, K.D. (eds.), FAO/WHO/OIE Guidelines for the Surveillance, Management, Prevention and Control of Trichinellosis, Chapter 1. World Organisation for Animal Health, Paris, 2007. 3. Zarlenga, D.S. et al. Post-Miocene expansion, colonization, and host switching drove speciation among extant nematodes of the archaic genus Trichinella. Proc. Natl. Acad. Sci. USA. 103, 7354, 2006. 4. Pozio, E., and Zarlenga, D.S. Recent advances on the taxonomy, systematics and epidemiology of Trichinella. Int. J. Parasitol. 35, 1191, 2005. 5. Pozio, E. et al. Taxonomic revision of the genus Trichinella. J. Parasitol. 78, 654, 1992. 6. Bruschi, F. et al. Anaphylactic response to parasite antigens: IgE and IgG1 independently induce death in Trichinellainfected mice. Int. Arch. Allergy. Immunol. 119, 291, 1999. 7. Gomez Morales, M.A. et al. Increased CD8 + -T-cell expression and a type 2 cytokine pattern during the muscular phase of Trichinella infection in humans. Inf. Immun. 70, 233, 2002. 8. Forbes, L.B. Zoonotic aspects of trichinellosis. In: Rhode, K. (ed.), Marine Parasitology, Chapter 11. CSIRO, Melbourne, 2005. 9. Dick, T.A., and Pozio, E. Trichinella spp. and trichinellosis. In: Samuel, W.M., Pybus, M.J., and Kocan, A.A. (eds), Parasitic Diseases of Wild Mammals, Chapter 15. Iowa State University Press, Ames, 2001. 10. Serhir, B. et al. Outbreak of trichinellosis associated with arctic walruses in northern Canada. Can. Com. Dis. Rep. 27, 31, 2001. 11. Schellenberg, R.S. et al. An outbreak of trichinellosis due to consumption of bear meat infected with Trichinella nativa, in 2 northern Saskatchewan communities. J. Infec. Dis. 188, 835, 2003. 12. Nelson, M. et al. A common-source outbreak of trichinosis from consumption of bear meat. J. Environ. Health 65, 16, 2003. 13. Ancelle, T. et al. Outbreak of trichinellosis due to consumption of bear meat from Canada, France, September 2005. Eurosurveillance 10, 10, 2005. 14. La Rosa, G. et al. Molecular identi�cation of natural hybrids between Trichinella nativa and Trichinella T6 provides evidence of gene �ow and ongoing genetic divergence. Int. J. Parasitol. 33, 209, 2003. 15. Gari-Toussaint, M. et al. Human trichinellosis due to Trichinella britovi in southern France after consumption of frozen wild boar meat. Eur. Surveill. 10, 117, 2005. 16. Rodriguez de las Parras, E. et al. Trichinellosis outbreaks in Spain (1990–2001). Enferm. Infec. Microbiol. Clin. 22, 70, 2004. 17. Ozdemir, D. et al. Acute trichinellosis in children compared with adults. Pediatric. Infect. Dis. J. 24, 897, 2005. 18. Nezri, M. et al. Premiere observation d’un cas de humain de trichinellose a Trichinella britovi en Algerie après consummation de viande de chacal (Canis aureus). Bull. Soc. Pathol. Exotique 99, 94, 2006. 19. Pozio, E. et al. Human outbreak of trichinellosis in the Mediterranean island of Sardinia, Italy. Vet. Parasitol. 140, 177, 2006. 20. Pozio, E., Gomez Morales, M.A., and Dupouy Camet, J. Clinical aspects, diagnosis and treatment of trichinellosis. Exp. Rev. Anti-infect. Ther. 1, 471, 2003. 21. Marucci, G. et al. Trichinella nelsoni and Trichinella T8 mixed infection in a lion (Panthera leo) of the Kruger National Park (South Africa). Vet. Parasitol. 159, 225, 2009. 22. Kapel, C.M.O., and Gamble, H.R. Infectivity, persistence, and antibody response to domestic and sylvatic Trichinella spp. in experimentally infected pigs. Int. J. Parasitol. 30, 215, 2000.
23. Kanai, Y. et al. Epizootiological survey of Trichinella spp. infection in carnivores, rodents and insectivores in Hokkaido, Japan. Jpn. J. Vet. Res. 54, 175, 2007.
24. Pozio, E. et al. Trichinella britovi etiological agent of sylvatic trichinellosis in the Republic of Guinea (West Africa) and a re-evaluation of geographical distribution for encapsulated species in Africa. Int. J. Parasitol. 35, 955, 2005.
25. Forrester, A.T. et al. The �rst record of an outbreak of trichinosis in Africa south of the Sahara. Trans. Roy. Soc. Trop. Med. Hyg. 55, 503, 1961.
26. Bura, M.W., and Willett, W.C. An outbreak of trichinosis in Tanzania. East African Med. J. 54, 185, 1977.
27. Krivokapich, S. et al. Molecular evidence for a novel encapsulated genotype of Trichinella from Patagonia, Argentina, Abstract book, p. 13, 12th International Conference on Trichinellosis, National Park Plitvice Lakes, Croatia, pp. 25–30, September, 2007.
28. La Rosa, G. et al. Trichinella pseudospiralis populations of the Palearctic region and their relationship with populations of the Nearctic and Australian regions. Int. J. Parasitol. 31, 297, 2001.
29. Pozio, E. The broad spectrum of Trichinella hosts: From cold- to warm-blooded animals. Vet. Parasitol. 132, 3, 2005.
30. Andrews J.R.H. et al. Identi�cation of Trichinella pseudospiralis from a human case using random ampli�ed polymorphic DNA. Am. J. Trop. Med. Hyg. 53, 185, 1995.
31. Britov, V.A. Trichinellosis in Kamchatka. Wiad. Parazytolog 43, 287, 1997.
32. Jongwutiwes, S. et al. First outbreak of human trichinellosis caused by Trichinella pseudospiralis. Clin. Infect. Dis. 26, 111, 1998.
33. Ranque, S. et al. Trichinella pseudospiralis outbreak in France. Em. Infect. Dis. 6, 543, 2000.
34. Pozio, E. et al. Trichinella papuae n. sp. (Nematoda), a new non-encapsulated species from domestic and sylvatic swine of Papua New Guinea. Int. J. Parasitol. 29, 1825, 1999.
35. Pozio, E. et al. Inappropriate feeding practice favors the transmission of Trichinella papuae from wild pigs to saltwater crocodiles in Papua New Guinea. Vet. Parasitol. 127, 245, 2005.
36. Kusolsuk, T. et al. Trichinella papuae in Ban-Rai district, Uthaithani province, Thailand, Abstract book, p. 47, 5th Food and Waterborne Parasitic Zoonoses, Bangkok, Thailand, pp. 28–30, November, 2006.
37. Owen, I.L. et al. Trichinella infection in a hunting population of Papua New Guinea suggests an ancient relationship of Trichinella with human beings. Trans. Roy. Soc. Trop. Med. Hyg. 99, 618, 2005.
38. Pozio, E. et al. Trichinella zimbabwensis n.sp. (Nematoda), a new non-encapsulated species from crocodiles (Crocodylus niloticus) in Zimbabwe also infecting mammals. Int. J. Parasitol. 32, 1787, 2002.
39. Pozio, E. et al. Trichinella zimbabwensis in wild reptiles of Zimbabwe and Mozambique and in farmed reptiles of Ethiopia. Vet. Parasitol. 143, 305, 2007.
40. Pozio, E. et al. Trichinellosis in the European Union: Epidemiology, ecology and economic impact. Parasitol. Today 14, 35, 1998.
41. Pozio, E. et al. Hunting practices increase the prevalence of Trichinella infection in wolves from European Russia. J. Parasitol. 87, 1498, 2001.
42. Dupouy-Camet, J. et al. Opinion on the diagnosis and treatment of human trichinellosis. Exp. Opinion Pharmacother. 3, 1117, 2002. 43. Dupouy-Camet, J., and Bruschi, F. Management and diagnosis of human trichinellosis. In: Dupouy-Camet, J., and Murrell, K.D. (eds), FAO/WHO/OIE Guidelines for the Surveillance, Management, Prevention and Control of Trichinellosis, Chapter 2. World Organisation for Animal Health, Paris, 2007. 44. Kociecka, W., Trichinellosis: Human disease, diagnosis and treatment. Vet. Parasitol. 93, 365, 2000. 45. Capó, V.A., Despommier, D.D., and Polvere, R.I. Trichinella spiralis: Vascular endothelial growth factor is up-regulated within the nurse cell during the early phase of its formation. J. Parasitol. 84, 209, 1998. 46. Dick, T.A. et al. The use of the polymerase chain reaction to identify porcine isolates of Trichinella. J. Parasitol. 78, 145, 1992. 47. Soulé, C. et al. Differentiation of Trichinella isolates by polymerase chain reaction. Parasitol. Res. 79, 461, 1993. 48. Bandi, C. et al. Random ampli�ed polymorphic DNA technique for the identi�cation of Trichinella species. Parasitology 107, 419, 1993. 49. Bandi, C. et al. Random ampli�ed polymorphic DNA �ngerprints of the eight taxa of Trichinella and their comparison with allozyme analysis. Parasitology 110, 401, 1995. 50. Wang, H. et al. Restriction fragment length polymorphism (RFLP) analysis of genomic DNA of 5 strains of Trichinella spiralis in China. Chin. Med. Sci. J. 10, 131, 1995. 51. Gasser, R.B. et al. PCR-SSCP of rDNA for the identi�cation of Trichinella isolates from mainland China. Mol. Cell. Probes 12, 27, 1998. 52. Gasser, R.B. et al. Nonisotopic single-strand conformation polymorphism analysis of sequence variability in ribosomal DNA expansion segments within the genus Trichinella (Nematoda: Adenophorea). Electrophoresis 25, 3357, 2004. 53. Wu, Z. et al. Polymerase chain reaction primers to identify Trichinella spiralis or T. pseudospiralis. Parasitol. Int. 46, 149, 1997. 54. Wu, Z., Nagano, I., and Takahashi, Y. The detection of Trichinella with polymerase chain reaction (PCR) primers constructed using sequences of random ampli�ed polymorphic DNA (RAPD) or sequences of complementary DNA encoding excretory-secretory (E–S) glycoproteins. Parasitology 117, 173, 1998. 55. Nagano, I. et al. Identi�cation of Trichinella isolates by polymerase chain reaction—Restriction fragment length polymorphism of the mitochondrial cytochrome c-oxidase subunit I gene. Int. J. Parasitol. 29, 1113, 1999. 56. Appleyard, G.D. et al. Differentiation of Trichinella genotypes by polymerase chain reaction using sequence-speci�c primers. J. Parasitol. 85, 556, 1999. 57. Wu, Z. et al. Polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) for the identi�cation of Trichinella isolates. Parasitology. 118, 211, 1999. 58. Wu, Z. et al. DNA �ngerprints of Trichinella as revealed by restriction fragment length polymorphism and single-strand conformational polymorphism (RFLP-SSCP). Mol. Cell. Probes 14, 291, 2000. 59. Zarlenga, D.S. et al. A single multiplex PCR for unequivocal differentiation of six distinct genotypes of Trichinella and three geographical genotypes of Trichinella pseudospiralis. Int. J. Parasitol. 29, 1859, 1999. 60. Pozio, E., and La Rosa, G. PCR-derived methods for the identi�cation of Trichinella parasites from animal and human samples. Methods Mol. Biol. 216, 299, 2003.
61. Rombout, Y.B., Bosch. S., and Van Der Giessen, J.W. Detection and identi�cation of eight Trichinella genotypes by reverse line blot hybridization. J. Clin. Microbiol. 39, 642, 2001.
62. MacPherson, J.M. et al. Variability of the random ampli�ed polymorphic DNA assay among thermal cyclers, and effects of primer and DNA concentration. Mol. Cell. Probes 7, 293, 1993.
63. Zarlenga, D.S., and Dame, J.B. The identi�cation and characterization of a break within the large subunit ribosomal RNA of Trichinella spiralis: Comparison of gap sequences within the genus, Mol. Biochem. Parasitol. 51, 281, 1992. 64. Pozio, E. et al. Twelve years of activity of the International Trichinella Reference Centre. Parasite, 8, S44, 2001. 65. Kapel, C.M., Webster, P., and Gamble, H.R. Muscle distribution of sylvatic and domestic Trichinella larvae in production animals and wildlife. Vet. Parasitol. 132, 101, 2005. 66. Nöckler, K., and Kapel, C.M.O. Detection and surveillance for Trichinella meat inspection and hygiene, and legislation. In: Dupouy-Camet, J., and Murrell, K.D. (eds), FAO/WHO/OIE Guidelines for the Surveillance, Management, Prevention and Control of Trichinellosis, Chapter 3. World Organisation for Animal Health, Paris, 2007. 68 Chapter 68 - Trichostrongylus
FIGURE 68.2 (a) Schematic representation of part of the rRNA transcriptional unit and relative locations of primers (NC5, NC2, jmAD, jmNA, and jhTsp) used in this study. The region harboring ITS-1, ITS-2, and 5.8S rDNA is ampli�ed with NC5 and NC2 primers. The species-speci�c primers, jmAD, jmNA, and jhTsp target the region within the ITS-1 sequence. (b) Speci�city of ITS-PCR in the recognition of the genomic DNA of other bacteria and protozoa. Lane M, 100 bp ladder, 600 bp and 100 bp bands are shown. Lane 1, N. americanus; lane 2, A. duodenale; lane 3, Trichostrongylus colubriformis; lane 4, Escherichia coli; lane 5, Entamoeba histolytica; lane 6, Giardia lam blia. Only speci�cally ampli�ed PCR products are detected on lanes 1–3. (From Yong, T.S. et al. Korean J. Parasitol. 45, 69, 2007. With permission.)
5. Rim, H.J. et al. Prevalence of intestinal parasite infections on a national scale among primary schoolchildren in Laos. Parasitol. Res. 91, 267, 2003.
6. Yong, T.S. et al. Differential diagnosis of Trichostrongylus and hookworm eggs via PCR using ITS-1 sequence. Korean J. Parasitol. 45, 69, 2007.
7. Faust, E.C., Russell, P.F., and Jung, R.C. Craig and Faust’s Clinical Parasitology. 8th ed. Lea & Febiger, Philadelphia, p. 289, 1970.
8. Gasser, R.B. et al. Rapid sequencing of rDNA from single worms and eggs of parasitic helminthes. Nucleic Acids Res. 21, 2525, 1993.
9. Schnieder, T., Heise, M., and Epe, C. Genus-speci�c PCR for the differentiation of eggs or larvae from gastrointestinal nematodes of ruminants. Parasitol. Res. 85, 895, 1999. 10. McMurtry, L.W. et al. Distinguishing morphological features of the third larval stage of ovine Trichostrongylus spp. Vet. Parasitol. 90, 73, 2000. 11. Gasser, R.B. et al. Species identi�cation of trichostrongyle nematodes by PCR-linked RFLP. Int. J. Parasitol. 24, 291, 1994. 12. Hoste, H. et al. Lack of intraspeci�c variation in the second internal transcribed spacer (ITS-2) of Trichostrongylus colubriformis ribosomal DNA. Int. J. Parasitol. 23, 1069, 1993. 13. Hoste, H. et al. A comparison of the �rst internal transcribed spacer of ribosomal DNA in seven species of Trichostrongylus (Nematoda: Trichostrongylidae). Int. J. Parasitol. 28, 1251, 1998. 14. Bott, N.J. et al. A combined microscopic-molecular method for the diagnosis of strongylid infections in sheep. Int. J. Parasitol. 39, 1277, 2009. 69 Chapter 69 - Trichuris
2. Gonçalves, M.L. et al. Human intestinal parasites in the past: New �ndings and a review. Mem. Ints. Oswaldo Cruz 98, 103, 2003.
3. Harter-Lailheugue, S., and Bouchet, F. Palaeoparasitological study of a typical elements of the low and high Nile Valley. Bull. Soc. Pathol. Exot. 99, 53, 2006.
4. Araujo, A. et al. Parasites as probes for prehistoric human migrations? Trends Parasitol. 24, 112, 2008.
5. Reinhard, K.J., Hevly, R.H., and Anderson, G.A. Helminth remains from prehistoric Indian coprolites on the Colorado Plateau. J. Parasitol. 73, 630, 1987.
6. Chabaud, A.G. CIH keys to the nematode parasites of vertebrates. In: Anderson, R.C., Chabaud, A.G., and Willmott, S. (eds), Class Nematoda. Commonwealth Agricultural Bureaux, Slough, Inglaterra, pp. 7–17, 1974.
7. de Ley, P., and Blaxter, M.L. Systematic position and phylogeny. In: Lee, D.L. (ed.), The Biology of Nematode. Taylor & Francis, London, pp. 1–30, 2002.
8. Lorenzen, S. Entwurf eines phylogenetischen systems der freilebenden nematoden. Veröffentlichungen des Institut für Meeresforshungen Bremerhaven 7, 472, 1981.
9. Inglis, W.G. An outline classi�cation of the phylum Nematoda. Aust. J. Zool. 31, 243, 1983.
10. de Ley, P.A. A Quick Tour of Nematode Diversity and Backbone of Nematode Phylogeny. WormBook, pp. 1–8, 2006. Website: http://www.wormbook.org. USA.
11. Gagarin, V.G. Critical evaluation of measured characters in the differentiation of helminth species (on the model of trichocephalids) (in Russian). In: Tokobaev, M.M. (ed), Fauna gelmintov zivotnych y rastenij kirgizii. ILIM, Frunze, pp. 3–9, 1974.
12. Dujardin, F. Histoire naturalle des helminthes ou vers intestinaux. Paris, 1841.
13. Chandler, A.C. Speci�c characters in the genus Trichuris with a description of the new species Trichuris tenuis from a camel. J. Parasitol., 16, 198, 1930.
14. Creplin, F.C.H. Cited by Schwartz (1926), 1825.
15. Leuckart, R. Cited by Schwartz (1926), 1876.
16. Schwartz, B. J. Agric. Res., Washington DC, 33, 311, 1926.
17. Shneider, A. Cited by Schwartz (1926), 1866.
18. Sondak, V.A. On the question of species independence of whipworms of man and swine. Parazitol. Sbornik. Zool. Inst. AN. SSSR, 10:195–204.
19. Pavlovski, E.N., and Sondak, V.A. On the speci�c differences between the whipworm of man and swine. Parazitol. Sbornik. Zool. Inst. AN. SSSR, 13:35–44.
20. Dinnik, N.N. Differences between Trichocephalus trichiura and Trichocephalus suis (Schrank, 1938). Med. Parazit. 1, 907, 1938.
21. Beer, R.J. The relationship between Trichuris trichiura (Linnaeus 1758) of man and Trichuris suis (Schrank 1889 of the pig). Res. Vet. Sci. 20, 47, 1976.
22. Cutillas, C. et al. Trichuris trichiura and Trichuris suis are different nematode species. Acta Trop. 111, 299, 2009.
23. Gállego Berenguer, J. Manual de parasitología. Morfología y biología de los parásitos de interés sanitario, Ed. Universitat de Barcelona, Spain, 2nd ed., pp. 342–346, 2003.
24. Bogitsh, B.J, Carter, C.E., and Oeltmann, T.N. Human Parasitology, 3rd ed., Elsevier Academic Press, USA/UK, pp. 324–327, 2005.
25. Skrjabin, K.I, Shikhobalova, N.P., and Orlow, I.V. Tricocephalidae and Capillariidae of Animals and the Man and the Diseases Caused by them. Izd. Akademii Nauk SSRR, Moskva, 587, 1957.
26. Schmidt, G.D., and Robert´s, L.S. Foundations of Parasitology, 7th ed., McGraw-Hill, New York, USA, pp. 397–400, 2005. 27. Tenora, F., Hovorka, I., and Hejlková, D. A supplement to the scanning electron microscopy of some Trichocephalus spp (Nematoda). Helminthologia 25, 227, 1988. 28. Tenora, F. et al. Some novel features of male posterior end of Trichuris discolor as revealed by scanning electron microscopy. Jpn. J. Parasitol. 41, 487, 1992. 29. Ooi, H.K. et al. Comparative study of Trichuris trichiura from non-human primates and from man and their difference with T. suis. J. Vet. Med. Sci. 55, 363, 1993. 30. Mahmoud, L.H. Scanning electron microscopy of Trichuris trichiura. J. Egypt. Soc. Parasitol. 32, 469, 2002. 31. Beaver, P.C., Jung, R.C., and Cupp, E.W. Parasitología Clínica, Ed. Salvat, Barcelona, 2nd ed., pp. 241–245, 1994. 32. Monroe, L.S. Gastrointestinal parasites. In: Haubrich, W.S., Schaffner, F., and Berk, J.E. (eds), Bockus Gastroenterology, 5th ed., Phil. Saunders, Philadelphia, USA, 3113, 1995. 33. Fishman, J.A., and Perrone, T.L. Colonic obstruction and perforation due to Trichuris trichiura. Am. J. Med. 77, 154, 1984. 34. Santos, F.L, Cerqueira, E.J., and Soares, N.M. Comparison of the thick smear and Kato–Katz techniques for diagnosis of intestinal helminth infections. Rev. Soc. Bras. Med. Trop., 38, 196, 2005. 35. Chandra, B., and Long, J.D. Diagnosis of Trichuris trichiura (whipworm) by colonoscopic extraction. J. Clin. Gastroenterol. 27, 152, 1998. 36. Bundy, D.A.P., and Cooper, E.S. Trichuris and trichuriasis in humans. Adv. Parasitol. 28, 107, 1989. 37. de Silva, N.R. et al. Soil-transmitted helminth infections: Updating the global picture. Trends Parasitol. 19, 547, 2003. 38. Cooper, E.S., and Bundy, D.A.P. Trichuris is not trivial. Parasitol. Today 4, 301, 1988. 39. Khuroo, M.S. et al. Trichuris dysentery syndrome: A common cause of chronic iron de�ciency anemia in adults in an endemic area (with videos). Gastrointest. Endoscopy 71, 200, 2010. 40. Bethony, J. et al. Soil-transmitted helminth infections: Ascariasis, trichuriasis, and hookworm. Seminar 367, 1521, 2006. 41. Galvani, A.P. Age-dependent epidemiological patterns and strain diversity in helminth parasites. J. Parasitol. 91, 24, 2005. 42. Cooper, E.S., and Bundy, D.A.P. Trichuriasis. In: Pawloswki, Z.-S. (ed.), Intestinal Helminthic Infections, Baillière’s Clin., Bailliere Tindall, London and Philadelphia, Trop. Med. Com. Dis. 2, 629, 1987. 43. Dunn, J.J. et al. Trichuris vulpis recovered from a patient with chronic diarrhea and �ve dogs. J. Clin. Microbiol. 40, 2703, 2002. 44. Mirdha, B.R. et al. Trichuris vulpis infection in slum children. Indian J. Gastroenterol. 17, 154, 1998. 45. Vásquez, O. et al. Infección Mixta por Trichuris trichiura y por Trichuris vulpis. Rev. Gastroenterol. Perú 17, 255–258, 1997. 46. Singh, S. et al. Trichuris vulpis infection in an Indian tribal population. J. Parasitol. 79, 457, 1993. 47. Yoshikawa, H. et al., Variations in egg size of Trichuris trichiura. Parasitol. Res. 75, 649, 1989. 48. Ferrer-Rodríguez, I., and Kozek, W.J. Abnormal Trichuris trichiura eggs detected during an epidemiological survey. P. R. Health Sci. J. 26, 237, 2007. 49. Kenney M., and Yermakov V. Infection of man with Trichuris vulpis, the whipworm of dogs. Am. J. Trop. Med. Hyg. 29, 1205, 1980. 50. Horii, Y., and Usui, M. Experimental transmission of Trichuris ova from monkeys to man. Trans. Roy. Soc. Trop. Med. Hyg. 79, 423, 1984.
51. Zaccone, P. et al. Schistosoma mansoni antigens modulate the activity of the innate immune response and prevent onset of type 1 diabetes. Eur. J. Inmunol. 33, 1439, 2003.
52. Cooper, E.S. et al. Intestinal nematode infections in children: The pathophysiological price paid. Parasitology 104, S91, 1992.
53. Kitis, G., Thompson, H., and Allan, R.N. Finger clubbing in in�ammatory bowel disease: Its prevalence and pathogenesis. Br. Med. J. 2(6194), 825–828, 1979.
54. Nokes, C. et al., Moderate to heavy infections of Trichuris trichiura affect cognitive function in Jamaican school children. Parasitology 104, 539, 1992.
55. Fort, M.M. et al. IL-25 induces IL-4, IL-5, and IL-13 and Th2associated pathologies in vivo. Immunity 15, 985, 2001.
56. Harnett, W., and Harnett, M.M. Molecular basis of worminduced immunomodulation. Parasite Immunol. 28, 535, 2006.
57. Finkelman, F.D. Interleukin-4 and interleukin-13 mediated host protection against intestinal nematode parasites. Inmunol. Rev. 201, 139, 2004.
58. Voehringer, D. et al. Type 2 immunity is controlled by IL-4/ IL-13 expression in hematopoietic non-eosinophil cells of the innate immune system. J. Exp. Med. 203, 1435, 2006.
59. Fox, J.G. et al. Concurrent enteric helminth infection modulates in�ammation and gastric immune responses and reduces helicobacter-induced gastric atrophy. Nat. Med. 6, 536, 2000.
60. Summers, R.W. et al. Trichuris suis therapy for active ulcerative colitis: A randomized controlled trial. Gastroenterology 128, 825, 2005.
61. Wang, L.J., Cao, Y., and Shi, H.N. Helminth infections and intestinal in�ammation. World J. Gastroenterol. 14, 5125, 2008.
62. WHO. Preventive Chemotheraphy in Human Helminthiasis: Coordinated use of Anthelminthic Drugs in Control Interventions: A Manual for Health Profesionals and Programme Managers. World Health Organization, Geneva, pp. 1–62, 2006.
63. Knopp, S. Diagnosis of soil-transmitted helminths in the era of preventive chemotherapy: Effect of multiple stool sampling and use of different diagnostic techniques. PloS Negl. Trop. Dis. 2, 331, 2008.
64. Ridley, D.S., and Hawgood, B.C. The value of formol-ether concentration of faecal cysts and ova. J. Clin. Pathol. 9, 74, 1956.
65. Allen, A.V.H., and Ridley, D.S. Further observations on the formol-ether concentration technique for faecal parasites. J. Clin. Pathol. 23, 545, 1970.
66. Katz, N., Chaves, A., and Pellegrino, J. A simple device for quantitative stool thick-smear technique in schistosomiasis mansoni. Rev. Inst. Med. Trop. São Paulo 14, 397, 1972.
67. Peters, P.A. et al. Quick Kato smear for �eld quanti�cation of Schistosoma mansoni eggs. Am. J. Trop. Med. Hyg. 29, 217, 1980.
68. WHO. Monitoring Anthelmitic Ef�cacy for Soil Transmitted Helminths (STH). World Health Organization, Geneva, 2008.
69. Cringoli, G. FLOTAC, a novel apparatus for a multivalent faecal egg count technique. Parassitologia 48, 381, 2006.
70. Goodman, D. et al. A comparison of methods for detecting the eggs of Ascaris, Trichuris, and hookworm in infant stool, and the epidemiology of infection in Zanzibari infants. Am. J. Trop. Med. Hyg. 74, 725, 2007.
71. Utzinger, J. et al. FLOTAC: A new sensitive technique for the diagnosis of hookworm infections in humans. Trans. Roy. Soc. Trop. Med. Hyg. 102, 84, 2008. 72. WHO. Bench Aids for the Diagnosis of Intestinal Parasites. World Heath Organization, Geneva, 1994. 73. Montresor, A. et al. Guidelines for the Evaluation of SoilTransmitted Helthminthiasis and Schistosomiasis at Community Level. A Guide for Managers of Control Programmes. World Heath Organization, Geneva, pp. 1–45, 1998. 74. Hall, A. Quantitative variability of nematode egg counts in faeces: A study among rural Kenyans. Trans. R. Soc. Trop. Med. Hyg. 75, 682, 1981. 75. Anderson, R.M., and Schad G.A. Hookworm burdens and faecal egg counts: An analysis of the biological basis variation. Trans. R. Soc. Trop. Med. Hyg. 79, 812, 1985. 76. Marti, H.P., and Koella, J.C. Multiple stool examinations for ova and parasites and rate of false-negative results. J. Clin. Microbiol. 31, 3044, 1993. 77. Booth, M. et al. The in�uence of sampling effort and performance of the Kato–Katz technique in diagnosing Schistosoma mansoni and hookworm co-infections in rural Cộte dÍvore. Parasitology 127, 525, 2003. 78. Knopp, S. et al. A single FLOTAC is more sensitive than triplicate Kato–Katz for the diagnosis of low-intensity soiltransmitted helminth infections. Trans. Roy. Soc. Trop. Med. Hyg. 103, 347, 2009. 79. Tarafder, M.R. et al. Estimating the sensitivity and speci�city of Kato–Katz stool examination technique for detection of hookworms, Ascaris lumbricoides and Trichuris trichiura infections in humans in the absence of a `gold standard`. Int. J. Parasitol. 40, 399, 2010. 80. Steinmann, P. et al. Extensive multiparasitism in a village of Yunnan province, Peoplés Republic of China, revealed by a suit of diagnostic methods. Am. J. Top. Med. Hyg. 78, 760, 2008. 81. Walden, J. Parasitic diseases. Other roundworms. Trichuris, hookworm and Strongyloides. Prim. Care 18, 53, 1991. 82. Mullen, K., and Prost, A. Decreased micro�larial load and its effect on the calculation of prevalence and the rate of false negatives in the detection of onchocerciasis. Int. J. Epidemiol. 12, 102, 1983. 83. Levecke, B. et al. Field validity and feasibility of four techniques for the detection of Trichuris in simians: A model for monitoring drug ef�cacy in public health? Negl. Trop. Dis. 3, 366, 2009. 84. Munene, E. et al. Helminth and protozoan gastrointestinal tract parasites in captive and wild-trapped African non-human primates. Vet. Parasitol. 78, 195, 1998. 85. Muriuki, S.M.K. et al. Some gastro-intestinal parasites of zoonotic (public health) importance commonly observed in old world non-human primates in Kenya. Acta Trop. 71, 73, 1998. 86. Ritchie, L.S. An ether sedimentation technique for routine stool examinations. Bull. U.S. Army Med. Dept. 8, 326, 1948. 87. Ahmadi, N.A., and Pakdad, K. Tween as a substitute for diethyl ether in the formalin-ether sedimentation technique. Iranian J. Publ. Health 36, 91, 2007. 88. Young, K.H. et al. Ethyl acetate as a substitute for diethyl ether in the formalin-ether sedimentation technique. J. Clin. Microbiol. 10, 852, 1979. 89. Kightlinger, L., and Kightlinger, M.B. Examination of faecal specimens by the formalin-detergent technique. Trans. R. Soc. Trop. Med. Hyg. 84, 417, 1990. 90. Parija, S.C. et al. Evaluation of formalin-acetone sedimentation in the concentration of stool for intestinal parasites. Trop. Doct. 33, 163, 2003. 91. Ahmadi, N.A. et al. Potency of wet mount, formalin-acetone and formalin-ether methods in detection of intestinal parasitic infections. Iranian J. Dis. Trop. Med. 12, 43, 2007.
92. Cheesbrough, M. District Laboratory Practice in Tropical Countries, Part 1. Cambridge University Press, Cambridge, 1998.
93. Boswell, M.V., and Collins, V.J. Diethyl ether and chloroform. In: Collins, V.J. (ed), Physiologic and Pharmacologic bases of Anesthesia. Williams & Wilkins, Pennsylvania, pp. 650–662, 1996.
94. Ahmadi, N.A., and Damraj, F-A. A �eld evaluation of formalin-gasoline technique in the concentration of stool for detection of intestinal parasites. Parasitol. Res. 104, 553, 2009.
95. Bruschi, F., and Castagna, B. The serodiagnosis of parasitic infections. Parassitologia 46, 141, 2004.
96. Joseph, L. et al. Bayesian estimation of disease prevalence and the parameters of diagnostic tests in the absence of a gold standard. Am. J. Epidemiol., 141, 263, 1995.
97. Dendukuri, N. et al. Bayesian sample size determination for prevalence and diagnostic test studies in the absence of a gold standard test. Biometrics 60, 388, 2004.
98. Kyung-Sun, O. et al. Trichuris trichiura infection diagnosed by colonoscopy: Case reports and review of literature. Korean J. Parasitol., 47, 273, 2009.
99. Tokmak, N. et al. Computed tomographic �ndings of trichuriasis. World J. Gastroenterol. 12, 4270, 2006.
100. Cancrini, G., and Iori, A. Traditional and innovative diagnostic tools: When and why they should be applied. Parassitologia 46, 173, 2004. 101. Silva, A.J. et al. Fast reliable extraction of protozoan parasite DNA from fecal specimens. Mol. Diag. 4, 57, 1999.
102. Shnieder, T., Heise, M., and Epe, C. Genus-speci�c PCR for differentiation of eggs or larvae from gastrointestinal nematodes of ruminants. Parasitol. Res. 85, 895, 1999.
103. Machado, E. et al. Random ampli�ed polymorphic DNA analysis of DNA extracted from Trichuris trichiura (Linnaeus, 1771) eggs and its prospective application to paleoparasitological studies. Mem. Inst. Oswaldo Cruz 98, 59, 2003.
104. Nagano, I. et al. Identi�cation of Trichinella isolates by polymerase chain reaction-restriction fragment length polymorphism of the mitochondrial cytochome c oxidase subunit I gene. Int. J. Parasitol. 29, 1113, 1999.
105. Zarlenga, D.S. et al. Characterization and detection of a newly described Asian Taeniid using closed ribosomal DNA fragments and sequence ampli�cation by polimerase chain reaction. Exp. Parasitol. 72, 174, 1991.
106. Zarlenga, D.S. Cloning and characterization of ribosomal RNA genes from three species of Haemonchus (Nematoda: Trichostrongyloidea) and identi�cation of PCR primers for rapid differentiation. Exp. Parasitol. 8, 28, 1994. 107. Oliveros, R. et al. Characterization of four species of Trichuris (Nematoda: Enoplida) by their second internal transcribed spacer ribosomal DNA sequence. Parasitol. Res. 86, 1008, 2000. 108. Foreyt, W.J. Diagnostic parasitology. Vet. Clin. Am. Small Anim. Pract. 19, 979, 1989. 109. Mochizuki, R. et al. PCR-based species-speci�c ampli�cation of ITS of Mecistocirrus digitatus and its application in identi�cation of G.I. nematode eggs in bovine faeces. J. Vet. Med. Sci. 68, 345, 2006. 110. Gasser, R.B. et al. Species markers for equine strongyles detected in intergenic spacer rDNA by PCR-RFLP. Mol. Cell. Prob. 10, 371, 1996. 111. Stoll, N.R. This wormy world. J. Parasitol. 33, 1–18, 1947. 112. Maizels, R.M. et al. Vaccination against helminth parasites: The ultimate challenge for vaccinologists? Inmunol. 171, 125, 1999. 113. Hotez, P.J. et al. Helminth infections: Soil-transmitted helminth infections and schistosomiasis. In: Jamison, D.T., Breman, J.G., Measham, A.R., Alleyne, G., Claeson, M., Evans, D.B., Jha, P., Mills, A., and Musgrove, P. (eds). Disease Control Priorities in Developing Countries, 2nd edition. Oxford University Press, New York, Chapter 24, pp. 467–482, 2006. 114. Molyneux, D.H., Hotez, P.J., and Fenwick, A. Rapid-impact interventions: How a policy of integrated control for Africás neglected tropical diseases could bene�t the poor. PloS Med. 2, 336, 2005. 115. Loukas, A. et al. Hookworm vaccines: Past, present, and future. Lancet Infect. Dis. 6, 733, 2006. 116. Utzinger, J., and Keiser, J., Schistosomiasis and soiltransmitted helminthiasis: Common drugs for treatment and control. Expert Opin. Pharmacother. 5, 263, 2004. 117. Lammie, P.J., Fenwick, A., and Utzinger, J. A blueprint for success: Integration of neglected tropical disease control programmes. Trends Parasitol. 22, 313, 2006. 118. Hotez, P.J. et al. Control of neglected tropical diseases. N. Engl. J. Med. 357, 1018, 2007. 119. Kabaterine, N.B. The control of schistosomiasis and soil-transmitted helminths in East Africa. Trends Parasitol. 22, 332, 2006. 120. Keiser, J., and Utzinger, J. Ef�cacy of current drugs against soil-transmitted helminth infections. J. Am. Med. Assoc. 299, 23, 2008. 121. Olsen, A. et al. Albendazole and mebendazole have low ef�cacy against Trichuris trichiura in school-age children in Kabale District, Uganda. Trans. Roy. Soc. Trop. Med. 103, 443, 2009. 122. ten Hove, R.J. et al. Molecular diagnostics of intestinal parasites in returning travellers. Eur. J. Clin. Microbiol. Infect. Dis. 28, 1045, 2009. 70 Chapter 70 - Wuchereria
1. Pfarr, K.M. et al., Filariasis and lymphoedema. Parasite Immunol., 31, 664, 2009.
2. Southgate, B., The signi�cance of low density micro�laremia in the transmission of lymphatic �larial parasites. J. Trop. Med. Hyg., 95, 79, 1992.
3. Leite, A.B. et al., Assessment of family and neighbors of an individual infected with Wuchereria bancrofti from a nonendemic area in the city of Maceió, Brazil. Braz. J. Infect. Dis., 14, 125, 2010.
4. Rosenblatt, J.E., Laboratory diagnosis of infections due to blood and tissue parasites. Clin. Infect. Dis., 49, 1103, 2009.
5. Amaral, F. et al., Live adult worms detected by ultrasonography in human Bancroftian �lariasis. Am. J. Trop. Med. Hyg., 50, 753, 1994.
6. Weil, G.J., and Ramzy, R.M.R., Diagnostic tools for �lariasis elimination programmes. Trends Parasitol., 23, 78, 2007.
7. Chularerk, P., and Desowitz, R.S., A simpli�ed membrane �ltration technique for the diagnosis of micro�laremia. J. Parasitol., 56, 623, 1970.
8. Hoti, S.L. et al., A method for detecting micro�laraemia, �larial speci�c antigens and antibodies and typing of parasites for drug resistance and genotypes using �nger prick blood sample. Acta Trop., 107, 268, 2008.
9. Nuchprayoon, S. et al., Endemic bancroftian �lariasis in Thailand: Detection by Og4C3 antigen capture ELISA and the polymerase chain reaction. J. Med. Assoc. Thai., 84, 1300, 2001.
10. Lammie, P.J. et al., Recombinant antigen-based antibody assays for the diagnosis and surveillance of lymphatic �lariasis—a multicenter trial. Filaria J., 3, 9, 2004.
11. Ramzy, R.M., Field application of PCR-based assays for monitoring Wuchereria bancrofti infection in Africa. Ann. Trop. Med. Parasitol., 96, S55, 2002. 12. Ramzy, R.M.R. et al., A polymerase chain reaction-based assay for detection of Wuchereria bancrofti in human blood and Culex pipiens. Trans. R. Soc. Trop. Med. Hyg., 91, 156, 1997. 13. Williams, S.A. et al., Development and standardization of a rapid, PCR-based method for the detection of Wuchereria bancrofti in mosquitoes, for xenomonitoring the human prevalence of bancroftian �lariasis. Ann. Trop. Med. Parasitol., 96, S41, 2002. 14. Fischer, P., Boakye, D., and Hamburger, J., Polymerase chain reaction-based detection of lymphatic �lariasis. Med. Microbiol. Immunol., 192, 3, 2003. 15. Vasuki, V. et al., A simple and rapid DNA extraction method for the detection of Wuchereria bancrofti infection in the vector mosquito, Culex quinquefasciatus by Ssp I PCR assay. Acta Trop., 86, 109, 2003. 16. Vasuki, V., Hoti, S.L., and Patra, K.P., RT-PCR assay for the detection of infective (L3) larvae of lymphatic �larial parasite, Wuchereria bancrofti, in vector mosquito Culex quinquefasciatus. J. Vector Borne Dis., 45, 207, 2008. 17. Helmy, H. et al., Test strip detection of Wuchereria bancrofti ampli�ed DNA in wild-caught Culex pipiens and estimation of infection rate by a PoolScreen algorithm. Trop. Med. Int. Health, 9, 158, 2004. 18. Supali, T. et al., Estimation of the prevalence of lymphatic �lariasis by a pool screen PCR assay using blood spots collected on �lter paper. Trans. R. Soc. Trop. Med. Hyg., 100, 753, 2006. 19. Boakye, D.A. et al., Monitoring lymphatic �lariasis interventions: Adult mosquito sampling, and improved PCR-based pool screening method for Wuchereria bancrofti infection in Anopheles mosquitoes. Filaria J., 6, 13, 2007. 20. Kagai, J.M. et al., Molecular technique utilising sputum for detecting Wuchereria bancrofti infections in Malindi, Kenya. East Afr. Med. J., 85, 118, 2008. 21. Chambers, E.W. et al., Xenomonitoring of Wuchereria bancrofti and Diro�laria immitis infections in mosquitoes from American Samoa: Trapping considerations and a comparison of polymerase chain reaction assays with dissection. Am. J. Trop. Med. Hyg., 80, 774, 2009. 22. Intapan, P.M. et al., Rapid detection of Wuchereria bancrofti and Brugia malayi in mosquito vectors (Diptera: Culicidae) using a real-time �uorescence resonance energy transfer multiplex PCR and melting curve analysis. J. Med. Entomol., 46, 158, 2009. 23. Zhong, M. et al., A PCR assay for detection of the parasite Wuchereria bancrofti in human blood samples. Am. J. Trop. Med. Hyg., 54, 357, 1996. 24. Dassanayake, R.S., Chandrasekharan, N.V., and Karunanayake, E.H., Trans-spliced leader RNA, 5S-rRNA genes and novel variant orphan spliced-leader of the lymphatic �larial nematode Wuchereria bancrofti, and a sensitive polymerase chain reaction based detection assay. Gene, 269, 185, 2001. 25. Chansiri, K. et al., Detection of Plasmodium falciparum and Wuchereria bancrofti infected blood samples using multiplex PCR. Mol. Cell. Probes, 15, 201, 2001. 26. Chansiri, K., and Phantana, S., A polymerase chain reaction assay for the survey of bancroftian �lariasis. Southeast Asian J. Trop. Med. Public Health, 33, 504, 2002. 27. Lulitanond, V. et al., Rapid detection of Wuchereria bancrofti in mosquitoes by LightCycler polymerase chain reaction and melting curve analysis. Parasitol. Res., 94, 337, 2004. 28. Kanjanavas, P. et al., Detection of lymphatic Wuchereria bancrofti in carriers and long-term storage blood samples using semi-nested PCR. Mol. Cell. Probes, 19, 169, 2005.
29. Mishra, K. et al., Combined detection of Brugia malayi and Wuchereria bancrofti using single PCR. Acta Trop., 93, 233, 2005.
30. Rao, R.U. et al., A real-time PCR-based assay for detection of Wuchereria bancrofti DNA in blood and mosquitoes. Am. J. Trop. Med. Hyg., 74, 826, 2006.
31. Rao, R.U. et al., A qPCR-based multiplex assay for the detection of Wuchereria bancrofti, Plasmodium falciparum and Plasmodium vivax DNA. Trans. R. Soc. Trop. Med. Hyg., 103, 365, 2009.
32. Nuchprayoon, S., Junpee, A., and Poovorawan, Y., Random ampli�ed polymorphic DNA (RAPD) for differentiation between Thai and Myanmar strains of Wuchereria bancrofti. Filaria J., 6, 6, 2007.
33. Kamal, I.H. et al., Evaluation of a PCR-ELISA to detect Wuchereria bancrofti in Culex pipiens from an Egyptian village with a low prevalence of �lariasis. Ann. Trop. Med. Parasitol., 95, 833, 2001.
34. Jiménez, M. et al., Detection and discrimination of Loa loa, Mansonella perstans and Wuchereria bancrofti by PCRRFLP and nested-PCR of ribosomal DNA ITS1 region. Exp. Parasitol., 127, 282, 2011.
35. Laney, S.J. et al., Detection of Wuchereria bancrofti L3 larvae in mosquitoes: A reverse transcriptase PCR assay evaluating infection and infectivity. PLoS Negl. Trop. Dis., 4, e602, 2010.
36. Mehlotra, R.K. et al., Molecular-based assay for simultaneous detection of four Plasmodium spp. and Wuchereria bancrofti infections. Am. J. Trop. Med. Hyg., 82, 1030, 2010. 37. Torres, E.P. et al., Detection of bancroftian �lariasis in human blood samples from Sorsogon province, the Philippines by polymerase chain reaction. Parasitol. Res., 87, 677, 2001. 38. Bhandari, Y. et al., Analysis of polymorphism of 18S rRNA gene in Wuchereria bancrofti micro�lariae. Microbiol. Immunol., 49, 909, 2005. 39. Nuchprayoon, S. et al., Detection and differentiation of �larial parasites by universal primers and polymerase chain reactionrestriction fragment length polymorphism analysis. Am. J. Trop. Med. Hyg., 73, 895, 2005. 40. Farid, H.A. et al., A critical appraisal of molecular xenomonitoring as a tool for assessing progress toward elimination of lymphatic �lariasis. Am. J. Trop. Med. Hyg., 77, 593, 2007. 41. Hoti, S.L. et al., Laboratory evaluation of Ssp I PCR assay for the detection of Wuchereria bancrofti infection in Culex quinquefasciatus. Indian J. Med. Res., 114, 59, 2001. 42. Sakthidevi, M. et al., Lymphatic �larial species differentiation using evolutionarily modi�ed tandem repeats: Generation of new genetic markers. Infect. Genet. Evol., 10, 591, 2010. 43. Van Hoegaerden, M., and Ivanoff, B., A rapid, simple method for isolation of viable micro�lariae. Am. J. Trop. Med. Hyg., 35, 148, 1986. 44. Nuchprayoon, S., DNA-based diagnosis of lymphatic �lariasis. Southeast Asian J. Trop. Med. Public Health, 40, 904, 2009. 45. Hoti, S.L. et al., An allele speci�c PCR assay for screening for drug resistance among Wuchereria bancrofti populations in India. Indian J. Med. Res., 130, 193, 2009. 46. Palumbo, E., Filariasis: Diagnosis, treatment and prevention. Acta Biomed., 79, 106, 2008.
Section VI
Arthropoda: Arachnida 71 Chapter 71 - Argasidae (Soft Ticks)
13. Thewes, M. et al., Argas re�exus (the pigeon tick)–a household pest. Acta Derm. Venereol., 77, 173-4, 1997.
14. Haag-Wackernagel, D., and Bircher, A.J., Ectoparasites from feral pigeons affecting humans. Dermatology, 220, 82, 2010.
15. Dautel, H., Scheurer, S., and Kahl, O., The pigeon tick (Argas re°exus): Its biology, ecology, and epidemiological aspects. Zentralbl. Bakteriol., 289, 745, 1999.
16. Reeves, W.K., Molecular evidence for a novel Coxiella from Argas monolakensis (Acari: Argasidae) from Mono Lake, California, USA. Exp. Appl. Acarol., 44, 57, 2008.
17. Condy, J.B. et al., The effects of the bites of Argas brumpti (Acarina: Argasidae) on humans. Cent. Afr. J. Med., 26, 212, 1980.
18. Fukunaga, M. et al. Molecular phylogenetic analysis of Ixodid ticks based on the ribosomal DNA spacer, internal transcribed spacer 2, sequences. J. Parasitol., 86, 38, 2000.
19. Cutler, S.J., Browning, P., and Scott, J.C., Ornithodoros moubata, a soft tick vector for Rickettsia in east Africa? Ann. N. Y. Acad. Sci., 1078, 373, 2006.
20. Beck, A.F., Holscher, K.H., and Butler, J.F., Life cycle of Ornithodoros turicata americanus (Acari: Argasidae) in the laboratory. J. Med. Entomol., 23, 313, 1986.
21. Helmy, N., Seasonal abundance of Ornithodoros (O.) savignyi and prevalence of infection with Borrelia spirochetes in Egypt. J. Egypt. Soc. Parasitol., 30, 607, 2000.
22. Charrel, R.N. et al., Alkhurma hemorrhagic fever virus in Ornithodoros savignyi ticks. Emerg. Infect. Dis., 13, 153, 2007.
23. Krinsky, W.L., Dermatoses associated with the bites of mites and ticks (Arthropoda: Acari). Int. J. Dermatol., 22, 75, 1983.
24. Loftis, A.D. et al., Detection of Rickettsia, Borrelia, and Bartonella in Carios kelleyi (Acari: Argasidae). J. Med. Entomol., 42, 473, 2005.
25. Mumcuoglu, K.Y. et al., Argasid ticks as possible vectors of West Nile virus in Israel. Vector Borne Zoonotic Dis., 5, 65, 2005.
26. Tosti, A., Peluso, A.M., and Spedicato, S., Urticaria–angioedema syndrome caused by an Argas re°exus sting. Contact Dermatitis, 19, 315, 1988.
27. Veraldi, S., Scarabelli, G., and Grimalt, R., Acute urticaria caused by pigeon ticks (Argas re°exus). Int. J. Dermatol., 35, 34, 1996.
28. Veraldi, S. et al., Skin manifestations caused by pigeon ticks (Argas re°exus). Cutis, 61, 38, 1998.
29. Sirianni, M.C. et al., Anaphylaxis after Argas re°exus bite. Allergy, 55, 303, 2000.
30. Rolla, G. et al., Allergy to pigeon tick (Argas re°exus): Demonstration of speci�c IgE-binding components. Int. Arch. Allergy Immunol., 135, 293, 2004.
31. Hilger, C. et al., IgE-mediated anaphylaxis caused by bites of the pigeon tick Argas re�exus: Cloning and expression of the major allergen Arg r 1. J. Allergy Clin. Immunol., 115, 617, 2005.
32. Quercia, O. et al., Anaphylactic shock to Argas re°exus bite. Eur. Ann. Allergy Clin. Immunol., 37, 66, 2005.
33. Kleine-Tebbe, J. et al., Bites of the European pigeon tick (Argas re°exus): Risk of IgE-mediated sensitizations and anaphylactic reactions. J. Allergy Clin. Immunol., 117, 190, 2006.
34. Spiewak, R. et al., Allergy to pigeon tick (Argas re°exus) in Upper Silesia, Poland. Ann. Agric. Environ. Med., 13, 107, 2006.
35. Weckesser, S. et al., Anaphylactic reactions to bites of the pigeon tick Argas re°exus. Eur. J. Dermatol., 20, 244, 2010. 36. Dworkin, M.S., Schwan, T.G., and Anderson, D.E. Jr., Tickborne relapsing fever in North America. Med. Clin. North. Am., 86, 413, 2002. 37. Dworkin, M.S., Tick-borne relapsing fever. Infect. Dis. Clin. North. Am., 22, 449, 2008. 38. Aher, A.R. et al., A case report of relapsing fever. Indian J. Pathol. Microbiol., 51, 292, 2008. 39. Moemenbellah-Fard, M.D. et al., Tick-borne relapsing fever in a new highland endemic focus of western Iran. Ann. Trop. Med. Parasitol., 103, 529, 2009. 40. Hoskins, J.D., Ixodid and argasid ticks. Keys to their identi�cation. Vet. Clin. North. Am. Small. Anim. Pract., 21, 185, 1991. 41. Black, W.C., and Piesman, J., Phylogeny of hard- and softtick taxa (Acari: Ixodida) based on mitochondrial 16S rDNA sequences. Proc. Natl. Acad. Sci. USA, 91, 10034, 1994. 42. Black, W.C., Klompen, J.S.H., and Keirans, J.E., Phylogenetic relationships among tick subfamilies (Ixodida: Ixodidae: Argasidae) based on the 18S nuclear rDNA gene. Mol. Phylog. Evol., 7, 129, 1997. 43. Crampton, A., McKay, Y., and Barker, S.C., Phylogeny of ticks (Ixodida) inferred from nuclear ribosomal DNA. Int. J. Parasitol., 26, 511, 1996. 44. Mangold, A.J., Bargues, M.D., and Mas-Coma, S., Mitochondrial 16S rRNA sequences and phylogenetic relationships of Rhipicephalus and other tick genera among Metastriata (Acari: Ixodidae). Parasitol. Res., 84, 478, 1998. 45. Hwang, U.W., and Kim, W., General properties and phylogenetic utilities of nuclear ribosomal DNA and mitochondrial DNA commonly used in molecular systematics, Korean J. Parasitol., 37, 215, 1999. 46. Cruickshank, R.H., Molecular marker for the phylogenetics of mites and ticks. Syst. Appl. Acarol., 3, 1, 2002. 47. Anderson, J.M., Ammerman, N.C., and Norris, D.E., Molecular differentiation of metastriate tick immatures. Vector Borne Zoonotic Dis., 4, 334, 2004. 48. Vial, L. et al., Molecular divergences of the Ornithodoros sonrai soft tick species, a vector of human relapsing fever in West Africa. Microbes Infect., 8, 2605, 2006. 49. Geraci, N.S. et al., Variation in genome size of argasid and ixodid ticks. Insect. Biochem. Mol. Biol., 37, 399, 2007. 50. Marrelli, M.T. et al., Taxonomic and phylogenetic relationships between neotropical species of ticks from genus Amblyomma (Acari: Ixodidae) inferred from second internal transcribed spacer sequences of rDNA. J. Med. Entomol., 44, 222, 2007. 51. Chitimia, L. et al., Molecular characterization of hard and soft ticks from Romania by sequences of the internal transcribed spacers of ribosomal DNA. Parasitol. Res., 105, 907, 2009. 52. Chitimia, L. et al., Genetic characterization of ticks from southwestern Romania by sequences of mitochondrial cox1 and nad5 genes. Exp. Appl. Acarol., 52, 305, 2010. 53. Halos, L. et al., Determination of an ef�cient and reliable method for DNA extraction from ticks. Vet. Res. 5, 1, 2004. 54. Shone, S.M. et al., A novel real-time PCR assay for the speciation of medically important ticks. Vector Borne Zoonotic Dis., 6, 152, 2006. 55. Chao, L.L., Wu, W.J., and Shih, C.M., Species identi�cation of Ixodes granulatus (Acari: Ixodidae) based on internal transcribed spacer 2 (ITS2) sequences. Exp. Appl. Acarol., 54, 51, 2011. 56. Dergousoff, S.J., and Chilton, N.B., Differentiation of three species of ixodid tick, Dermacentor andersoni, D. variabilis and D. albipictus, by PCR-based approaches using markers in ribosomal DNA. Mol. Cell. Probes, 21, 343, 2007. 72 Chapter 72 - Demodex (Hair Follicle Mite)
10. Ozdemir, M.H. et al., Hilal, Prevalence of Demodex in health personnel working in the autopsy room. Am. J. Forensic Med. Pathol., 26, 18, 2005.
11. Yao, Y.E., Guo, N., and Wu, L.P., The effect of temperature on the viability of Demodex folliculorum and Demodex brevis. Parasitol. Res., 105, 1623, 2009.
12. Zhao, Y.E., Guo, N., and Wu, L.P., In�uence of temperature and medium on viability of Demodex folliculorum and Demodex brevis (Acari: Demodicidae). Exp. Appl. Acarol., 54, 421, 2011.
13. Kemal, M. et al., The prevalence of Demodex folliculorum in blepharitis patients and the normal population. Ophthalmic Epidemiol., 12, 287, 2005.
14. Aylesworth, R., and Vance, J.C., Demodex folliculorum and Demodex brevis in cutaneous biopsies. J. Am. Acad. Dermatol., 7, 583, 1982.
15. Bonnar, E., Eustace, P., and Powell, F.C., Demodex mite in normal skin. Lancet, 337, 1168, 1991.
16. Pena, G.P., and Andrade Filho, J.S., Is Demodex really nonpathogenic? Rev. Inst. Med. Trop Sao Paulo, 42, 171, 2000.
17. Ayres, S. Jr., Demodex folliculorum as a pathogen. Cutis, 37, 441, 1986.
18. Heacock, C.E., Clinical manifestations of demodicosis, J. Am. Optom. Assoc., 57, 914, 1986.
19. Sahn, E.E., and Sheridan, D.M., Demodicidosis in a child with leukaemia. J. Am. Acad. Dermatol., 27, 799, 1992.
20. Redondo Mateo, J. et al., Demodex-attributed rosacea-like lesions in AIDS. Acta Derm. Venereol., 73, 437, 1993.
21. Barrio, J. et al., Rosacea-like demodicidosis in an HIVpositive child. Dermatology, 192, 143, 1996.
22. Castanet, J. et al., Demodicidosis in an immunode�cient child. Pediatr. Dermatol., 14, 219, 1997.
23. Patrizi, A. et al., Demodicidosis in immunocompetent young children: Report of eight cases. Dermatology, 195, 239, 1997.
24. Pallotta, S. et al., Unilateral demodicidosis. Eur. J. Dermatol., 8, 191, 1998.
25. Aydıngoz, I.E., Dervent, B., and Guney, O., Demodex folliculorum in pregnancy. Int. J. Dermatol., 39, 743, 2000.
26. Jansen, T. et al., Rosacea-like demodicidosis associated with acquired immunode�ciency syndrome. Br. J. Dermatol., 144, 139, 2001.
27. Kastner, U., Kreuter, A., and Altmeyer, P., Rosacea-like demodicidosis associated with acquired immunode�ciency syndrome. Br. J. Dermatol., 144, 139, 2001.
28. Akdeniz, S. et al., Is Demodex folliculorum larger in diabetic patients? J. Eur. Acad. Dermatol. Venereol., 16, 539, 2002.
29. Baima, B., and Sticherling, M., Demodicidosis revisited. Acta Derm. Venereol., 82, 3, 2002.
30. Seyhan, M.E. et al., Density of Demodex folliculorum in haematological malignancies. J. Int. Med. Res., 32, 411, 2004.
31. Akilov, O.E., Butov, Y.S., and Mumcuoglu, K.Y., A clinicopathological approach to the classi�cation of human demodicosis. J. Dtsch. Dermatol. Ges., 3, 607, 2005.
32. Ciftci, I.H. et al., Demodex folliculorum in patients with rheumatoid arthritis. Acta Parasitol., 2007, 52, 70–73.
33. Ozcelik, S. et al., The incidence of Demodex folliculorum in patients with chronic kidney de�ciency. Turkiye Parazitol. Derg., 31, 66, 2007.
34. Kulac, M. et al., Clinical importance of Demodex folliculorum in patients receiving phototherapy. Int. J. Dermatol., 47, 72, 2008.
35. Hsu, C.K., Hsu, M.M., and Lee, J.Y., Demodicosis: A clinicopathological study. J. Am. Acad. Dermatol., 60, 453, 2009. 36. Kligman, A.M., and Christensen, M.S., Demodex folliculorum: Requirements for understanding its role in human skin disease. J. Invest. Dermatol., 131, 8, 2011. 37. Purcell, S.M., Hayes, T.J., and Dixon, S.L., Pustular folliculitis associated with Demodex folliculorum. J. Am. Acad. Dermatol., 15, 1159, 1986. 38. Shelley, W.B., Shelley, E.D., and Burmeister, V., Unilateral demodectic rosacea. J. Am. Acad. Dermatol., 20, 915, 1989. 39. Sibenge, S., and Gawkrodger, D.J., Rosacea: A study of clinical patterns, blood �ow, and the role of Demodex folliculorum. J. Am. Acad. Dermatol., 26, 590, 1992. 40. Bonnar, E., Eustace, P., and Powell, F.C., The Demodex mite population in rosacea. J. Am. Acad. Dermatol., 28, 443, 1993. 41. Vollmer, R.T., Demodex-associated folliculitis. Am. J. Dermatopathol., 18, 589, 1996. 42. Hwang, S.M. et al., Demodecidosis manifested on the external genitalia. Int. J. Dermatol., 37, 634, 1998. 43. Georgala, S. et al., Increased density of Demodex folliculorum and evidence of delayed hypersensitivity reaction in subjects with papulopustular rosacea. J. Eur. Acad. Dermatol. Venereol., 15, 441, 2001. 44. Cribier, B., Pathophysiology of rosacea: Redness, telangiectasia, and rosacea. Ann. Dermatol. Venereol., 138, S184, 2011. 45. Erbagci, Z., and Ozgoztasi, O., The signi�cance of Demodex folliculorum density in rosacea, Int. J. Dermatol., 37, 421, 1998. 46. Forton, F. et al., Demodicosis and rosacea: Epidemiology and signi�cance in daily dermatologic practice. J. Am. Acad. Dermatol., 52, 74, 2005. 47. Morras, P.G. et al., Rosacea-like demodicidosis in an immunocompromised child. Pediatr. Dermatol., 20, 8, 2003. 48. Schaller, M., Sander, C.A., and Plewig, G., Demodex abscesses: Clinical and therapeutic challenges. J. Am. Acad. Dermatol., 49, S272, 2003. 49. Karincaoglu, Y. et al., The clinical importance of Demodex folliculorum presenting with nonspeci�c facial signs and symptoms. J. Dermatol., 31, 618, 2004. 50. Dolenc-Voljc, M., Pohar, M., and Lunder, T., Density of Demodex folliculorum in perioral dermatitis, Acta Derm. Venereol., 85, 211, 2005. 51. el-Bassiouni, S.O. et al., A study on Demodex folliculorum mite density and immune response in patients with facial dermatoses. J. Egypt. Soc. Parasitol., 35, 899, 2005. 52. Okyay, P. et al., Prevalence of Demodex folliculorum in young adults: Relation with sociodemographic/hygienic factors and acne vulgaris. J. Eur. Acad. Dermatol. Venereol., 20, 474, 2006. 53. García-Vargas, A., Mayorga-Rodríguez, J.A., and SandovalTress, C., Scalp demodicidosis mimicking favus in a 6-yearold boy. J. Am. Acad. Dermatol., 57, S19, 2007. 54. Lee, J.Y., and Hsu, C.K., Granulomatous rosacea-like demodicidosis. Dermatol. Online J., 13, 9, 2007. 55. Moravvej, H. et al., Association of rosacea with demodicosis. Arch. Iran. Med., 10, 199, 2007. 56. Fernandez-Flores, A., and Alija, A., Scalp folliculitis with Demodex: Innocent observer or pathogen? Braz. J. Infect. Dis., 13, 81, 2009. 57. Zhao, Y.E. et al., Retrospective analysis of the association between Demodex infestation and rosacea. Arch. Dermatol., 146, 896, 2010; Erratum: 146, 1412, 2010. 58. Humiczewska, M., Demodex folliculorum and Demodex brevis (Acarida) as the factors of chronic marginal blepharitis. Wiad. Parazytol., 37, 127, 1991.
59. Kamoun, B. et al., Blepharitis due to Demodex: myth or reality? J. Fr. Ophtalmol., 22, 525, 1999.
60. Koksal, M. et al., A rare agent of chalazion: Demodectic mites. Can. J. Ophthalmol., 38, 605, 2003.
61. Arıcı, M.K. et al., The Prevalence of Demodex folliculorum in blepharitis patients and the normal population. Ophthalmic Epidemiol., 12, 287, 2005.
62. Rodriguez, A.E., Ferrer, C., and Alio, J.L., Chronic blepharitis and Demodex. Arch. Soc. Esp. Ophthalmol., 80, 635, 2005.
63. Czepita, D. et al., Demodex folliculorum and Demodex brevis as a cause of chronic marginal blepharitis. Ann. Acad. Med. Stetin., 53, 63, 2007.
64. Lee, S.H. et al., The relationship between demodex and ocular discomfort. Invest. Ophthalmol. Vis. Sci., 51, 2906, 2010.
65. Lacey, N., Kavanagh, K., and Tseng, S.C., Under the lash: Demodex mites in human diseases. Biochemistry (Lond), 31, 2, 2009.
66. Liang, L. et al., Ocular demodicosis as a potential cause of pediatric blepharoconjunctivitis. Cornea, 29, 1386, 2010.
67. English, F.P. et al., The presence of the parasite Demodex folliculorum on the skin surface of the eyelid. Aust. N. Z. J. Ophthalmol., 19, 229, 1991.
68. Mor�n Maciel, B.M., Demodicidosis in a female patient treated as allergic blepharoconjunctivitis. A case report. Rev. Alerg. Mex., 50, 232, 2003.
69. Kheirkhah, A. et al., Corneal manifestations of ocular Demodex infestation Am. J. Ophthalmol., 143, 743, 2007.
70. Hoekzema, R., Hulsebosch, H.J., and Bos, J.D., Demodicidosis or rosacea: What did we treat? Br. J. Dermatol., 133, 294, 1995.
71. Crawford, G.H., Pelle, M.T., and James, W.D., Rosacea: I Etiology, pathogenesis, and subtype classi�cation. J. Am. Acad. Dermatol., 51, 327, 2004.
72. Buechner, S.A., Rosacea: An update, Dermatology, 210, 100, 2005.
73. Benessahraoui, M. et al., Demodicidosis in a child with xantholeukaemia associated with type 1 neuro�bromatosis. Eur. J. Dermatol., 13, 311, 2003.
74. Türk, M. et al., Comparison of incidence of Demodex folliculorum on the eyelash follicule in normal people and blepharitis patients. Turkiye Parazitol. Derg., 31, 296, 2007.
75. Liu, J., Sheha, H., and Tseng, S.C., Pathogenic role of Demodex mites in blepharitis. Curr. Opin. Allergy Clin. Immunol., 10, 505, 2010.
76. Clifford, C.W., and Fulk, G.W., Association of diabetes, lash loss, and Staphylococcus aureus with infestation of eyelids by Demodex folliculorum (Acari: Demodicidae). J. Med. Entomol., 27, 467, 1990.
77. Li, J. et al., Correlation between ocular Demodex infestation and serum immunoreactivity to bacillus proteins in patients with facial rosacea. Ophthalmology, 117, 870, 2010.
78. Lacey, N. et al., Mite-related bacterial antigens stimulate in�ammatory cells in rosacea. Br. J. Dermatol., 157, 474, 2007.
79. Akilov, O.E., and Mumcuoglu K.Y., Association between human demodicosis and HLA class I. Clin. Exp. Dermatol., 28, 70, 2003.
80. Mumcuoglu, K.Y., and Akilov, O.E., The role of HLA A2 and Cw2 in the pathogenesis of human demodicosis. Dermatology, 210, 109, 2005. 81. Akilov, O.E., and Mumcuoglu, K.Y., Immune response in demodicosis. J. Eur. Acad. Dermatol. Venereol., 18, 440, 2004. 82. Gao, Y.-Y. et al., Clinical treatment of ocular demodicosis by lid scrub with tea tree oil, Cornea, 26, 136, 2007. 83. Demmler, M. et al., Blepharitis. Demodex folliculorum, associated pathogen spectrum and speci�c therapy. Ophthalmologe, 94, 191, 1997. 84. Damian, D., and Rogers, M., Demodex infestation in a child with leukaemia: Treatment with ivermectin and permethrin. Int. J. Dermatol., 42, 724, 2003. 85. Antille, C., Saurat, J.H., and Lubbe, J., Induction of rosaceiform dermatitis during treatment of facial in�ammatory dermatoses with tacrolimus ointment. Arch. Dermatol., 140, 457, 2004. 86. El-Shazly, A.M. et al., Treatment of human Demodex folliculorum by camphor oil and metronidazole. J. Egypt. Soc. Parasitol., 34, 107, 2004. 87. Clyti, E., Sayavong, K., and Chanthavisouk, K., Demodecidosis in a patient infected by HIV: Successful treatment with ivermectin. Ann. Dermatol. Venereol., 132, 459, 2005. 88. Yagdiran Düzgün, O., and Aytekin, S., Comparison of Demodex folliculorum density in haemodialysis patients with a control group. J. Eur. Acad. Dermatol. Venereol., 21, 480, 2007. 89. Bikowski, J.B., and Del Rosso, J.Q., Demodex dermatitis: A retrospective analysis of clinical diagnosis and successful treatment with topical crotamiton. J. Clin. Aesthet. Dermatol., 2, 20, 2009. 90. Inceboz, T. et al., Diagnosis and treatment of demodectic blepharitis. Turkiye Parazitol. Derg., 33, 32, 2009. 91. Nara, T. et al., Eosinophilic folliculitis with a Demodex folliculorum infestation successfully treated with ivermectin in a man infected with human immunode�ciency virus. Clin. Exp. Dermatol., 34, e981, 2009. 92. Kheirkhah, A. et al., Fluorescein dye improves microscopic evaluation and counting of Demodex in blepharitis with cylindrical dandruff. Cornea, 26, 697, 2007. 93. Forton, F., and Seys, B., Density of Demodex folliculorum in rosacea: A case-control study using standardized skin-surface biopsy. Br. J. Dermatol., 128, 650, 1993. 94. As¸kin, U., and Seçkin, D., Comparison of the two techniques for measurement of the density of Demodex folliculorum: Standardized skin surface biopsy and direct microscopic examination. Br. J. Dermatol., 162, 1124, 2010. 95. Noge, K. et al., Identi�cation of astigmatid mites using the second internal transcribed spacer (ITS2) region and its application for phylogenetic study. Exp. Appl. Acarol., 35, 29, 2005. 96. Wong, S.F. et al., Molecular identi�cation of house dust mites and storage mites. Exp. Appl. Acarol., 55, 123, 2011. 97. Borgo, S.N. et al., PCR analysis for Wolbachia in human and canine Demodex mites. Arch. Dermatol. Res., 301, 747, 2009. 98. Toops, E. et al., Extraction and characterization of DNA from Demodex canis. Int. J. Appl. Res. Vet. Med., 8, 31, 2010. 99. Ravera, I. et al., Development of a real-time PCR to detect Demodex canis DNA in different tissue samples. Parasitol. Res., 108, 305, 2011. 100. Desloire, S. et al., Comparison of four methods of extracting DNA from D. gallinae (Acari: Dermanyssidae). Vet. Res., 37, 725, 2006.
101. Basta-Juzbasic, A., Subic, J.S., and Ljubojevic, S., Demodex folliculorum in development of dermatitis rosaceiformis steroidica and rosacea-related diseases. Clin. Dermatol., 20, 135, 2002.
102. Erbagci, Z., Erbagci, I., and Erkilic, S., High incidence of demodicidosis in eyelid basal cell carcinomas. Int. J. Dermatol., 42, 567, 2003. 103. Gao, Y.-Y. et al., High prevalence of ocular Demodex in eyelashes with cylindrical dandruffs, Invest. Ophthalmol. Vis. Sci., 46, 3089, 2005. 104. Forton, F., and Song, M., Limitation of standardized skin surface biopsy in measurement of the density of Demodex folliculorum. A case report. Br. J. Dermatol., 139, 697, 1998. 73 Chapter 73 - Dermanyssus, Ornithonyssus, and Trombicula (Red, Fowl, and Harvest Mites)
6. Baker, A.S., Mites and ticks of domestic animals: An identi�cation guide and information source. London: The Stationary Of�ce, pp. 134–136, 1999.
7. Hutcheson, H.J., and Oliver, J.H. Jr., Spermiogenesis and reproductive biology of Dermanyssus gallinae (DeGeer) (Parasitiformes: Dermanyssidae). J. Med. Entomol., 25, 321, 1988.
8. Lucky, A.W. et al., Avian mite bites acquired from a new source-pet gerbils. Arch. Dermatol., 137, 167, 2001.
9. Oliver, J.H., Cytogenetics of mites and ticks. Annu. Rev. Entomol., 22, 407, 1977.
10. Sparagano, O. et al., Prevalence and key �gures for the poultry red mite Dermanyssus gallinae infections in poultry farm systems. Exp. Appl. Acarol., 48, 3, 2009.
11. Valiente-Moro, C., Chauve, C., and Zenner, L., Vectorial role of some Dermanyssoid mites, Parasite, 12, 99, 2005.
12. Valiente Moro, C. et al., The poultry red mite (Dermanyssus gallinae): A potential vector of pathogenic agents. Exp. Appl. Acarol., 48, 93, 2009.
13. Daniel, M., and Stekol’nikov, A.A., Chigger mites (Acari: Trombiculidae) from Makalu region in Nepal Himalaya, with a description of three new species. J. Med. Entomol., 46, 753, 2009.
14. Brown, W.A., Description of six new species of the genus Leptotrombidium from Mindanao Island and notes on the medically important genera of chiggers (Acari: Trombiculidae) of the Philippine Islands. J. Med. Entomol., 29, 284, 1992.
15. Shatrov, A.B., Stylostome formation in trombiculid mites (Acariformes: Trombiculidae). Exp. Appl. Acarol., 49, 261, 2009.
16. Iwasa, M. et al., Trombiculid mites (Acari: Trombiculidae) and Rickettsia tsutsugamushi isolated from wild rodents in a new endemic area of Japan. J. Med. Entomol., 27, 501, 1990.
17. Takahashi, M. et al., A new member of the trombiculid mite family Neotrombicula nagayoi (Acari: Trombiculidae) induces human dermatitis. Southeast Asian J. Trop. Med. Public Health, 35, 113, 2004.
18. Daniela, M. et al., Chigger mites (Acari, Trombiculidae) parasitizing small mammals in the Eastern Hindu Kush and some other Afghan areas. Parasitol. Res., 107, 1221, 2010.
19. Vargo, J., Ginsberg, M., and Mizrahi, R., Human infestation by the pigeon mite: A case report. Am. J. Infect. Control, 11, 24, 1983.
20. Regan, A.M., Metersky, M.L., and Craven, D.E., Nosocomial dermatitis and pruritus caused by pigeon mite infestation. Arch. Intern. Med., 174, 2185, 1987.
21. Yassien, N.A. et al., Immunohistopathological status of the skin in cases infested with three species of mites. J. Egypt. Soc. Parasitol., 26, 567, 1996.
22. Rossiter, A., Occipational otitis externa in chicken catchers. J. Laryngol. Otol., 111, 366, 1997.
23. Chauve, C., The poultry red mite Dermanyssus gallinae: Current situation and future prospects. Vet. Parasitol., 8, 364, 1998.
24. Rosen, S., Yeruham, I., and Braverman, Y., Dermatitis in humans associated with the mites Pyemotes tritici, Dermanyssus gallinae, Ornithonyssus bacoti and Androlaelaps casalis in Israel. Med. Vet. Entomol., 16, 442, 2002.
25. Nisbet, A.J. et al., A house dust mite allergen homologue from poultry red mite Dermanyssus gallinae (De Geer). Parasite Immunol., 28, 401, 2006.
26. Green, B., Sparling, J., and Sparling, M.B., What’s eating you? Pigeon mite (Dermanyssus gallinae). Cutis, 80, 461, 2007. 27. Akdemir, C., Gülcan, E., and Tanritanir, P., Case report: Dermanyssus gallinae in a patient with pruritus and skin lesions. Turkiye Parazitol. Derg., 33, 242, 2009. 28. McClain, D., Dana, A.N., and Goldenberg, G., Mite infestations. Dermatol. Therapy, 22, 327, 2009. 29. Dogramaci, A.C., Culha, G., and Ozçelik, S., Dermanyssus gallinae infestation: An unusual cause of scalp pruritus treated with permethrin shampoo. J. Dermatol. Treat., 21, 319, 2010. 30. Haag-Wackernagel, D., and Bircher, A.J., Ectoparasites from feral pigeons affecting humans. Dermatology, 220, 82, 2010. 31. Ca�ero, M.A. et al., Pseudoscabies caused by Dermanyssus gallinae in Italian city dwellers: A new setting for an old dermatitis. J. Eur. Acad. Dermatol. Venereol., 22, 1382, 2008. 32. Chirico, J. et al., The poultry red mite, Dermanyssus gallinae, a potential vector of Erysipelothrix rhusiopathiae causing erysipelas in hens. Med. Vet. Entomol., 17, 232, 2003. 33. De Luna, C.J. et al., The poultry red mite Dermanyssus gallinae as a potential carrier of vector-borne diseases. Ann. N. Y. Acad. Sci., 1149, 255, 2008. 34. Bellanger, A.P. et al., Nosocomial dermatitis caused by Dermanyssus gallinae. Infect. Control Hosp. Epidemiol., 29, 282, 2008. 35. Gupta, A.K., Billings, J.K., and Ellis, C.N., Chronic pruritus: An uncommon cause. Avian mite dermatitis caused by Ornithonyssus sylviarum (Northern fowl mite). Arch. Dermatol., 124, 1102, 1988. 36. Watson, C.R., Human infestation with bird mites in Wollongong. Commun. Dis. Intel., 27, 259, 2003. 37. Orton, D.I., Warren, L.J., and Wilkinson, J.D., Avian mite dermatitis. Clin. Exp. Dermatol., 25, 129, 2000. 38. Baumstark, J., Beck, W., and Hofmann, H., Outbreak of tropical rat mite (Ornithonyssus bacoti) dermatitis in a home for disabled persons. Dermatology, 215, 66, 2007. 39. Beck, W., Occurrence of a house-infesting tropical rat mite (Ornithonyssus bacoti) on murides and human beings. Travel Med. Infect. Dis., 6, 245, 2008. 40. Axman, W.R., and Brummer, J.J., Chigger mite infestation. J. Am. Podiatr. Med. Assoc., 93, 399, 2003. 41. Thind, B.B., and Ford, H.L., Assessment of susceptibility of the poultry red mite Dermanyssus gallinae (Acari: Dermanyssidae) to some acaricides using an adapted �lter paper based bioassay. Vet. Parasitol., 144, 344, 2007. 42. Marangi, M. et al., Evaluation of the poultry red mite, Dermanyssus gallinae (Acari: Dermanyssidae) susceptibility to some acaricides in �eld populations from Italy. Exp. Appl. Acarol., 48, 11, 2009. 43. Roy, L. et al., Diversity of phylogenetic information according to the locus and the taxonomic level: An example from a parasitic mesostigmatid mite genus. Int. J. Mol. Sci., 11, 1704, 2010. 44. Söller, R. et al., Phylogenetic relationships within terrestrial mites (Acari: Prostigmata, Parasitengona) inferred from comparative DNA sequence analysis of the mitochondrial cytochrome oxidase subunit I gene. Mol. Phylogenet. Evol., 18, 47, 2001. 45. Cruickshank, R.H., Molecular markers for the phylogenetics of mites and ticks. Syst. Appl. Acarol., 7, 3, 2002. 46. Jeyaprakash, A., and Hoy M.A., Mitochondrial 12S rRNA sequences used to design a molecular ladder assay to identify six commercially available phytoseiids (Acari: Phytoseiidae). Biol. Control, 25, 136, 2002. 47. Wuyts, J. et al., The European ribosomal RNA database. Nucleic Acids Res., 32, D101, 2004.
48. Klompen, H., Lekveishvili, M., and Black, W.C., Phylogeny of parasitiform mites (Acari) based on rRNA. Mol. Phylogenet. Evol., 43, 936, 2007.
49. Potenza, L. et al., Characterization of Dermanyssus gallinae (Acarina: Dermanissydae) by sequence analysis of the ribosomal internal transcribed spacer regions. Vet. Res. Commun., 33, 611, 2009.
50. Brännström, S. et al., Genetic differences in internal transcribed spacer 1 between Dermanyssus gallinae from wild birds and domestic chickens. Med. Vet. Entomol., 22, 152, 2008. 51. Marangi, M. et al., Phylogenetic relationship between Dermanyssus gallinae populations in European countries based on mitochondrial COI gene sequences. Exp. Appl. Acarol., 48, 143, 2009. 52. Stekolnikov, A.A., and Klimov, P.B., A revision of chiggers of the minuta species-group (Acari: Trombiculidae: Neotrombicula Hirst, 1925) using multivariate morphometrics. Syst. Parasitol., 77, 55, 2010. 53. Desloire, S. et al., Comparison of four methods of extracting DNA from D. gallinae (Acari: Dermanyssidae). Vet. Res., 37, 725, 2006. 74 Chapter 74 - Ixodidae (Hard Ticks)
20. Labruna, M.B. et al., Amblyomma latepunctatum, a valid tick species (Acari: Ixodidae) long misidenti�ed with both Amblyomma incisum and Amblyomma scalpturatum. J. Parasitol., 91, 527, 2005.
21. Labruna, M.B. et al., Human parasitism by the capibara tick, Amblyomma dubitatum (Acari: Ixodida). Entomol. News, 118, 77, 2007.
22. Labruna, M.B. et al., Redescription of the female, description of the male, and several new records of Amblyomma parkeri (Acari: Ixodidae), a South America tick species. Exp. Appl. Acarol., 49, 243, 2009.
23. Aguirre, D.H. et al., Amblyomma tigrinum (Acari: Ixodidae): New data on hosts and biology of immature stages and on DNA composition. Vet. Parasitol., 134, 267, 2005.
24. Nava, S., Mangold, A.J., and Guglielmone, A.A., The natural hosts of larvae and nymphs of Amblyomma tigrinum Koch, 1844 (Acari: Ixodidae). Vet. Parasitol., 140, 124, 2006.
25. Nava, S. et al., Distribution, hosts, 16S rDNA sequences and phylogenetic position of the Neotropical tick Amblyomma parvum (Acari: Ixodidae). Ann. Trop. Med. Parasitol., 102, 409, 2008.
26. Nava, S. et al., Ecology of Amblyomma neumanni (Acari: Ixodidae). Acta Trop., 111, 226, 2009.
27. Goddard, J., A review of the disease agents harbored and transmitted by the lone star tick, Amblyomma americanum (L.). Southwest. Entomol., 12, 158, 1987.
28. Goddard, J., and Varela-Stokes, A.S., Role of the lone star tick, Amblyomma americanum (L.), in human and animal diseases. Vet. Parasitol., 160, 1, 2009.
29. Childs, J.E., and Paddock, C.D., The ascendancy of Amblyomma americanum as a vector of pathogens affecting humans in the United States. Ann. Rev. Entomol., 48, 307, 2003.
30. Paddock, C.D., and Yabsley, M.J., Ecological havoc, the rise of white-tailed deer, and the emergence of Amblyomma americanum-associated zoonoses in the United States. Curr. Top. Microbiol. Immunol., 315, 289, 2007.
31. Trout, R.T., Steelman, C.D., and Szalanski, A.L., Population genetics of Amblyomma americanum (Acari; Ixodidae) collected from Arkansas. J. Med. Entomol., 47, 152, 2010.
32. Lockhart, J.M. et al., Temporal association of Amblyomma americanum with the presence of Ehrlichia chaffeensis reactive antibodies in white-tailed deer. J. Wildl. Dis. 31, 119, 1995.
33. Loftis, A.D. et al., Geographic distribution and genetic diversity of the Ehrlichia sp. from Panola Mountain in Amblyomma americanum. BMC Infect. Dis. 8, 54, 2008.
34. Anderson, B.E. et al. Amblyomma americanum: A potential vector of human ehrlichiosis. Am. J. Trop. Med. Hyg. 49, 239, 1993.
35. Jasinskas, A., Zhong, J., and Barbour, A.G., Highly prevalent Coxiella sp. bacterium in the tick vector Amblyomma americanum. Appl. Environ. Microbiol., 73, 334, 2007.
36. Mixson, T.R. et al., Prevalence of Ehrlichia, Borrelia, and Rickettsial agents in Amblyomma americanum (Acari: Ixodidae) collected from nine states. J. Med. Entomol., 43, 1261, 2006.
37. Szabo, M.P.J. et al., Biology and life cycle of Amblyomma incisum (Acari: Ixodidae). Exp. Appl. Acarol., 48, 263, 2009.
38. Kang, Y.B. et al. Amblyomma testudinarium Koch, 1844: Discovery and record in Korea, and indenti�cation and redescription of male tick. Korean J. Vet. Res., 21, 65, 1981.
39. Kim, J. et al., A case of Amblyomma testudinarium tick bite in a Korean woman. Korean J. Parasitol., 48, 313, 2010. 40. Sanches, G.S., Bechara, G.H., and Camargo-Mathias, M.I., Morphological description of Amblyomma brasiliense Aragão, 1908 (Acari: Ixodidae) larvae and nymphs. Rev. Bras. Parasitol. Vet., 18, 15, 2009. 41. Yamada, Y. et al., A case of tick bite from Amblyomma testudinarium on the glans penis. J. Dermatol., 23, 136, 1996. 42. Zahler, M., Gothe, R., and Rinder, H., Genetic evidence against a morphologically suggestive conspeci�city of Dermacentor reticulatus and D. marginatus (Acari: Ixodidae). Int. J. Parasitol., 25, 1413, 1995. 43. Zahler, M., and Gothe, R., Evidence for the reproductive isolation of Dermacentor marginatus and Dermacentor reticulatus (Acari: Ixodidae) ticks based on cross-breeding, morphology and molecular studies. Exp. Appl. Acarol., 21, 685, 1997. 44. Bursali, A. et al., Ixodid ticks (Acari: Ixodidae) infesting humans in Tokat Province of Turkey: species diversity and seasonal activity. J. Vector Ecol., 35, 180, 2010. 45. Bursali, A. et al., Species diversity of ixodid ticks feeding on humans in Amasya, Turkey: Seasonal abundance and presence of Crimean-Congo hemorrhagic fever virus. J. Med. Entomol., 48, 85, 2011. 46. Omeragic, J., Ixodid ticks in Bosnia and Herzegovina. Exp. Appl. Acarol., 53, 301, 2011. 47. Patterson, E.I., Dergousoff, S.J., and Chilton, N.B., Genetic variation in the 16S mitochondrial DNA gene of two Canadian populations of Dermacentor andersoni (Acari: Ixodidae). J. Med. Entomol., 46, 475, 2009. 48. Ahmed, B.M., Taha, K.M., and Hussein, A.M., Life cycle of Hyalomma anatolicum Koch, 1844 (Acari: Ixodidae) fed on rabbits, sheep and goats. Vet. Parasitol., 177, 353, 2011. 49. Zheng, H. et al., Development and biological characteristics of Haemaphysalis longicornis (Acari: Ixodidae) under �eld conditions. Exp. Appl. Acarol., 53, 377, 2011. 50. Poucher, K.L. et al. Molecular genetic key for the identi�cation of 17 Ixodes species of the United States (Acari: Ixodidae): A methods model. J. Parasitol., 85, 623, 1999. 51. Labruna, M.B. et al. Study of the taxonomic status of Ixodes didelphidis (Acari: Ixodidae). J. Med. Entomol., 39, 242, 2002. 52. Xu, G. et al., Molecular phylogenetic analysis indicate that the Ixodes ricinus complex is a paraphyletic group. J. Parasitol., 89, 452, 2003. 53. Eremeeva, M.E., and Stromdahl, E.Y., Short report: New spotted fever group Rickettsia in a Rhipicephalus turanicus tick removed from a child in eastern Sicily, Italy. Am. J. Trop. Med. Hyg., 84, 99, 2011. 54. Wikswo, M.E. et al., Detection of Rickettsia rickettsii and Bartonella henselae in Rhipicephalus sanguineus ticks from California. J. Med. Entomol., 44, 158, 2007. 55. Dantas-Torres, F., The brown dog tick, Rhipicephalus sanguineus (Latreille, 1806) (Acari: Ixodidae): From taxonomy to control. Vet. Parasitol., 152, 173, 2008. 56. Demma, L.J. et al., An outbreak of Rocky Mountain spotted fever associated with a novel tick vector, Rhipicephalus sanguineus, in Arizona, 2004: Preliminary report. Ann. N. Y. Acad. Sci., 1078, 342, 2006. 57. Parola, P., Socolovschi, C., and Raoult, D., Deciphering the relationships between Rickettsia conorii conorii and Rhipicephalus sanguineus in the ecology and epidemiology of Mediterranean spotted fever. Ann. N. Y. Acad. Sci., 1166, 49, 2009. 58. de Oliveira, P.R. et al., Comparison of the external morphology of Rhipicephalus sanguineus (Latreille, 1806) (Acari: Ixodidae) ticks from Brazil and Argentina. Vet. Parasitol., 129, 139, 2005.
59. Yamasaki, T., Oda, T., and Kitaoka, S., A case of human infestation with a female of Amblyomma testudinarium Koch, 1844 (Acarina; Ixodidae). Trop. Med., 21, 167, 1979.
60. Krinsky, W.L., Dermatoses associated with the bites of mites and ticks (Arthropoda: Acari). Int. J. Dermatol., 22, 75, 1983.
61. Nakamura-Uchiyama, F. et al., Amblyomma testudinarium tick bite: One case of engorged adult and a case of extraordinary number of larval tick infestation. J. Dermatol., 27, 774, 2000.
62. Kahl, O. et al., Tick infection rates with Borrelia: Ixodes ricinus versus Haemaphysalis concinna and Dermacentor reticulatus in two locations in eastern Germany. Med. Vet. Entomol., 6, 363, 1992.
63. Chao, L.L., Wu, W.J., and Shih, C.M., Molecular analysis of Ixodes granulatus, a possible vector tick for Borrelia burgdorferi sensu lato in Taiwan. Exp. Appl. Acarol., 48, 329, 2009.
64. Dumler, J.S., and Bakken, J.S., Human ehrlichioses: Newly recognized infections transmitted by ticks. Annu. Rev. Med. 49, 201, 1998.
65. Felek, S. et al., Sensitive detection of Ehrlichia chaffeensis in cell culture, blood, and tick specimens by reverse transcription- PCR. J. Clin. Microbiol., 39, 460, 2001.
66. Steiert, J.G., and Gilfoy, F., Infection rates of Amblyomma americanum and Dermacentor variabilis by Ehrlichia chaffeensis and Ehrlichia ewingii in southwest Missouri. Vector Borne Zoonotic Dis., 2, 53, 2002.
67. Anderson, J.F. et al., Transstadial transfer of West Nile virus by three species of ixodid ticks (Acari: Ixodidae). J. Med. Entomol., 40, 528, 2003.
68. Apperson, C.S. et al., Tick-borne diseases in North Carolina: Is “Rickettsia amblyommii” a possible cause of rickettsiosis reported as Rocky Mountain spotted fever? Vector Borne Zoonotic Dis., 8, 597, 2008.
69. Venzal, J.M. et al., Guglielmone AA Amblyomma triste Koch, 1844 (Acari: Ixodidae): Hosts and seasonality of the vector of Rickettsia parkeri in Uruguay. Vet. Parasitol., 155, 104, 2008.
70. Oliveira, K.A. et al., Amblyomma imitator ticks as vectors of Rickettsia rickettsii, Mexico. Emerg. Infect. Dis., 16, 1282, 2010.
71. Hilburn, L.R., and Sattler, P.W., Electrophoretically detectable protein variation in natural population of the lone star tick, Amblyomma americanum (Acari: Ixodidae). Heredity, 56, 67, 1986.
72. Wesson, D.M. et al., Investigation of the validity of species status of Ixodes dammini (Acari: Ixodidae) using rDNA. Proc. Natl. Acad. Sci. USA, 90, 10221, 1993.
73. Barker, S.C., Distinguishing species and populations of Rhipicephalini ticks with ITS 2 ribosomal RNA. J. Parasitol., 84, 887, 1998.
74. Fournier, P.E. et al., Genetic identi�cation of rickettsiae isolated from ticks in Japan. J. Clin. Microbiol., 40, 2176, 2002.
75. Anderson, J.M., Ammerman, N.C., and Norris, D.E., Molecular differentiation of metastriate tick immatures. Vector Borne Zoonotic Dis., 4, 334, 2004.
76. Geraci, N.S. et al., Variation in genome size of argasid and ixodid ticks. Insect Biochem. Mol. Biol., 37, 399, 2007.
77. Pagel Van Zee, J. et al., Tick genomics: The Ixodes genome project and beyond. Int. J. Parasitol., 37, 1297, 2007.
78. Nava, S., Guglielmone, A.A., and Mangold, A.J., An overview of systematics and evolution of ticks. Front. Biosci., 14, 2857, 2009. 79. Dharmarajan, G., and Rhodes, O.E., Evaluating levels of PCR ef�ciency and genotyping error in DNA extracted from engorged and non-engorged female Dermacentor variabilis ticks. Med. Vet. Entomol., 25, 109, 2011. 80. Black, W.C., and Piesman, J., Phylogeny of hard- and softtick taxa (Acari: Ixodida) based on mitochondrial 16S rDNA sequences. Proc. Natl. Acad. Sci. USA, 91, 10034, 1994. 81. Black, W.C., Klompen, J.S.H., and Keirans, J.E., Phylogenetic relationships among tick subfamilies (Ixodida: Ixodidae: Argasidae) based on the 18S nuclear rDNA gene. Mol. Phylogenet. Evol., 7, 129, 1997. 82. Caporale, D.A. et al., Discriminating between Ixodes ticks by means of mitochondrial DNA sequences. Mol. Phylogenet. Evol., 4, 361, 1995. 83. McLain, D.K. et al., Variation in ribosomal DNA internal transcribed spaces 1 among eastern populations of Ixodes scapularis (Acari: Ixodidae). J. Med. Entomol., 32, 353, 1995. 84. Rich, S.M. et al., Distribution of the Ixodes ricinus-like ticks of eastern North America. Proc. Natl. Acad. Sci. USA, 92, 6284, 1995. 85. Crampton, A., McKay, Y., and Barker, S.C., Phylogeny of ticks (Ixodida) inferred from nuclear ribosomal DNA. Int. J. Parasitol., 26, 511, 1996. 86. Norris, D.E. et al., Population genetics of Ixodes scapularis (Acari: Ixodidae) based on mitochondrial 16S and 12S genes. J. Med. Entomol., 33, 78, 1996. 87. Norris, D.E. et al., Taxonomic status of Ixodes neotomae and I. spinipalpis (Acari: Ixodidae) based on mitochondrial DNA evidence. J. Med. Entomol., 34, 696, 1997. 88. Mangold, A.J., Bargues, M.D., and Mas-Coma, S., Mitochondrial 16S rRNA sequences and phylogenetic relationships of Rhipicephalus and other tick genera among Metastriata (Acari: Ixodidae). Parasitol. Res., 84, 478, 1998. 89. Fukunaga, M. et al., Molecular phylogenetic analysis of Ixodid ticks based on the ribosomal DNA spacer, internal transcribed spacer 2, sequences. J. Parasitol., 86, 38, 2000. 90. Cruickshank, R.H., Molecular marker for the phylogenetics of mites and ticks. Syst. Appl. Acarol., 3, 1, 2002. 91. Hlinka, O., Murrell, A., and Barker, S.C., Evolution of the secondary structure of the rRNA internal transcribed spacer 2 (ITS2) in hard tick (Ixodidae, Arthropoda). Heredity, 88, 275, 2002. 92. Estrada-Peña, A. et al., The Amblyomma maculatum Koch, 1844 (Acari: Ixodidae: Amblyomminae) tick group: Diagnostic characters, description of the larva of A. parvitarsum Neumann, 1901, 16S rDNA sequences, distribution and hosts. Syst. Parasitol., 60, 99, 2005. 93. Reichard, M.V. et al., Sequence variation of the ribosomal DNA second internal transcribed spacer region in two spatially distinct population of Amblyomma americanum (L.) (Acari: Ixodidae). J. Parasitol., 91, 260, 2005. 94. Marrelli, M.T. et al., Taxonomic and phylogenetic relationships between neotropical species of ticks from genus Amblyomma (Acari: Ixodidae) inferred from second internal transcribed spacer sequences of rDNA. J. Med. Entomol., 44, 222, 2007. 95. Chitimia, L. et al., Molecular characterization of hard and soft ticks from Romania by sequences of the internal transcribed spacers of ribosomal DNA. Parasitol. Res., 105, 907, 2009. 96. Chitimia, L. et al., Genetic characterization of ticks from southwestern Romania by sequences of mitochondrial cox1 and nad5 genes. Exp. Appl. Acarol., 52, 305, 2010.
97. Chao, L.L., Wu, W.J., and Shih, C.M., Species identi�cation of Ixodes granulatus (Acari: Ixodidae) based on internal transcribed spacer 2 (ITS2) sequences. Exp. Appl. Acarol., 54, 51, 2010.
98. Rumer, L. et al., Differentiation of medically important EuroAsian tick species Ixodes ricinus, Ixodes persulcatus, Ixodes hexagonus, and Dermacentor reticulatus by polymerase chain reaction. Vector Borne Zoonotic Dis., 11, 899, 2010.
99. Tian, Z. et al., Discrimination between Haemaphysalis longicornis and H. qinghaiensis based on the partial 16S rDNA and the second internal transcribed spacer (ITS-2). Exp. Appl. Acarol., 54, 165, 2011. 100. Halos, L. et al., Determination of an ef�cient and reliable method for DNA extraction from ticks. Vet. Res., 5, 1, 2004. 101. Shone, S.M. et al., A novel real-time PCR assay for the speciation of medically important ticks. Vector Borne Zoonotic Dis., 6, 152, 2006. 102. Dergousoff, S.J., and Chilton, N.B., Differentiation of three species of ixodid tick, Dermacentor andersoni, D. variabilis and D. albipictus, by PCR-based approaches using markers in ribosomal DNA. Mol. Cell. Probes, 21, 343, 2007. 75 Chapter 75 - Pyroglyphidae and Glycyphagidae (House Dust and Storage Mites)
18. Fernández-Caldas, E., and Lockey, R.F., Blomia tropicalis, a mite whose time has come. Allergy, 59, 1161, 2004.
19. Thind, B.B., and Clarke, P.G., The occurrence of mites in cereal-based foods destined for human consumption and possible consequences of infestation. Exp. Appl. Acarol., 25, 203, 2001.
20. Müsken, H. et al., Sensitization to different mite species in German farmers: Clinical aspects. J. Investig. Allergol. Clin. Immunol., 10, 346, 2000.
21. Arlian, L.G. et al., Cross-antigenicity between the scabies mite, Sarcoptes scabiei, and the house dust mite, Dermatophagoides pteronyssinus. J. Invest. Dermatol., 96, 349, 1991.
22. Arlian, L.G. et al., Allergenicity of the mite, Blomia tropicalis. J. Allergy Clin. Immunol., 91, 1042, 1993.
23. Arlian, L.G., Morgan, M.S., and Neal, J.S., Dust mite allergens: Ecology and distribution. Curr. Allergy Asthma Rep., 2, 401, 2002.
24. Arlian, L.G. et al., Cross-reactivity between storage and dust mites and between mites and shrimp. Exp. Appl. Acarol., 47, 159, 2009.
25. Erben, A.M. et al., Anaphylaxis after ingestion of beignets contaminated with Dermatophagoides farinae. J. Allergy Clin. Immunol., 92, 846, 1993.
26. Matsumoto, T. et al., Systemic anaphylaxis after eating storage- mite-contaminated food. Int. Arch. Allergy Immunol., 109, 197, 1996.
27. Stanaland, B.E. et al., Positive nasal challenge response to Blomia tropicalis. J. Allergy Clin. Immunol., 97, 1045, 1996.
28. Li, C.P. et al., Acaroid mite, intestinal and urinary acariasis. World J. Gastroenterol., 9, 874, 2003.
29. Milián, E., and Díaz, A.M., Allergy to house dust mites and asthma. P.R. Health Sci. J., 23, 47, 2004.
30. Wen, D.C. et al., Systemic anaphylaxis after the ingestion of pancake contaminated with the storage mite Blomia freemani. Ann. Allergy Asthma Immunol., 95, 612, 2005.
31. Kuljanac, I., The role of Dermatophagoides pteronyssinus in atopic dermatitis. Acta Dermatovenerol. Croat., 14, 86, 2006.
32. Nuttall, T.J. et al., House dust and forage mite allergens and their role in human and canine atopic dermatitis. Vet. Dermatol., 17, 223, 2006.
33. Solarz, K., Indoor mites and forensic acarology. Exp. Appl. Acarol., 49, 135, 2009.
34. Sade, K., Roitman, D., and Kivity, S., Sensitization to Dermatophagoides, Blomia tropicalis, and other mites in atopic patients. J. Asthma, 47, 849, 2010.
35. Spiegel, W.A. et al., Anaphylaxis associated with dust mite ingestion. Ann. Allergy, 72, 56, 1994.
36. Matsumoto, T., and Satoh, A., The occurrence of mitecontaining wheat �our. Pediatr. Allergy Immunol., 15, 469, 2004.
37. Sanchez-Borges, M., Capriles-Hulett, A., and CaballeroFonesca, F., Oral mite anaphylaxis (pancake syndrome) also observed in children. Ann. Allergy Asthma Immunol., 96, 755, 2006.
38. Tay, S.Y. et al., Anaphylaxis following the ingestion of �our contaminated by house dust mites—a report of two cases from Singapore. Asian Pac. J. Allergy Immunol., 26, 165, 2008.
39. van Hage-Hamsten, M., and Johansson, E., Clinical and immunologic aspects of storage mite allergy. Allergy, 53, 49, 1998.
40. Iglesias-Souto, J. et al., Oral mite anaphylaxis by Thyreophagus entomophagus in a child: A case report. Clin. Mol. Allergy, 7, 1, 2009. 41. Grif�n, P. et al., Allergenic and antigenic relationship between three species of storage mite and the house dust mite, Dermatophagoides pteronyssinus. J. Allergy Clin. Immunol., 84, 108, 1989. 42. Puerta, L. et al., Cross-reactivity among Blomia tropicalis, Lepidoglyphus destructor and Dermatophagoides farinae. J. Allergy Clin. Immunol., 87, 190, 1991. 43. Arruda, L.K. et al., Sensitisation to Blomia tropicalis in patients with asthma: Molecular cloning and expression of a major allergen (Blo t 5) with homology to Dermatophagoides pteronyssinus Der p 5. Am. J. Respir. Crit. Care Med., 155, 343, 1997. 44. Simpson, A., Arruda, K., and Chapman, M., Antigenic interrelationships among mite allergens (Blomia and Dermatophagoides spp). Clin. Rev. Allergy Immunol., 15, 461, 1997. 45. Flores, I. et al., Cloning and molecular characterization of a cDNA from Blomia tropicalis homologous to dust mite group 3 allergens (trypsin-like proteases). Int. Arch. Allergy Immunol., 130, 12, 2003. 46. Mora, C. et al., Cloning and expression of Blo t 1, a novel allergen from the dust mite Blomia tropicalis, homologous to cysteine proteases. Clin. Exp. Allergy, 33, 28, 2003. 47. Boquete, M. et al., In vivo and in vitro allergenicity of the domestic mite Chortoglyphus arcuatus. Ann. Allergy Asthma Immunol., 97, 203, 2006. 48. Chua, K.Y. et al., The Blomia tropicalis allergens. Protein Pept. Lett., 14, 325, 2007. 49. Liao, E.C. et al., Dermatophagoides pteronyssinus and Tyrophagus putrescentiae allergy in allergic rhinitis caused by cross-reactivity not dual-sensitization. J. Clin. Immunol., 30, 830, 2010. 50. O’Brien, R. et al., Immunologic responses following respiratory sensitization to house dust mite allergens in mice. Immunol. Cell Biol., 74, 174, 1996. 51. Morgan, M.S., Arlian, L.G., and Fernández-Caldas, E., Crossallergenicity of the house dust mites Euroglyphus maynei and Blomia tropicalis. Ann. Allergy Asthma Immunol., 77, 386, 1996. 52. Caraballo, L. et al., Cloning and IgE binding of a recombinant allergen from the mite Blomia tropicalis, homologous with fatty acid-binding proteins. Int. Arch. Allergy Immunol., 112, 341, 1997. 53. Chew, F.T. et al., Allergenic differences between the domestic mites Blomia tropicalis and Dermatophagoides pteronyssinus. Clin. Exp. Allergy, 29, 982, 1999. 54. Yi, F.C. et al., Identi�cation of shared and unique immunoglobulin E epitopes of the highly conserved tropomyosins in Blomia tropicalis and Dermatophagoides pteronyssinus. Clin. Exp. Allergy, 32, 1203, 2002. 55. Yi, F.C. et al., Molecular cloning of Blomia tropicalis allergens—a major source of dust mite allergens in the tropics and subtropics. In°amm. Allergy Drug Targets, 5, 261, 2006. 56. Boner, A.L. et al., Skin and serum reactivity to some storage mites in children sensitive to Dermatophagoides pteronyssinus. Ann. Allergy, 63, 82, 1989. 57. Boner, A., Pescollderungg, L., and Silverman, M., The role of house dust mite elimination in the management of childhood asthma: An unresolved issue. Allergy, 57 Suppl 74, 23, 2002. 58. Sheikh, A., and Hurwitz, B., House dust mite avoidance measures for perennial allergic rhinitis: A systematic review of ef�cacy. Br. J. Gen. Pract., 53, 318, 2003.
59. Mitchell, E.B. et al., Reduction of house dust mite allergen levels in the home: Uses of the acaracide, pirimiphos methyl. Clin. Allergy, 15, 235, 1985.
60. Bush, R.K. et al., House dust mite sublingual immunotherapy: Results of a US trial. J. Allergy Clin. Immunol., 127, 974, 2011.
61. Pulsawat, P. et al., Production and immunogenicity of hypoallergenic codon-optimized DNA vaccine encoding mature Der p 1 allergen. J. Investig. Allergol Clin. Immunol., 20, 582, 2010.
62. Hirschmann, W., Simple method for identifying live house dust mites. Pract. Pest Controller, 38, 1, 1986.
63. Fernández-Caldas, E., Iraola, V., and Carnés, J., Molecular and biochemical properties of storage mites (except Blomia species). Protein Pept. Lett., 14, 954, 2007.
64. Dunn, J.A. et al., Rapid method for the detection of storage mites in cereals: Feasibility of an ELISA based approach. Bull. Entomol. Res., 98, 207, 2008.
65. Arruda, L.K., and Chapman, M.D., A review of recent immunochemical studies of Blomia tropicalis and Euroglyphus maynei allergens. Exp. Appl. Acarol., 16, 129, 1992.
66. Jung, Y.G. et al., Comparison of the skin-prick test and Phadia ImmunoCAP as tools to diagnose house-dust mite allergy. Am. J. Rhinol. Allergy, 24, 226, 2010.
67. Simpson, A. et al., Skin test reactivity to natural and recombinant Blomia and Dermatophagoides spp. allergens among mite allergic patients in the UK. Allergy, 58, 53, 2003.
68. Puccio, F.A. et al., Importance of including Blomia tropicalis in the routine diagnosis of Venezuelan patients with persistent allergic symptoms. Allergy, 59, 753, 2004.
69. Arias-Irigoyen, J. et al., Limited IgE cross-reactivity between Dermatophagoides pteronyssinus and Glycyphagus domesticus in patients naturally exposed to both mite species. J. Allergy Clin. Immunol., 120, 98, 2007.
70. Navajas, M., and Fenton, B., The application of molecular markers in the study of diversity in acarology: A review. Exp. Appl. Acarol., 24, 751, 2000.
71. Angus, A.C., Ong, S.T., and Chew, F.T., Sequence tag catalogs of dust mite-expressed genomes: Utility in allergen and acarologic studies. Am. J. Pharmacogenomics, 4, 357, 2004.
72. Jeyaprakash, A., and Hoy, M.A., A DNA extraction procedure that allows mite specimens to be slide mounted: Phytoseiid species evaluated as a model. Exp. Appl. Acarol., 52, 131, 2010. 73. Noge, K. et al., Identi�cation of astigmatid mites using the second internal transcribed spacer (ITS2) region and its application for phylogenetic study. Exp. Appl. Acarol., 35, 29, 2005. 74. de Rojas, M. et al., Utility of ITS1–5.8S–ITS2 and 16S mitochondrial DNA sequences for species identi�cation and phylogenetic inference within the Rhinonyssus coniventris species complex (Acari: Rhinonyssidae). Parasitol. Res., 100, 1041, 2006. 75. Wong, S.F. et al., Molecular identi�cation of house dust mites and storage mites. Exp. Appl. Acarol., 55, 123, 2011. 76. Yang, B., Cai, J., and Cheng, X., Identi�cation of astigmatid mites using ITS2 and COI regions. Parasitol. Res., 108, 497, 2011. 77. de Rojas, M. et al., Phylogenetic relationships in rhinonyssid mites (Acari: Rhinonyssidae) based on mitochondrial 16S rDNA sequences. Exp. Appl. Acarol., 25, 957, 2001. 78. Ros, V.I.D., and Breeuwer, J.A.J., Spider mite (Acari: Tetranychidae) mitochondrial COI phylogeny reviewed: Host plant relationships, phylogeography, reproductive parasites and barcoding. Exp. Appl. Acarol., 42, 239, 2007. 79. Suarez-Martinez, E.B. et al., Molecular identi�cation of pathogenic house dust mites using 12S rRNA sequences. Electrophoresis, 26, 2927, 2005. 80. Miyamoto, J., and Ishii, A., and Sasa, M., A successful method for mass culture of the house dust mite, Dermatophagoides pteronyssinus (Troussart, 1897). Jpn. J. Exp. Med., 45, 133, 1975. 81. Ferrándiz, R., Casas, R., and Dreborg, S., Sensitization to Dermatophagoides siboney, Blomia tropicalis, and other domestic mites in asthmatic patients. Allergy, 51, 501, 1996. 82. Puerta, L. et al., Nucleotide sequence analysis of a complementary DNA coding for a Blomia tropicalis allergen. J. Allergy Clin. Immunol., 98, 932, 1996. 83. Wayne, R. et al., Characterization and immunobiology of house dust mite allergens. Int. Arch. Allergy Immunol., 129, 1, 2002. 84. Edston, E., and van Hage-Hamsten, M., Death in anaphylaxis in a man with house dust mite allergy. Int. J. Legal Med., 117, 299, 2003. 76 Chapter 76 - Sarcoptes (Itch Mite)
3. Arlian, L., Biology, host relations and epidemiology of Sarcoptes scabiei. Ann. Rev. Entomol., 34: 139, 1989.
4. Walton, S.F. et al., Scabies: New future for a neglected disease. Adv. Parasitol., 57, 309, 2004.
5. Kemp, D. et al., The scourge of scabies. Biologist, 49, 19, 2002.
6. Burgess, I., Biology and epidemiology of scabies. Curr. Opin. Infect. Dis., 12, 177, 1999.
7. Montesu, M.A. et al., Discoverers of the parasitic origin of scabies. Am. J. Dermatopathol., 13, 425, 1991.
8. Bates, P., Sarcoptic mange (Sarcoptes scabiei var. vulpes) in a red fox (Vulpes vulpes) population in north-west Surrey. Vet. Rec., 152, 112, 2003.
9. Dagleish, M.P. et al., Fatal Sarcoptes scabiei infection of blue sheep (Pseudois nayaur) in Pakistan. J. Wildl. Dis., 43, 512, 2007.
10. Roncalli, R.A., The history of scabies in veterinary and human medicine from biblical to modern times. Vet. Parasitol., 25, 193, 1987.
11. Green, M.S., Epidemiology of scabies. Epidemiol. Rev., 11, 126, 1989.
12. Pence, D.B., and Ueckermann, E., Sarcoptic mange in wildlife. Rev. Sci. Tech. Off. Int. Epizoot., 21, 385, 2002.
13. Chosidow, O., Scabies and pediculosis. Lancet, 355, 819, 2000.
14. Chosidow, O., Scabies. N. Engl. J. Med., 354, 1718, 2006.
15. Menzano, A., Rambozi, L., and Rossi, L., Outbreak of scabies in human beings, acquired from chamois (Rupicapra rupicapra). Vet. Rec., 155, 568, 2004.
16. Cabrera, R., and Dahl, M.V., The immunology of scabies. Semin. Dermatol., 12, 15, 1993.
17. Holt, D.C. et al., Mechanisms for a novel immune evasion strategy in the scabies mite Sarcoptes scabiei: A multigene family of inactivated serine proteases. J. Investig. Dermatol., 121, 1419, 2003.
18. Walton, S.F., The immunology of susceptibility and resistance to scabies. Parasite Immunol., 32, 532, 2010.
19. Hengge, U.R. et al., Scabies: A ubiquitous neglected skin disease. Lancet Infect. Dis., 6, 769, 2006.
20. Anbar, T.S. et al., Scaly scalp associated with crusted scabies: Case series. Dermatol. Online J., 13, 18, 2007.
21. Parish, L.C., and Lomholt, G., Crusted scabies alias Norwegian scabies. Int. J. Dermatol., 15, 747, 1976.
22. Guggisberg, D. et al., Norwegian scabies in a patient with acquired immunode�ciency syndrome. Dermatology, 197, 306, 1998.
23. Van Der Wal, V.B. et al., Crusted (Norwegian) scabies in a patient with dystrophic epidermolysis bullosa. Br. J. Dermatol., 141, 918, 1999.
24. Walton, S.F. et al., New insights into disease pathogenesis in crusted (Norwegian) scabies: The skin immune response in crusted scabies. Br. J. Dermatol., 158, 1247, 2008.
25. Roberts, L.J. et al., Crusted scabies: Clinical and immunological �ndings in seventy-eight patients and a review of the literature. J. Infect., 50, 375, 2005.
26. Karthikeyan, K., Crusted scabies. Indian J. Dermatol. Venereol. Leprol., 75, 340, 2009.
27. Sampathkumar, K. et al., Norwegian scabies in a renal transplant patient. Indian J. Nephrol., 20, 89, 2010.
28. Subramaniam, G. et al., Norwegian scabies in a malnourished young adult: A case report. J. Infect. Dev. Ctries., 4, 349, 2010.
29. Heukelbach, J., and Feldmeier, H., Scabies. Lancet, 367, 1767, 2006. 30. Guldbakke, K.K., and Khachemoune, A., Crusted scabies: A clinical review. J. Drugs Dermatol., 5, 221, 2006. 31. Smets, K., and Vercruysse, J., Evaluation of different methods for the diagnosis of scabies in swine. Vet. Parasitol., 90, 137, 2000. 32. Taplin, D. et al., Comparison of crotamiton 10% cream (Eurax) and permethrin 5% cream (Elimite) for the treatment of scabies in children. Pediatr. Dermatol., 7, 67, 1990. 33. Pruksachatkunakorn, C., Damrongsak, M., and Sinthupuan, S., Sulfur for scabies outbreaks in orphanages. Pediatric Dermatol., 19, 448, 2002. 34. Chouela, E.N. et al., Equivalent therapeutic ef�cacy and safety of ivermectin and lindane in the treatment of human scabies. Arch. Dermatol., 135, 651, 1999. 35. Jin, S.P. et al., Scabies in a 2-month-old infant successfully treated with lindane. Ann. Dermatol., 21, 200, 2009. 36. Scott, G.R., and Chosidow, O., European guideline for the management of scabies. Int. J. STD AIDS, 22, 301, 2010. 37. Leone, P.A., Scabies and pediculosis pubis: An update of treatment regimens and general review. Clin. Infect. Dis., 44, S153, 2007. 38. Huffam, S.E., and Currie, B.J., Ivermectin for Sarcoptes scabiei hyperinfestation. Int. J. Infect. Dis., 2, 152, 1998. 39. Santoro, A.F., Rezac, M.A., and Lee, J.B., Current trend in ivermectin usage for scabies. J. Drugs Dermatol., 2, 397, 2003. 40. Alberici, F. et al., Ivermectin alone or in combination with benzyl benzoate in the treatment of human immunode�ciency virus associated scabies. Br. J. Dermatol., 142, 969, 2000. 41. Aubin, F., and Humbart, P., Ivermectin for crusted (Norwegian) scabies. N. Eng. J. Med., 333, 26, 1995. 42. Currie, B.J. et al., First documentation of in vivo and in vitro ivermectin resistance in Sarcoptes scabiei. Clin. Infect. Dis., 39, e8, 2004. 43. Currie, B.J., and McCarthy, J.S., Permethrin and ivermectin for scabies. N. Eng. J. Med., 362, 717, 2010. 44. Walton, S.F., Myerscough, M.R., and Currie, B.J., Studies in vitro on the relative ef�cacy of current acaricides for Sarcoptes scabiei var. hominis. Trans. R. Soc. Trop. Med. Hyg., 94, 92, 2000. 45. Meinking, T.L., and Elgart, W.E., Scabies therapy for the millenium. Pediatric Dermatol., 2, 154, 2000. 46. Scheinfeld, N., Controlling scabies in institutional settings: A review of medications, treatment models, and implementation. Am. J. Clin. Dermatol., 5, 31, 2004. 47. Karthikeyan, K., Treatment of scabies: Newer perspectives. Postgrad. Med. J., 81, 7, 2005. 48. Mumcuoglu, K.Y., and Gilead, L., Treatment of scabies infestations. Parasite, 15, 248, 2008. 49. Gould, D., Prevention, control and treatment of scabies. Nurs. Stand., 25, 42, 2010. 50. Gilmore, S.J., Control strategies for endemic childhood scabies. PLoS One, 6, e15990, 2011. 51. McCarthy, J.S. et al., Scabies: More than just an irritation. Postgrad. Med. J., 80, 382, 2004. 52. Walton, S.F., and Currie, B.J., Problems in diagnosing scabies, a global disease in human and animal populations. Clin. Microbiol. Rev., 20, 268, 2007. 53. Johnston, G., and Sladden, M., Scabies: Diagnosis and treatment. BMJ, 331, 619, 2005. 54. Leibowitz, M., Shave biopsy an underutilised technique in the diagnosis of scabies. N. Z. Med. J., 107, 111, 1994. 55. Zahler, M. et al., Molecular analyses suggest monospeci�city of the genus Sarcoptes (Acari: Sarcoptidae). Int. J. Parasitol., 29, 759, 1999.
56. Katsumata, K., and Katsumata, K., Simple method of detecting Sarcoptes Scabiei var hominis mites among bedridden elderly patients suffering from severe scabies infestation using an adhesive-tape. Intern. Med., 45, 857, 2006.
57. Turan, E. et al., The detection of Sarcoptes scabiei in human skin by in vivo confocal microscopy. Eur. J. Dermatol., 21, 1004, 2011.
58. Dupuy, A. et al., Accuracy of standard dermoscopy for diagnosing scabies. J. Am. Acad. Dermatol., 59, 530, 2008.
59. Grif�n, J.R., and Newman, C.C., Clinical, dermatoscopic, and microscopic �ndings of iInfestation with Sarcoptes scabiei var. hominis. Mayo Clin. Proc., 86, e47, 2011.
60. Walter, B. et al., Comparison of dermoscopy, skin scraping, and the adhesive tape test for the diagnosis of scabies in a resource-poor setting. Arch. Dermatol., 147, 468, 2011.
61. Skerratt, L.F. et al., The mitochondrial 12S gene is a suitable marker of populations of Sarcoptes scabiei from wombats, dogs and humans in Australia. Parasitol. Res., 88, 376, 2002.
62. Cruickshank, R.H., Molecular markers for the phylogenetics of mites and ticks. Syst. Appl. Acarol., 7, 3, 2002.
63. Alasaad, S. et al., Sarcoptes mite from collection to DNA extraction: The lost realm of the neglected parasite. Parasitol. Res., 104, 723, 2008.
64. Alasaad, S. et al., HotSHOT Plus ThermalSHOCK, a new and ef�cient technique for preparation of PCR-quality Sarcoptes mite genomic DNA. Parasitol. Res., 103, 1455, 2008.
65. Alasaad, S. et al., Skin-scale genetic structure of Sarcoptes scabiei populations from individual hosts: Empirical evidence from Iberian ibex-derived mites. Parasitol. Res., 104, 101, 2008.
66. Alasaad, S. et al., Effectiveness of postponed isolation (postfrozen isolation) method for PCR-quality Sarcoptes mite gDNA. Exp. Appl. Acarol., 47, 173, 2009. 67. Alasaad, S. et al., Is ITS-2 rDNA suitable marker for genetic characterization of Sarcoptes mites from different wild animals in different geographic areas? Vet. Parasitol., 159, 181, 2009. 68. Alasaad, S. et al., Sarcoptes-World Molecular Network (Sarcoptes-WMN): Integrating research on scabies. Int. J. Infect. Dis., 15, e294, 2011. 69. Berrilli, F., D’Amelio, S., and Rossi, L., Ribosomal and mitochondrial DNA sequence variation in Sarcoptes mites from different hosts and geographical regions. Parasitol. Res., 88, 772, 2002. 70. Walton, S.F., Currie, B.J., and Kemp, D.J., A DNA �ngerprinting system for the ectoparasite Sarcoptes scabiei. Mol. Biochem. Parasitol., 85, 187, 1997. 71. Walton, S.F. et al., Genetically distinct dog-derived and human-derived Sarcoptes scabiei in scabies-endemic communities in northern Australia. Am. J. Trop. Med. Hyg., 61, 542, 1999. 72. Walton, S.F. et al., Crusted scabies: A molecular analysis of Sarcoptes scabiei variety hominis population from patients with repeated infestations. Clin. Infect. Dis., 29, 1226, 1999. 73. Walton, S.F. et al., Genetic epidemiology of Sarcoptes scabiei (Acari: Sarcoptidae) in northern Australia. Int. J. Parasitol., 34, 839, 2004. 74. Soglia, D. et al., Microsatellites as markers for comparison among different populations of Sarcoptes scabiei. It. J. Anim. Sci., 7, 214, 2007. 75. Bezold, G. et al., Hidden scabies: Diagnosis by polymerase chain reaction. Br. J. Dermatol., 144, 614, 2001. 76. Soglia, D. et al., Two simple techniques for the safe Sarcoptes collection and individual mite DNA extraction. Parasitol. Res., 105, 1465, 2009.
Section VII
Arthropoda: Insecta 77 Chapter 77 - Calliphoridae, Oestridae, and Sarcophagidae (Myiasis-Causing Flies)
1. Noutsis, C., and Millikan, L.E., Myiasis. Dermatol. Clin., 12, 729, 1994.
2. Millikan, L.E., Myiasis. Clin. Dermatol., 17, 191, 1999.
3. Stevens, J.R., and Wallman, J.F., The evolution of myiasis in humans and other animals in the Old and New Worlds (part I): Phylogenetic analyses. Trends Parasitol., 22, 129, 2006.
4. Stevens, J.R. et al., The evolution of myiasis in humans and other animals in the Old and New Worlds (part II): Biological and life-history studies. Trends Parasitol., 22, 181, 2006.
5. Erzinçlioglu, Y.Z., The origin of parasitism in blow�ies. Brit. J. Entomol. Nat. Hist., 2, 125, 1989.
6. Rognes, K., The Calliphoridae (blow�ies) (Diptera: Oestroidea) are not a monophyletic group. Cladistic, 13, 27, 1997.
7. Stevens, J.R., The evolution of myiasis in blow�ies (Calliphoridae). Int. J. Parasitol., 33, 1105, 2003.
8. Wallman, J.F., Third-instar larvae of common carrion-breeding blow�ies of the genus Calliphora in South Australia. Invertebr. Taxon, 15, 37, 2001.
9. Alexander, J.L., Screwworms. J. Am. Vet. Med. Assoc., 228, 357, 2006.
10. Stevens, J., and Wall, R., Species, subspecies and hybrid populations of the blow�ies Lucilia cuprina and Lucilia sericata (Diptera: Calliphoridae). Proc. Biol. Sci., 263, 1335, 1996. 11. Stevens, J., and Wall, R., The evolution of ectoparasitism in the genus Lucilia (Diptera: Calliphoridae). Int. J. Parasitol., 27, 51, 1997. 12. Stevens, J., and Wall, R., Genetic relationships between blow�ies (Calliphoridae) of forensic importance. Forensic Sci. Int., 120, 116, 2001. 13. Norris, K.R., Evidence for the multiple exotic origin of Australian populations of the sheep blow�y, Lucilia cuprina (Wiedemann) (Diptera: Calliphoridae). Aust. J. Zool., 38, 635, 1990. 14. Hall, M.J.R. et al., Old World screwworm �y, Chrysomya bezziana, occurs as two geographical races. Med. Vet. Entomol., 15, 393, 2001. 15. Wyss, J., Screwworm eradication in the Americas. Ann. N.Y. Acad. Sci., 916, 186, 2000. 16. Coronado, A., and Kowalski, A., Current status of the New World screwworm Cochliomyia hominivorax in Venezuela. Med. Vet. Entomol., 23 (Suppl 1), 106, 2009. 17. Pape, T., Phylogeny of Oestridae (Insecta: Diptera). Syst. Entomol., 26, 133, 2001. 18. Hall, M.J.R., and Wall, R., Myiasis of humans and domestic animals. Adv. Parasitol., 35, 257, 1995. 19. Powers, N., and Yorgensen, M., Myiasis in humans: An overview and a report of two cases in the Republic of Panama. Mil. Med., 161, 495, 1996. 20. MacNamara, A., and Durham, S., Dermatobia hominis in the accident and emergency department: “I’ve got you under my skin.” J. Accid. Emerg. Med., 14, 179, 1997. 21. Bapat, S., Neonatal myiasis. Pediatrics, 106, e6, 2000. 22. Jiang, C., A collective analysis on 54 cases of human myiasis in China from 1995-2001. Chin. Med. J. (Engl), 115, 1445, 2002. 23. Cestari, T.F., Pessato, S., and Ramos-e-Silva, M., Tungiasis and myiasis. Clin, Dermatol., 25, 158, 2007. 24. Paris, L.A. et al., Pin-site myiasis: A rare complication of a treated open fracture of tibia. Surg. Infect. (Larchmt), 9, 403, 2008. 25. Otranto, D., The immunology of myiasis: Parasite survival and host defense strategies. Trends Parasitol., 17, 176, 2001. 26. Robbins, K., and Khachemoune, A., Cutaneous myiasis: A review of the common types of myiasis. Int. J. Dermatol., 49, 1092, 2010. 27. Gordon, P. et al., Cutaneous myiasis due to Dematobia hominis: A report of six cases. Brit. J. Dermatol., 132, 811, 1995. 28. Kpea, N., and Zywocinski, C., Flies in the �esh: A case report and review of cutaneous myiasis. Cutis, 55, 47, 1995. 29. Swetter, S., Stewart, M., and Smoller, B., Cutaneous myiasis following travel to Belize. Int. J. Dermatol., 35, 118, 1996. 30. Tsuda, S. et al., Furuncular cutaneous myiasis caused by Dermatobia hominis larvae following travel to Brazil. Int. J. Dermatol., 35, 121, 1996. 31. Shorter, N. et al., Furuncular cuterebrid myiasis. J. Pediatr. Surg., 22, 1511, 1997. 32. Hasegawa, M. et al., An imported case of furuncular myiasis due to Cordylobia anthropophaga which emerged in Japan. Br. J. Dermatol., 143, 912, 2000. 33. Jelinek, T. et al., Cutaneous myiasis: Review of 13 cases of travelers returning from tropical countries. Int. J. Dermatol., 39, 689, 2000. 34. Robert, L., and Yelton, J., Imported furuncular myiasis in Germany. Mil. Med., 167, 990, 2002. 35. Haruki, K. et al., Myiasis with Dermatobia hominis in a traveler returning from Costa Rica: Review of 33 cases imported from South America to Japan. J. Travel Med., 12, 285, 2005.
36. Kokcam, I., and Saki, C.E., A case of cutaneous myiasis caused by Wohlfahrtia magni�ca. J. Dermatol., 32, 459, 2005.
37. Curtis, S.J. et al., Case of the month: Cutaneous myiasis in a returning traveler from the Algarve: First report of tumbu maggots, Cordylobia anthropophaga, acquired in Portugal. Emerg. Med. J., 23, 236, 2006.
38. Delshad, E. et al., Cuterebra cutaneous myiasis: Case report and world literature review. Int. J. Dermatol., 47, 363, 2008.
39. McGraw, T.A., and Turiansky, G.W., Cutaneous myiasis. J. Am. Acad. Dermatol., 58, 907, 2008; quiz 58, 927, 2008.
40. Krajewski, A. et al., Cutaneous myiasis. J. Plast Reconstr. Aesthet. Surg., 62, e383, 2009.
41. Sherman, R.A., Wound myiasis in urban and suburban United States. Arch. Intern. Med., 160, 2004, 2000.
42. Hemmings, S.C., Matthews, K.J., and Alexander, J., Human myiasis in western Jamaica: Five years after the implementation of a screwworm eradication programme. West Indian Med. J., 56, 341, 2007.
43. Sesterhenn, A.M. et al., Cutaneous manifestation of myiasis in malignant wounds of the head and neck. Eur. J. Dermatol., 19, 64, 2009.
44. Lane, J.E. et al., Furuncular myiasis secondary to Dermatobia hominis. J. Drugs Dermatol., 4, 365, 2005.
45. Fujisaki, R. et al., Exotic myiasis caused by 19 larvae of Cordylobia anthropophaga in Namibia and identi�ed using molecular methods in Japan. Trans. Royal Soc. Trop. Med. Hyg., 102, 599, 2008.
46. Geary, M.J. et al., Exotic myiasis with Lund’s �y (Cordylobia rodhaini). Med. J. Aust., 171, 654, 1999.
47. Royce, L.A. et al., Recovery of a second instar Gasterophilus larva in a human infant: A case report. Am. J. Trop. Med. Hyg., 60, 403, 1999.
48. Karaman, E. et al., Otomyiasis by Wohlfahrtia magni�ca. J. Craniofac. Surg., 20, 2123, 2009.
49. Tuygun, N. et al., Furuncular myiasis in a child caused by Wohlfahrtia magni�ca (Diptera: Sarcophagidae) associated with eosinophilia. Turk. J. Pediatr., 51, 279, 2009.
50. Romero-Cabello, R. et al., Cutaneous myiasis caused by Chrysomya bezziana larvae, Mexico. Emerg. Infect. Dis., 16, 2014, 2010.
51. Starr, J. et al., Myiasis due to Hypoderma lineatum infection mimicking the hypereosinophilic syndrome. Mayo Clin. Proc., 75, 755, 2000.
52. Syrdalen, P., Nitter, T., and Mehl, R., Ophthalmomyiasis interna posterior: Report of case caused by the reindeer warble �y larva and review of previous reported cases. Br. J. Ophthalmol., 66, 589, 1982.
53. Passos, M.R. et al., Vulvar myiasis. Infect. Dis. Obstet. Gynecol., 6, 69, 1998.
54. Droma, E.B. et al., Oral myiasis: A case report and literature review. Oral Surg. Oral. Med. Oral Pathol. Oral Radiol. Endod., 103, 92, 2007.
55. Lagacé-Wiens, P.R. et al., Human ophthalmomyiasis interna caused by Hypoderma tarandi, Northern Canada. Emerg. Infect. Dis., 14, 64, 2008.
56. Terterov, S. et al., Posttraumatic human cerebral myiasis. World Neurosurg., 73, 557, 2010.
57. Ng, K.H. et al., A case of oral myiasis due to Chrysomya bezziana. Hong Kong Med. J., 9, 454, 2003.
58. Sharma, A., and Hedge, A., Primary oral myiasis due to Chrysomya bezziana treated with Ivermectin. A case report. J. Clin. Pediatr. Dent., 34, 259, 2010. 59. Gealh, W.C. et al., Treatment of oral myiasis caused by Cochliomyia hominivorax: Two cases treated with ivermectin. Br. J. Oral Maxillofac. Surg., 47, 23, 2009. 60. Das, A. et al., Accidental intestinal myiasis caused by genus Sarcophaga. Indian J. Med. Microbiol., 28, 176, 2010. 61. Brewer, T. et al., Bacon therapy and furuncular myiasis. JAMA, 270, 2087, 1993. 62. Caissie, R. et al., Cutaneous myiasis: Diagnosis, treatment, and prevention. J. Oral Maxillofac. Surg., 66, 560, 2008. 63. Sherman, R.A. et al., Medicinal maggots: An ancient remedy for some contemporary af�ictions. Annu. Rev. Entomol., 45, 55, 2000. 64. Kitching, J., Tropical myiasis: An unwanted holiday souvenir. J. Accid. Emerg. Med., 14, 178, 1997. 65. Lucchina, L., Wilson, M., and Drake, L., Dermatology and the recently returned traveler: Infectious diseases with dermatologic manifestations. Int. J. Dermatol., 36, 167, 1997. 66. Diaz, J.H., The epidemiology, diagnosis, management, and prevention of ectoparasitic diseases in travelers. J. Travel Med., 13, 100, 2006. 67. Hakeem, M.J., and Bhattacharyya, D.N., Exotic human myiasis. Travel Med. Infect. Dis., 7, 198, 2009. 68. Bowry, R., and Cottingham, R., Use of ultrasound to aid management of late presentation of Dermatobia hominis larva infestation. J. Accid. Emerg. Med., 14, 177, 1997. 69. Greenberg, B., and Singh, D., Species identi�cation of calliphorid (Diptera) eggs. Med. Entomol., 32, 21, 1995. 70. Moiré, N. et al., Sequencing and gene expression of hypodermins A, B, C in larval stages of Hypoderma lineatum. Mol. Biochem. Parasitol., 66, 233, 1994. 71. Boulard, C., Villejoubert, C., and Moiré, N., Cross-reactive, stage-speci�c antigens in the Oestridae family. Vet. Res., 27, 535, 1996. 72. Goddard, P., Bates, P., and Webster, K.A., Evaluation of a direct ELISA for the serodiagnosis of Oestrus ovis infections in sheep. Vet. Rec., 144, 497, 1999. 73. Figarola, J.L. et al., Identi�cation of screwworm, Cochliomyia hominivorax (Coquerel) (Diptera: Calliphoridae), with a monoclonal antibody-based enzyme-linked immunosorbent assay (MAb-ELISA). Vet. Parasitol., 102, 341, 2001. 74. Kocher T.D., and Xiong, B., Comparison of mitochondrial DNA sequences of seven morphospecies of black �ies (Diptera: Simulidae). Genome, 34, 306, 1991. 75. Sperling, F.A.H., Anderson, G.S., and Hickey, D.A., A DNAbased approach to the identi�cation of insect species used for post-mortem interval estimation. J. Forensic Sci., 39, 418, 1994. 76. Infante-Malachias, M.E. et al., Random ampli�ed polymorphic DNA of screwworm �y populations (Diptera: Calliphoridae) from Southeastern Brazil and Northern Argentina. Genome, 42, 772, 1999. 77. Caterino, M.S., Cho, S., and Sperling, F.A.H., The current state of insect molecular systematics. Annu. Rev. Entomol., 45, 1, 2000. 78. Nirmala, X. et al., Molecular phylogeny of Calyptratae (Diptera: Brachycera): The evolution of 18S and 16S ribosomal rDNAs in higher dipterans and their use in phylogenetic inference. Insect Mol. Biol., 10, 475, 2001. 79. Hillis, D.M., and Dixon, M.T., Ribosomal DNA: Molecular evolution and phylogenetic inference. Q. Rev. Biol., 66, 411, 1991. 80. Lunt, D.H. et al., The insect cytochrome oxidase I gene: Evolutionary patterns and conserved primers for phylogenetic studies. Ins. Mol. Biol., 5, 153, 1996.
81. Wells, J.D., and Sperling, F.A., Molecular phylogeny of Chrysomya albiceps and C. ru�facies. J. Med. Entomol., 36, 222, 1999. 82. Wells, J.D. et al., DNA-based identi�cation and molecular systematics of forensically important Sarcophagidae (Diptera). J. Forensic Sci., 46, 1098, 2001.
83. Wells, J.D. et al., Human and insect mitochondrial DNA analysis from maggots. J. Forensic Sci., 46, 685, 2001.
84. Lessinger, A.C., and Azeredo-Espin, A.M.L., Evolution and structural organisation of mitochondrial DNA control region of myiasis-causing �ies. Med. Vet. Entomol., 14, 71, 2000.
85. Lessinger, A.C. et al., The mitochondrial genome of the primary screwworm �y Cochliomyia hominivorax (Diptera: Calliphoridae). Insect Mol. Biol., 9, 521, 2000.
86. Otranto, D. et al., Differentiation by polymerase chain reaction-restriction fragment length polymorphism of some Oestridae larvae causing myiasis. Vet. Parasitol., 90, 305, 2000.
87. Otranto D., and Stevens, J.R., Molecular approaches to the study of myiasis-causing larvae. Int. J. Parasitol., 32, 1345, 2002.
88. Otranto, D. et al., Utility of mitochondrial and ribosomal genes for the differentiation and phylogenesis of species of gastrointestinal bot�ies. J. Econ. Entomol., 98, 2235, 2005.
89. Simon, C. et al., Evolution, weighting and phylogenetic utility of mitochondrial gene sequences and a compilation of conserved polymerase chain reaction primers. Ann. Entomol. Soc. Am., 87, 651, 1994. 90. Taylor, D.B., Szalanski, A.L., and Peterson, R.D., Mitochondrial DNA variation in screwworm. Med. Vet. Entomol., 10, 161, 1996. 91. Litjens, P., Lessinger, A.C., and Azeredo-Espin, A.M.L., Characterization of the screwworm �ies Cochliomyia hominivorax and Cochliomyia macellaria by PCRRFLP of mitochondrial DNA. Med. Vet. Entomol., 15, 183, 2001. 92. Wallman, J.F., and Donnellan, S.C., The utility of mitochondrial DNA sequences for the identi�cation of forensically important blow�ies (Diptera: Calliphoridae) in southeastern Australia. Forensic Sci. Int., 120, 60, 2001. 93. Wallman, J.F. et al., Molecular systematics of Australian carrion-breeding blow�ies (Diptera: Calliphoridae) based on mitochondrial DNA. Invert. Syst., 19, 1, 2005. 94. Zhang, D.X., and Hewitt, G.M., Assessment of the universality and utility of a set of conserved mitochondrial COI primers in insects. Insect Mol. Biol., 6, 143, 1996. 95. Li Loong, P.T., Lui, H., and Buck, H.W., Cutaneous myiasis: A simple and effective technique for extraction of Dermatoba hominis larvae. Int. J. Dermatol., 31, 657, 1992. 96. Olumide, Y., Cutaneous miasis: A simple and effective technique for extraction of Dermatobia hominis larvae. Int. J. Dermatol., 33, 148, 1994. 97. Adams, Z.J.O., and Hall, M.J.R., Methods used for killing and preservation of blow�y larvae, and their effect on postmortem larval length. Forensic Sci. Int., 138, 50, 2003. 78 Chapter 78 - Cimex (Bedbug)
1. Thomas, I., Kihiczak, G.G., and Schwartz, R.A., Bed bug bites: A review. Int. J. Dermal., 43, 430, 2004.
2. Harlan, H.J., Bed bugs 101: The basics of Cimex lectularius. Am. Entomol., 52, 99, 2006.
3. Kolb, A. et al., Bedbugs. Dermatol. Ther., 22, 347, 2009.
4. Reinhardt, K., and Siva-Jothy, M.T., The biology of bedbugs (Cimicidae). Ann. Rev. Entomol., 52, 351, 2007.
5. How, Y.F., and Lee, C.Y., Fecundity, nymphal development and longevity of �eld-collected tropical bedbugs, Cimex hemipterus. Med. Vet. Entomol., 24, 108, 2010.
6. Araujo, R.N. et al., The feeding process of Cimex lectularius (Linnaeus 1758) and Cimex hemipterus (Fabricius 1803) on different bloodmeal sources. J. Insect Physiol., 55, 1151, 2009.
7. Ryan, N., Peters, B., and Miller, P., A survey of bed bugs in short-stay lodges. NSW Public Health Bull., 15, 215, 2004.
8. Hwang, S.W. et al., Bed bug infestations in an urban environment. Emerg. Infect. Dis., 11, 533, 2005.
9. Wang, C. et al., Characteristics of Cimex lectularius (Hemiptera: Cimicidae), infestation and dispersal in a highrise apartment building. J. Econ. Entomol., 103, 172, 2010.
10. Boase, C.J., Bedbugs: Back from the brink. Pest Outl., 12, 159, 2001.
11. Myamba, J. et al., Pyrethroid resistance in tropical bedbugs, Cimex hemipterus, associated with use of treated bed nets. Med. Vet. Entomol., 16, 448, 2002. 12. Doggett, S.L., Geary, M.J., and Russell, R.C., The resurgence of bed bugs in Australia, with notes on their ecology and control. Environ. Health, 4, 30, 2004. 13. Ter Poorten, M.C., and Prose, N.S., The return of the common bedbug. Pediatr. Dermatol., 22, 183, 2005. 14. Karunaratne, S.H.P.P. et al., Insecticide resistance in the tropical bedbug Cimex hemipterus. Pest Biochem. Physiol., 88, 102, 2007. 15. Romero, A. et al., Insecticide resistance in the bed bug: A factor in the pest’s sudden resurgence? J. Med. Entomol., 44, 175, 2007. 16. Lee, I.Y. et al., Reemergence of the bedbug Cimex lectularius in Seoul, Korea. Korean J. Parasitol., 46, 269, 2008. 17. Anderson, A.L., and Lef�er, K., Bedbug infestations in the news: A picture of an emerging public health problem in the United States. J. Environ. Health, 70, 24, 2008. 18. Levy Bencheton, A. et al., Resurgence of bedbugs in southern France: A local problem or the tip of the iceberg? J. Eur. Acad. Dermatol. Venereol., 25, 599, 2011. 19. Manuel, J., Invasion of the bedbugs. Environ. Health Perspect., 118, A429, 2010. 20. Zhu, F. et al., Widespread distribution of knockdown resistance mutations in the bed bug, Cimex lectularius (Hemiptera: Cimicidae), populations in the United States. Arch. Insect Biochem. Physiol., 73, 245, 2010. 21. Stibich, A.S., Carbonaro, P.A., and Schwartz, R.A., Insect bite reactions: An update. Dermatology, 202, 193, 2001. 22. Doggett, S.L., and Russell, R., Bed bugs—What the GP needs to know. Aust. Fam. Physician, 38, 880, 2009. 23. Goddard, J., and deShazo, R., Bed bugs (Cimex lectularius) and clinical consequences of their bites. J. Am. Med. Assoc., 301, 1358, 2009. 24. Lorenzen, R., Bedbugs and health. CMAJ, 181, 617, 2009. 25. Reinhardt, D. et al., Sensitivity to bites by the bedbug, Cimex lectularius. Med. Vet. Entomol., 23, 163, 2009. 26. Scarupa, M.D., and Economides, A., Bedbug bites masquerading as urticaria. J. Allergy Clin. Immunol., 117, 1508, 2006. 27. Stucki, A., and Ludwig, R., Images in clinical medicine: Bedbug bites. N. Engl. J. Med., 359, 1047, 2008. 28. Delaunay, P. et al., Bedbugs and infectious diseases. Clin. Infect. Dis., 52, 200, 2011. 29. Reinhardt, K., Wong, C.H., and Georgiou, A.S., Detection of seminal �uid proteins in the bed bug, Cimex lectularius, using two-dimensional gel electrophoresis and mass spectrometry. Parasitology, 136, 283, 2009. 30. Abdel-Naser, M.B. et al., Patients with papular urticaria have IgG antibodies to bedbug (Cimex lectularius) antigens. Parasitol. Res., 98, 550, 2006. 31. Leverkus, M. et al., Bullous allergic hypersensitivity to bed bug bites mediated by IgE against salivary nitrophorin. J. Invest. Dermatol., 126, 91, 2006. 32. Pritchard, M.J., and Hwang, S.W., Cases: Severe anemia from bedbugs. CMAJ, 181, 287, 2009. 33. Moore, D.J., and Miller, D.M., Laboratory evaluations of insecticide product ef�cacy for control of Cimex lectularius. J. Econ. Entomol., 99, 2080, 2006. 34. Kweka, E.J. et al., Insecticides susceptibility status of the bedbugs (Cimex lectularius) in a rural area of Magugu, Northern Tanzania. J. Glob. Infect. Dis., 1, 102, 2009. 35. Gangloff-Kaufmann, J. et al., Bed bugs in America: A pest management industry survey. Am. Entomol., 52, 105, 2006. 36. Naylor, R.A., and Boase, C.J., Practical solutions for treating laundry infested with Cimex lectularius (Hemiptera: Cimicidae). J. Econ. Entomol., 103, 136, 2010.
37. Potter, M.F., A bed bug state of mind: Emerging issues in bed bug management. Pest Control Technol., 33, 82, 2005.
38. Delaunay, P. et al., Bedbugs and healthcare-associated dermatitis, France. Emerg. Infect. Dis., 15, 989, 2009.
39. Rossi, L., and Jennings, S., Bed bugs: A public health problem in need of a collaborative solution. J. Environ. Health, 72, 34, 2010.
40. Szalanski, A.L. et al., Time course analysis of bed bug, Cimex lectularius L. (Hemiptera: Cimicidae) blood meals using PCR. J. Agr. Urban Entomol., 23, 237, 2006. 41. Szalanski, A.L. et al. Mitochondrial and ribosomal internal transcribed spacer (ITS1) diversity of the bed bug Cimex lectularius L. (Heteroptera: Cimicidae). J. Med. Entomol., 45, 229, 2008. 42. Szalanski, A.L. et al. Isolation and characterization of human DNA from the bed bug, Cimex lectularius L. (Heteroptera: Cimicidae). J. Agr. Urban Entomol., 23, 189, 2006. 43. Panagiotakopulu, E., and Buckland, P.C., Cimex lectularius L., the common bed bug from Pharaonic Egypt. Antiquity, 73, 908, 1999. 79 Chapter 79 - Pediculus (Body and Head Lice)
22. Yong, Z. et al., The geographical separation of human lice preceded that of Pediculus humanus capitis and Pediculus humanus humanus. C. R. Biol., 326, 565, 2003.
23. Raoult, D. et al., Evidence for louse-transmitted diseases in soldiers of Napoleon’s grand army in Vilnius. J. Infect. Dis., 193, 112, 2006.
24. Odronitz, F., Becker, S., and Kollmar, M., Reconstructing the phylogeny of 21 completely sequenced arthropod species based on their motor proteins. BMC Genomics, 10, 173, 2009. 27. Desalle, R. et al., DNA sequences from a fossil termite in Oligo-Miocene amber and their phylogenetic implications. Science, 257, 1933, 1992. 28. Tamura, K. et al., MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol., 28, 2731, 2011. 80 Chapter 80 - Phthirus (Crab Louse)
1. Burkhart, C.N., Gunning, W., and Burkhart, C.G., Scanning electron microscopic examination of the egg of the pubic louse (Anoplura: Pthirus pubis). Int. J. Dermatol., 39, 201, 2000.
2. Anderson, A.L., and Chaney, E., Pubic Lice (Pthirus pubis): History, biology and treatment vs. knowledge and beliefs of US College students. Int. J. Env. Res. Public Health, 6, 592, 2009.
3. Ko, C.J., and Elston, D.M., Pediculosis. J. Am. Acad. Dermatol., 50, 1, 2004.
4. Puri, P.K. et al., The histopathology of Phthirus pubis. J. Cutan. Pathol., 36, 80, 2009.
5. Routh, H.B. et al., Ectoparasites as sexually transmitted diseases. Semin. Dermatol., 13, 243, 1994.
6. Skinner, C.J., Viswalingam, N.D., and Goh, B.T., Phthirus pubis infestation of the eyelids: A marker for sexually transmitted diseases. Int. J. STD AIDS, 6, 451, 1995.
7. Mimouni, D. et al., Secular trends in epidemiology of pediculosis capitis and pubis among Israeli soldiers: A 27-year follow-up. Int. J. Derm., 40, 637, 2001.
8. Pierzchalski, J.L., Bretl, D.A., and Matson, S.C., Phthirus pubis as a predictor for chlamydia infections in adolescents. Sex. Transm. Dis., 29, 331, 2002.
9. Varela, J.A. et al., Pthirus pubis in a sexually transmitted diseases unit: A study of 14 years. Sex. Transm. Dis., 30, 292, 2003.
10. Orion, E., Hagit, M., and Wolf, R., Ectoparasitic sexually transmitted diseases: Scabies and Pediculosis. Clin. Dermatol., 22, 513, 2004.
11. Mimouni, D. et al., Seasonality trends of Pediculosis capitis and Phthirus pubis in a young adult population: Follow-up of 20 years. J. Eur. Acad. Dermatol. Venereol., 16, 257, 2002.
12. Flinders, D.C., and DeSchweinitz, P., Pediculosis and scabies. Am. Fam. Phys., 69, 341, 2004. 13. Poudel, S.K., and Barker, S.C., Infestation of people with lice in Kathmandu and Pokhara, Nepal. Med. Vet. Entomol., 18, 212, 2004.
14. Pakeer, O. et al., Four cases of pediculosis caused by Pthirus pubis Linnaeus, 1758 (Diptera: Anoplura) from peninsular Malaysia. Trop. Biomed., 24, 101, 2007.
15. Lee, H.L. et al., Review of forensically important entomological specimens in the period of 1972–2002. Trop. Biomed., 21, 69, 2004.
16. Kenward, H., Pubic lice in Roman and Medieval Britain. Trends Parasitol., 17, 167, 2001.
17. Rick, F.M. et al., Crab louse infestation in pre-Columbian America. J. Parasitol., 88, 1266, 2002.
18. Fornaciari, G. et al., ‘Royal’ pediculosis in Renaissance Italy: Lice in the mummy of the King of Naples Ferdinand II of Aragon (1467–1496). Mem. Inst. Oswaldo Cruz, 104, 671, 2009. 19. Manjunatha, N.P. et al., Pediculosis pubis: Presentation to ophthalmologist as phthriasis palpebrarum associated with corneal epithelial keratitis. Int. J. STD AIDS, 17, 424, 2006. 20. Ngai, J.W., Yuen, H.K., and Li, F.C., An unusual case of eye itchiness. Hong Kong Med. J., 14, 414, 2008. 21. Pinckney, J. 2nd. et al., Phthiriasis palpebrarum: A common culprit with uncommon presentation. Dermatol. Online J., 14, 7, 2008. 22. Niazi, M.K., and Arain, M.A., Phthiriasis palpebrarum. J. Coll. Physicians Surg. Pak., 19, 589, 2009. 23. Klaus, S., Shvil, Y., and Mumcuoglu, K.Y., Generalized infestation of a 31/2-year-old girl with the pubic louse. Pediatr. Dermatol., 11, 26, 1994. 24. Couch, J.M. et al., Diagnosing and treating Phthirus pubis palpebrarum. Surv. Ophthalmol., 26, 219, 1982. 24. Silburt, B.S., and Parsons, W.L., Scalp infestation by Phthirus pubis in a 6-week-old infant. Pediatr. Dermatol., 7, 205, 1990. 25. Singh, S. et al, Phthirus pubis infestation of the scalp: Report of three cases. Rev. Infect. Dis., 12, 560, 1990. 26. Mansour, A.M., Photo essay: Phthiriasis palpebrarum. Arch. Ophthalmol., 118, 1458, 2000. 27. Bessis, D. et al., Erythema annulare centrifugum induced by generalized Phthirus pubis infestation. Br. J. Dermatol., 149, 1291, 2003. 28. Ikeda, N. et al., Phthirus pubis infestation of the eyelashes and scalp hairs in a girl. Pediatr. Dermatol., 20, 356, 2003. 29. DeFazio, J.L., and Spencer, P., Images in clinical medicine. Dermoscopy of phthiriasis. N. Engl. J. Med., 362, e33, 2010. 30. Turgut, B. et al., Phthriasis palpebrarum mimicking lid eczema and blepharitis. J. Ophthalmol., 2009, 803951, 2009. 31. Ma, D.L., and Vano-Galvan, S., Infestation of the eyelashes with Phthirus pubis. CMAJ, 182, E187, 2010. 32. Keklikci, U. et al., Phthiriasis palpebrarum in an infant. J. Pediatr. Ophthalmol. Strabismus, 46, 173, 2009. 33. Newsome, J.H., Fiore, J.L., Jr., and Hackett, E., Treatment of infestation with Phthirus pubis: Comparative ef�cacies of synergized pyrethrins and gamma-benzene hexachloride. Sex. Transm. Dis., 6, 203, 1979. 35. Ashkenazi, I., Desatnik, H.R., and Abraham, F.A., Yellow mercuric oxide: A treatment of choice for Phthriasis palpebrarum. Brit. J. Ophthalmol., 75, 356, 1991. 36. Burkhart, C.N., and Burkhart, C.G., Oral ivermectin therapy for phthiriasis palpebrum. Arch. Ophthalmol., 118, 134, 2000. 37. Kumar, N., Dong, B., and Jenkins, C., Pubic lice effectively treated with Pilogel. Eye, 17, 538, 2003. 38. Yoon, K.C. et al. Mechanical treatment of phthiriasis palpebrarum. Korean J. Ophthalmol., 17, 71, 2003. 39. Elston, D.M., Drugs used in the treatment of pediculosis. J. Drugs Dermatol., 4, 207, 2005. 40. Leone, P.A., Scabies and pediculosis pubis: An update of treatment regimens and general review. Clin. Infect. Dis., 44, S153, 2007. 41. Scott, G.R., and Chosidow, O., European guideline for the management of pediculosis pubis, 2010. Int. J. STD AIDS, 22, 304, 2011. 42. Das, J.C., Sharma, P., and Singla, R., A new treatment modality for phthiriasis palpebrarum. J. Pediatr. Ophthalmol. Strabismus, 40, 304, 2003. 43. Burgess, I.F., Human lice and their management. Adv. Parasitol., 36, 271, 1995. 44. Galiczynski, E.M., Jr., and Elston, D.M., What’s eating you? Pubic lice (Pthirus pubis). Cutis, 81, 109, 2008.
45. Turow, V.D., Phthiriasis palpebrarum: An unusual cause of blepharitis. Arch. Pediatr. Adolesc. Med., 149, 704, 1995.
46. Lin, Y.-C. et al., Phthiriasis palpebrarum: An unusual blepharoconjunctivitis. Chin. Med. J., 65, 498, 2003.
47. Vandeweghe, K., and Zeyen, P., Phthiriasis palpebrarum: 2 case reports. Bull. Soc. Belge Ophtalmol., 300, 27, 2006.
48. Zvandasara, T., and Lam, F.C., Conjunctivitis due to Phthirus pubis: A trap for the unwary. Int. J. STD AIDS, 21, 519, 2010.
49. Rundle, P.A., and Hughes, D.S., Phthirus pubis infestation of the eyelids. Brit. J. Ophthalmol., 77, 815, 1993. 50. Desalle, R. et al. DNA sequences from a fossil termite in Oligo-Miocene amber and their phylogenetic implications. Science, 257, 1933, 1992. 51. Raoult, D. et al., Molecular identi�cation of lice from preColumbian mummies. J. Infect. Dis., 197, 535, 2008. 52. Leo, N.P., and Barker, S.C., Unravelling the evolution of the head lice and body lice of humans. Parasitol. Res., 98, 44, 2005. 53. Reed, D.L. et al. Genetic analysis of lice supports direct contact between modern and archaic humans. PloS Biol., 2, e340, 2004. 81 Chapter 81 - Pulex, Xenopsylla, and Ctenocephalides (Human, Rat, and Dog/Cat Fleas)
1. Buckland, P.C., and Sadler, J.P., A biogeography of the human �ea, Pulex irritans L. (Siphonaptera: Pulicidae). J. Biogeography, 16, 115, 1989. 2. Whiting, M.F. et al., A molecular phylogeny of �eas (Insecta: Siphonaptera): Origins and host associations. Cladistics 24, 1, 2008. 3. Elston, D.M., What’s eating you? Pulex irritans. Cutis, 66, 175, 2000. 4. Brouqui, P., and Raoult, D., Arthropod-borne diseases in homeless. Ann. N. Y. Acad, Sci., 1078, 223, 2006. 5. Gracia, M.J. et al., Pulex irritans infestation in dogs. Vet. Rec., 147, 748, 2000. 6. Ménier, K., Infestation of dairy goats with Pulex irritans. Vet. Rec., 153, 128, 2003. 7. Christodoulopoulos, G. et al., Biological, seasonal and environmental factors associated with Pulex irritans infestation of dairy goats in Greece. Vet. Parasitol., 137, 137, 2006. 8. Yin, J.X. et al., Predictors for abundance of host �ea and �oor �ea in households of villages with endemic commensal rodent plague, Yunnan Province, China. PLoS Negl. Trop. Dis., 5, e997, 2011. 9. Engelthaler, D.M., and Gage, K.L., Quantities of Yersinia pestis in �eas (Siphonaptera: Pulicidae, Ceratophyllidae, and Hystrichopsyllidae) collected from areas of known or suspected plague activity. J. Med. Entomol., 37, 422, 2000. 10. Loftis, A.D. et al., Surveillance of Egyptian �eas for agents of public health signi�cance: Anaplasma, Bartonella, Coxiella, Ehrlichia, Rickettsia, and Yersinia pestis. Am. J. Trop. Med. Hyg., 75, 41, 2006. 11. Laudisoit, A. et al., Plague and the human �ea, Tanzania. Emerg. Infect. Dis., 13, 687, 2007. 12. Hornok, S. et al., Molecular investigation of hard ticks (Acari: Ixodidae) and �eas (Siphonaptera: Pulicidae) as potential vectors of rickettsial and mycoplasmal agents. Vet. Microbiol., 140, 98, 2010. 13. Krug, L.E., and Elston, D.M., What’s eating you? Oriental rat �ea (Xenopsylla cheopis). Cutis, 86, 282, 2010. 14. Eremeeva, M.E. et al., Rickettsia typhi and R. felis in rat �eas (Xenopsylla cheopis), Oahu, Hawaii. Emerg. Infect. Dis., 14, 1613, 2008. 15. Billeter, S.A. et al., Molecular detection and identi�cation of Bartonella species in Xenopsylla cheopis (Siphonaptera: Pulicidae) collected from Rattus norvegicus in Los Angeles, California. Appl. Env. Microbiol., 77, 7850, 2011. 16. Morick, D. et al., Investigation of Bartonella acquisition and transmission in Xenopsylla ramesis �eas (Siphonaptera: Pulicidae). Mol. Ecol., 20, 2864, 2011. 17. Schotthoefer, A.M. et al., Effects of temperature on earlyphase transmission of Yersina pestis by the �ea, Xenopsylla cheopis. J. Med. Entomol., 48, 411, 2011. 18. Dryden, M.W., and Rust, M.K., The cat �ea: Biology, ecology and control. Vet. Parasitol., 52, 1, 1994. 19. Cañón-Franco, W.A., and Pérez-Bedoya, J.L., Siphonaptera (Pulicidae) in dogs and cats of Colombia: Clinical and epidemiological aspects. Vet. Parasitol., 173, 353, 2010. 20. Elston, D.M., and Do, H., What’s eating you? Cat �ea (Ctenocephalides felis), Part 1: Clinical features and role as a disease vector. Cutis, 85, 231, 2010. 21. Rust, M.K., and Dryden, M.W., The biology, ecology, and management of the cat �ea. Annu. Rev. Entomol., 42, 451, 1997. 22. Kelly, P.J., A review of bacterial pathogens in Ctenocephalides felis in New Zealand. N. Z. Vet. J., 52, 352, 2004. 23. Shaw, S.E. et al., Pathogen carriage by the cat �ea Ctenocephalides felis (Bouché) in the United Kingdom. Vet. Microbiol., 102, 183, 2004.
TABLE 81.1
Primers Used for Phylogenetic Analysis of Flea Species
Primer Sequence (5′–3′)
Sen-ITS1 GTA CAC ACC GCC CGT GCG TAC T
Rev-ITS1 GCT GCG TTC TTC ATC GAC CC
Sen-ITS2 GGG TCG ATG AAG AAC GCA GC
Rev-ITS2 GCG CAC ATG CTA GAC TCC GTG GTT CAA G
Sen-mt16S TAC ATA ACA CGA GAA GAC C
Rev-mt16S GTG ATT GCG CTG TTA TCC
24. Hawley, J.R., Shaw, S.E., and Lappin, M.R., Prevalence of Rickettsia felis DNA in the blood of cats and their �eas in the United States. J. Feline Med. Surg., 9, 258, 2007.
25. Reeves, W.K. et al., Association of Bartonella with the �eas (Siphonaptera) of rodents and bats using molecular techniques. J. Vector Ecol., 32, 118, 2007.
26. Eisen, R.J. et al., Early-phase transmission of Yersinia pestis by cat �eas (Ctenocephalides felis) and their potential role as vectors in a plague-endemic region of Uganda. Am. J. Trop. Med. Hyg., 78, 949, 2008.
27. Eisen, R.J., Eisen, L., and Gage, K.L., Studies of vector competency and ef�ciency of North American �eas for Yersinia pestis: State of the �eld and future research needs. J. Med. Entomol., 46, 737, 2009.
28. Oliveira, K.A. et al., Molecular identi�cation of Rickettsia felis in ticks and �eas from an endemic area for Brazilian Spotted Fever. Mem. Inst. Oswaldo Cruz, 103, 191, 2008.
29. Reif, K.E., and Macaluso, K.R., Ecology of Rickettsia felis: A review. J. Med. Entomol., 46, 723, 2009.
30. Kaewmongkol, G. et al., Zoonotic Bartonella species in �eas and blood from red foxes in Australia. Vector Borne Zoonotic Dis., 11, 1549, 2011.
31. Kernif, T. et al., Bartonella clarridgeiae in �eas, Tahiti, French Polynesia. Emerg. Infect. Dis., 17, 1773, 2011.
32. Parola, P., Rickettsia felis: From a rare disease in the USA to a common cause of fever in sub-Saharan Africa. Clin. Microbiol. Infect., 17, 996, 2011. 33. Tsai, K.H. et al., Prevalence of Rickettsia felis and the �rst identi�cation of Bartonella henselae Fizz/CAL-1 in cat �eas (Siphonaptera: Pulicidae) from Taiwan. J. Med. Entomol., 48, 445, 2011. 34. Williams, M. et al., First probable Australian cases of human infection with Rickettsia felis (cat-�ea typhus). Med. J. Aust., 194, 41, 2011. 35. Ugbomoiko, U.S., Ariza, L., and Heukelbach, J., Parasites of importance for human health in Nigerian dogs: High prevalence and limited knowledge of pet owners. BMC Vet. Res., 4, 49, 2008. 36. Rust, M.K., Advances in the control of Ctenocephalides felis (cat �ea) on cats and dogs. Trends Parasitol., 21, 232, 2005. 37. Dittmar, K. et al., Techniques of DNA-studies on prehispanic ectoparasites (Pulex sp., Pulicidae, Siphonaptera) from animal mummies of the Chiribaya culture, southern Peru. Mem. Inst. Oswaldo Cruz, 98, S53, 2003. 38. Vobis, M. et al., Molecular phylogeny of isolates of Ctenocephalides felis and related species based on analysis of ITS1, ITS2 and mitochondrial 16S rDNA sequences and random binding primers. Parasitol. Res., 94, 219, 2006. 39. Gamerschlag, S. et al., Repetitive sequences in the ITS1 region of the ribosomal DNA of Tunga penetrans and other �ea species (Insecta, Siphonaptera). Parasitol. Res., 102, 193, 2008. 82 Chapter 82 - Tunga (Jigger Flea)
9. Collins, G. et al., Tungiasis: A neglected health problem in rural Cameroon. Int. J. Collaborat. Res. Intern. Med. Public Health, 1, 2, 2009.
10. Maco, V., Tantaleán, M., and Gotuzzo, E., Evidence of tungiasis in pre-Hispanic America. Emerg. Infect. Dis., 17, 5, 2011.
11. Fioravanti, M.L., Pampiglione, S., and Trentini, M., A second species of Tunga (Insecta, Siphonaptera) infecting man: Tunga trimamillata. Parasite, 10, 282, 2003.
12. Fioravanti, M.I. et al., Presence of Tunga trimamillata (Insecta, Siphonaptera) in Peru. Parasite, 13, 85, 2006.
13. Fioravanti, M.L. et al., Sand �ea (Tunga spp.) infections in humans and domestic animals: State of the art. Med. Vet. Entomol., 23, 172, 2009.
14. Maco, V., Maco, V.P., and Gotuzzo, E., An ectopic case of Tunga spp. infection in Peru. Am. J. Trop. Med. Hyg., 82, 1076, 2010.
15. Heukelbach, J. et al., Tungiasis: A neglected health problem of poor communities. Trop. Med. Int. Health, 6, 267, 2001.
16. Heukelbach, J. et al., The animal reservoir of Tunga penetrans in severely affected communities of north-east Brazil. Med. Vet. Entomol., 18, 329, 2004.
17. Heukelbach, J., Franck, S., and Feldmeier, H., High attack rate of Tunga penetrans (Linnaeus 1758) infestation in an impoverished Brazilian community. Trans. R. Soc. Trop. Med. Hyg., 98, 431, 2004.
18. Wilcke, T. et al., High prevalence of tungiasis in a poor neighbourhood in Fortaleza, Northeast Brazil. Acta Trop., 83, 255, 2002.
19. de Carvalho, R.W. et al., The patterns of tungiasis in Araruama township, state of Rio de Janeiro, Brazil. Mem. Inst. Oswaldo Cruz, 98, 31, 2003.
20. Feldmeier, H. et al., Investigations on the biology, epidemiology, pathology and control of Tunga penetrans in Brazil: III. Cytokine levels in peripheral blood of infected humans. Parasitol. Res., 91, 298, 2003.
21. Feldmeier, H. et al., Investigations on the biology, epidemiology, pathology and control of Tunga penetrans in Brazil: IV. Clinical and histopathology. Parasitol. Res., 94, 275, 2004.
22. Heemskerk, J., van Empel, I., and Jakimowicz, J.J., Tunga penetrans. A case report and review of the literature. Acta Chir. Belg., 105, 548, 2005.
23. Ugbomoiko, U.S., Ofoezie, I.E., and Heukelbach, J., Tungiasis: High prevalence, parasite load, and morbidity in a rural community in Lagos State, Nigeria. Int. J. Dermatol., 46, 475, 2007.
24. Valentín, S.M. et al., Tungiasis: A case report. P.R. Health Sci. J., 26, 423, 2007.
25. Damazio, O.R., and Silva, M.V., Tungiasis in schoolchildren in Criciúma, Santa Catarina State, South Brazil. Rev. Inst. Med. Trop. Sao Paulo, 51, 103, 2009.
26. Mazigo, H.D. et al., Tungiasis infestation in Tanzania. J. Infect. Dev. Ctries., 4, 187, 2010.
27. Fein, H. et al., Tungiasis in North America: A report of 2 cases in internationally adopted children. J. Pediatr., 139, 744, 2001.
28. Pospísilová, A., and Pirochtová, K., Tungiasis (tungosis) comes to the Czech Republic. J. Cosmet. Dermatol., 1, 216, 2002.
29. Franck, S., Feldmeier, H., and Heukelbach, J., Tungiasis: More than an exotic nuisance. Travel Med. Infect. Dis., 1, 159, 2003.
30. Caputo, V. et al., Tungiasis: A report of four cases and a review of imported cases in Italy in the last 30 years. G. Ital. Med. Trop., 10, 33, 2005. 31. Sachse, M.M., Guldbakke, K.K., and Khachemoune, A., Tunga penetrans: A stowaway from around the world. J. Eur. Acad. Dermatol. Venereol., 21, 11, 2007. 32. Veraldi, S., and Valsecchi, M., Imported tungiasis: A report of 19 cases and review of the literature. Int. J. Dermatol., 46, 1061, 2007. 33. Escamilla-Martinez, E. et al., Tungiasis–traveler’s ectoparasitosis of the foot: A case report. Foot Ankle Int., 29, 354, 2008. 34. Hager, J. et al., Tungiasis in the United States: A travel souvenir. Dermatol. Online J., 14, 3, 2008. 35. Dialynas, M. et al., Imported human tungiasis in Greece. Travel Med. Infect. Dis., 7, 375, 2009. 36. Ferreira, L.A. et al., Tunga penetrans as a traveler’s disease. Travel Med. Infect. Dis., 7, 381, 2009. 37. Scalvenzi, M. et al. Tungiasis: Case report of a traveller to Kenya. Case Rep. Dermatol., 1, 29, 2009. 38. Hakeem, M.J. et al., Tungiasis—a cause of painful feet in a tropical traveller. Travel Med. Infect. Dis., 8, 29, 2010. 39. Rosmaninho, A. et al., Tunga penetrans: Painful lesions on the feet-the �rst imported case from Guinea-bissau. Case Report Med., 2010, 681302, 2010. 40. Wiwanitkit, V., Tungiasis among traveller. Travel Med. Infect. Dis., 8, 273, 2010. 41. Yotsu, R.R. et al., Imported tungiasis in a Japanese student returning from East Africa. J. Dermatol., 38, 185, 2011. 42. Feldmeier, H. et al., Investigations on the biology, epidemiology, pathology and control of Tunga penetrans in Brazil: VI. Natural history of the infestation in laboratory-raised Wistar rats. Parasitol. Res., 102, 1, 2007. 43. Witt, L. et al., Infestation of Wistar rats with Tunga penetrans in different microenvironments. Am. J. Trop. Med. Hyg., 76, 666, 2007. 44. Pilger, D. et al., Investigations on the biology, epidemiology, pathology, and control of Tunga penetrans in Brazil: VII. The importance of animal reservoirs for human infestation. Parasitol. Res., 102, 875, 2008. 45. Ugbomoiko, U.S., Ariza, L., and Heukelbach, J., Pigs are the most important animal reservoir for Tunga penetrans (jigger �ea) in rural Nigeria. Trop. Doct., 38, 226, 2008. 46. Luchetti, A. et al., Wolbachia infection in the newly described Ecuadorian sand �ea, Tunga trimamillata. Exp. Parasitol., 108, 18, 2004. 47. Heukelbach, J. et al., Ectopic localization of tungiasis. Am. J. Trop. Med. Hyg., 67, 214, 2002. 48. Feldmeier, H. et al., High exposure to Tunga penetrans (Linnaeus, 1758) correlates with intensity of infestation. Mem. Inst. Oswaldo Cruz, 101, 65, 2006. 49. Winter, B. et al., Tungiasis-related knowledge and treatment practices in two endemic communities in northeast Brazil. J. Infect. Dev. Ctries., 3, 458, 2009. 50. Kaimbo, D.K. et al., Upper eyelid localisation of Tunga penetrans. Ophthalmologica, 221, 439, 2007. 51. Macías, P.C., and Sashida, P.M., Cutaneous infestation by Tunga penetrans. Int. J. Dermatol., 39, 296, 2000. 52. Kehr, J.D. et al., Morbidity assessment in sand �ea disease (tungiasis). Parasitol. Res., 100, 413, 2007. 53. Cestari, T.F., Pessato, S., and Ramos-e-Silva, M., Tungiasis and myiasis. Clin. Dermatol. 25, 158, 2007. 54. Gibbs, S.S., The diagnosis and treatment of tungiasis. Br. J. Dermatol., 159, 981, 2008. 55. Ade-Serrano, M.A., Olomolehin, O.G., and Adenwunmi, A., Treatment of human tungiasis with niridazole (Ambilhar) in a double-blind placebo-controlled trial. Ann. Trop. Med. Parasitol., 76, 89, 1982.
56. Heukelbach, J. et al., Topical treatment of tungiasis: A randomized, controlled trial. Ann. Trop. Med. Parasitol., 97, 743, 2003.
57. Klimpel, S. et al., Field trial of the ef�cacy of a combination of imidacloprid and permethrin against Tunga penetrans (sand �ea, jigger �ea) in dogs in Brazil. Parasitol. Res., 97, S113, 2005.
58. Heukelbach, J., Revision on tungiasis: Treatment options and prevention. Expert Rev. Anti Infect. Ther., 4, 151, 2006.
59. Ariza, L. et al., A simple method for rapid community assessment of tungiasis. Trop. Med. Int. Health, 15, 856, 2010.
60. Di Stefani, A. et al., An additional dermoscopic feature of tungiasis. Arch. Dermatol., 141, 1045, 2005.
61. Marazza, G. et al., Tunga penetrans: Description of a new dermoscopic sign—the radial crown. Arch. Dermatol., 145, 348, 2009.
62. Bauer, J. et al., Dermoscopy of tungiasis. Arch. Dermatol., 40, 761, 2004.
63. Bakos, R.M., and Bakos, L., ‘Whitish chains’: A remarkable in vivo dermoscopic �nding of tungiasis. Br. J. Dermatol., 159, 991, 2008.
64. Cabrera, R., and Daza, F., Dermoscopy in the diagnosis of tungiasis. Br. J. Dermatol., 160, 1136, 2009.
65. Dunn, R., Asher, R., and Bowling, J., Tunga penetrans—egg head? Dermatol. Online J., 16, 14, 2010.
66. Cabrera, R., and Daza, F., Eggs seen with dermoscopy. Br. J. Dermatol., 158, 635, 2008.
67. Smith, M.D., and Procop, G.W., Typical histologic features of Tunga penetrans in skin biopsies. Arch. Pathol. Lab. Med., 126, 714, 2002. 68. Vobis, M. et al., Molecular biological investigations of Brazilian Tunga sp. isolates from man, dogs, cats, pigs and rats. Parasitol. Res., 96, 107, 2005. 69. Vobis, M. et al., Molecular phylogeny of isolates of Ctenocephalides felis and related species based on analysis of ITS1, ITS2 and mitochondrial 16S rDNA sequences and random binding primers. Parasitol. Res., 94, 219, 2004. 70. Luchetti A. et al., Molecular characterization of Tunga trimamillata and T. penetrans (Insecta, Siphonaptera, Tungidae): Taxonomy and genetic variability. Parasite, 12, 123, 2005. 71. Luchetti, A. et al., Genetic variability of Tunga penetrans (Siphonaptera, Tungidae) sand �eas across South America and Africa. Parasitol. Res., 100, 593, 2007. 72. Gamerschlag, S. et al., Repetitive sequences in the ITS1 region of the ribosomal DNA of Tunga penetrans and other �ea species (Insecta, Siphonaptera). Parasitol. Res., 102, 193, 2008. 73. Avelar, D.M., and Linardi, P.M., Use of multiple displacement ampli�cation as pre-polymerase chain reaction (pre-PCR) to amplify genomic DNA of siphonapterids preserved for long periods in scienti�c collections. Parasites Vectors, 3, 86, 2010. 74. Heukelbach, J., Tungiasis. Rev. Inst. Med. Trop. Sao Paulo, 47, 307, 2005. 75. Feldmeier, H. et al., Bacterial superinfection in human tungiasis. Trop. Med. Int. Health, 7, 559, 2002. 76. Heukelbach, J. et al., Seasonal variation of tungiasis in an endemic community. Am. J. Trop. Med. Hyg., 72, 145, 2005. 77. Muehlstaedt, M., Images in clinical medicine. Periungual tungiasis. N. Engl. J. Med., 359, e30, 2008.
Section VIII
Arthropoda: Pentastomida 83 Chapter 83 - Armillifer, Linguatula, and Porocephalus (Tongue Worms)
23. Khalil, G.M., Linguatula serrata (Pentastomida) parasitizing humans and animals in Egypt, neighbouring countries, and elsewhere: A review. J. Egypt. Public Health Assoc., 47, 364, 1972.
24. Gardiner, S.N. et al., Pentastomiasis in Canada. Arch. Pathol. Lab. Med., 115, 515, 1991.
25. Yagi, H. et al., The Marrara syndrome: A hypersensitivity reaction of the upper respiratory tract and buccopharyngeal mucosa to nymphs of Linguatula serrata. Acta Trop., 62, 127, 1996.
26. De Meneghi, D., Pentastomes (Pentastomiasis, Amillifer armillatus Wyman, 1848) in snakes from Zambia. Parasitogia, 41, 573, 1999.
27. Lazo, R.F. et al., Ocular linguatuliasis in Ecuador: Case report and morphometric study of the larva of Linguatula serrata. Am. J. Trop. Med. Hyg., 60, 405, 1999.
28. Siavashi, M.R., Assmar, M., and Vatankhah, A., Nasopharyngeal pentastomiasis (Halzoun): Report of 3 cases. Iran J. Mol. Sci., 27, 191, 2002.
29. Dakubo, J.C. et al., Human pentastomiasis: A case report. West Afr. J. Med., 25, 166, 2006.
30. Dakubo, J., Naaeder, S., and Kumodji, R., Totemism and the transmission of human pentastomiasis. Ghana Med. J., 42, 165, 2008.
31. Tappe, D. et al., Linguatuliasis in Germany. Emerg. Infect. Dis., 12, 1034, 2006.
32. Drabick, J.J., Pentastomiasis. Rev. Infect. Dis., 9, 1087, 1987.
33. Deweese, M.W., Murrah, W.F., and Caruthers, S.B., Case report of a tongue worm (Linguatula serrata) in the anterior chamber. Arch. Ophthalmol., 68, 587, 1962.
34. Sousaefaro, B., and Pinhao, R.C., An isolated case of ocular parasitosis caused by Linguatula serrata. J. Soc. Cienc. Med. Lisb., 128, 401, 1964.
35. Smith, J.A., Oladiran, B., and Lagundoye, S.B., Pentastomiasis and malignancy. Ann. Trop. Med. Parasitol., 69, 503, 1975.
36. Buslau, M., Kuhne, U., and Marsch, W.C., Dermatological signs of nasopharyngeal linguatulosis (Halzoun, Marrara syndrome): The possible role of a major basic protein. Dermatologica, 181, 327, 1990. 37. Pampiglione, S. et al., A nodular pulmonary lesion due to Linguatula serrata in an HIV-positive man. Parassitologia, 43, 105, 2001. 38. Gardiner, C.H., Dyke, J.W., and Shirley, S.F., Hepatic granuloma due to a nymph of Linguatula serrata in a woman from Michigan: A case report and review of the literature. Am. J. Trop. Med. Hyg., 33, 187, 1984. 39. Herzog, U., Marty, P., and Zak, F., Pentastomiasis: Case report of an acute abdominal emergency. Acta Trop., 42, 261, 1985. 40. Baird, J.K., Kassebaum, L.J., and Ludwig, G.K., Hepatic granuloma in a man from North America caused by a nymph of Linguatula serrata. Pathology, 20, 198, 1988. 41. du Plessis, V. et al. Pentastomiasis (Armillifer armillatus infestation). S. Afr. Med. J., 97, 928, 2007. 42. Tappe, D., and Büttner, D.W., Diagnosis of human visceral pentastomiasis. PLoS Negl. Trop. Dis., 3, E320, 2009. 43. Yapo Ette, H. et al., Human pentastomiasis discovered postmortem. Forensic Sci. Int., 137, 52, 2003. 44. Ma, K.C., Qiu, M.H., and Rong, Y.L., Pathological differentiation of suspected cases of pentastomiasis in China. Trop. Med. Int. Health, 7, 166, 2002. 45. Lang, Y. et al., Intraocular pentastomiasis causing unilateral glaucoma. Br. J. Ophthalmol., 71, 391, 1987. 46. Morsy, T.A., El-Sharkawy, I.M., and Lashin, A.H., Human nasopharyngeal linguatuliasis (Pentasomida) caused by Linguatula serrata. J. Egypt. Soc. Parasitol., 29, 787, 1999. 47. Yao, M.H., Wu, F., and Tang, L.F., Human pentastomiasis in China: Case report and literature review. J. Parasitol., 94, 1295, 2008. 48. Magnino, S. et al., Biological risks associated with consumption of reptile products. Int. J. Food Microbiol., 134, 163, 2009. 49. Machado, M.A.C. et al., Unusual case of pentastomiasis mimicking liver tumour. J. Gastroenterol. Hepatol., 21, 1218, 2006. 50. Lai, C. et al., Imaging features of pediatric pentastomiasis infection: A case report. Korean J. Radiol., 11, 480, 2010. 51. Mourra, N., Huerre, M., and Tiret, E., Liver pentastomiasis associated with rectal adenocarcinoma. Colorectal Dis., 12, 261, 2010.