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Gastrointestinal Helminthic Parasites of Habituated Wild Chimpanzees

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Aus dem Institut für Parasitologie und Tropenveterinärmedizin des Fachbereichs Veterinärmedizin der Freien Universität Berlin Gastrointestinal helminthic parasites of habituated wild chimpanzees (Pan troglodytes verus) in the Taï NP, Côte d’Ivoire − including characterization of cultured helminth developmental stages using genetic markers Inaugural-Dissertation zur Erlangung des Grades eines Doktors der Veterinärmedizin an der Freien Universität Berlin vorgelegt von Sonja Metzger Tierärztin aus München Berlin 2014 Journal-Nr.: 3727 Gedruckt mit Genehmigung des Fachbereichs Veterinärmedizin der Freien Universität Berlin Dekan: Univ.-Prof. Dr. Jürgen Zentek Erster Gutachter: Univ.-Prof. Dr. Georg von Samson-Himmelstjerna Zweiter Gutachter: Univ.-Prof. Dr. Heribert Hofer Dritter Gutachter: Univ.-Prof. Dr. Achim Gruber Deskriptoren (nach CAB-Thesaurus): chimpanzees, helminths, host parasite relationships, fecal examination, characterization, developmental stages, ribosomal RNA, mitochondrial DNA Tag der Promotion: 10.06.2015 Contents I INTRODUCTION ---------------------------------------------------- 1- 4 I.1 Background 1- 3 I.2 Study objectives 4 II LITERATURE OVERVIEW --------------------------------------- 5- 37 II.1 Taï National Park 5- 7 II.1.1 Location and climate 5- 6 II.1.2 Vegetation and fauna 6 II.1.3 Human pressure and impact on the park 7 II.2 Chimpanzees 7- 12 II.2.1 Status 7 II.2.2 Group sizes and composition 7- 9 II.2.3 Territories and ranging behavior 9 II.2.4 Diet and hunting behavior 9- 10 II.2.5 Contact with humans 10 II.2.6 Pathogens and diseases affecting Taï chimpanzees 11- 12 II.3 Helminth classification and taxonomy 12- 14 II.4 Gastrointestinal helminthic parasites reported in wild 14- 31 chimpanzees II.4.1 Overview of reported helminth diversity 14- 17 II.4.2 Helminth life cycles, morphology and pathogenicity 18- 31 II.4.2.1 NEMATODES 18- 28 II.4.2.1.1 Superfamily Strongyloidea (`strongyle´ nematodes) 18- 22 II.4.2.1.2 Genus Strongyloides 22- 23 II.4.2.1.3 Genus Capillaria 23- 24 II.4.2.1.4 Genus Trichuris 24- 25 II.4.2.1.5 Genus Enterobius 25- 26 II.4.2.1.6 Genus Ascaris 26- 27 II.4.2.1.7 Genus Probstmayria 27 II.4.2.1.8 Genus Physaloptera (Genus Abbreviata) 27- 28 II.4.2.2 TREMATODES 28- 29 II.4.2.2.1 Family Dicrocoeliidae 28- 29 Contents II.4.2.2.2 Other Trematodes 29 II.4.2.3 CESTODES 29- 31 II.4.2.3.1 Genus Bertiella 29- 30 II.4.2.3.2 Genus Hymenolepis 30- 31 II.4.2.3.3 Other Cestodes 31 II.5 Impacts of host-intrinsic and seasonal factors on 31- 35 patterns of parasitism II.5.1 Influence of host sex 31- 33 II.5.2 Influence of host age 33- 34 II.5.3 Influence of seasonality and seasonal climatic variations 35 II.6 Molecular markers used for identification and 35- 37 differentiation of helminthic parasites II.6.1 Nuclear ribosomal DNA markers 36 II.6.2 Mitochondrial DNA markers 36- 37 III METHODS & MATERIAL ---------------------------------------- 38- 59 III.1 Study site 38 III.2 Collection and processing of chimpanzee fecal samples 38- 40 III.2.1 Period and frequency of sample collection 38- 39 III.2.2 Method of sample collection 39 III.2.3 Sample preservation and storage 40 III.3 Collection and processing of fecal samples from 41 sympatric monkeys III.3.1 Sample collection 41 III.3.2 Sample processing 41 III.4 Recording of weather data 42 III.5 Sample screening using conventional coprological 42- 47 methods III.5.1 Modified Wisconsin Sugar Flotation (MWSF) 42- 44 III.5.1.1 Method description 42- 43 III.5.1.2 Number and distribution of examined samples 43- 44 III.5.2 Harada-Mori Fecal Cultures 44- 45 III.5.2.1 Method description 44- 45 III.5.2.2 Number and distribution of examined samples 45 III.5.3 Sodium-Nitrate Flotation and Sedimentation 45- 47 Contents III.5.3.1 Method description 45- 47 III.5.3.2 Number and distribution of examined samples 47 III.6 Statistical data analysis 47- 50 III.6.1 Descriptive statistics 47 III.6.2 Inferential statistics 47- 50 III.6.2.1 Model 1 (response variable: individual cumulative helminth 48- 49 morphotype richness) III.6.2.2 Model 2 (response variable: ppg) 49- 50 III.7 Preservation of larvae & free-living adult helminths for 51 DNA extraction III.8 DNA extraction 51 III.9 Polymerase chain reaction (PCR) 52- 59 III.9.1 Strongyle nematodes 52- 55 III.9.1.1 Ribosomal ITS-2 52- 53 III.9.1.2 12s RNA mtDNA 53- 55 III.9.2 Genus Strongyloides 55- 56 III.10 Visualization of amplification products 56 III.11 Purification of PCR products 56 III.12 Cloning 56- 57 III.13 Sequencing 57 III.13.1 Sequencing of cloned PCR products 57 III.13.2 Direct sequencing 57 III.14 Sequence- and phylogenetic analysis 57- 59 III.14.1 Sequence analysis 57- 58 III.14.2 Phylogenetic analysis 58- 59 IV RESULTS ------------------------------------------------------------- 60- 114 IV.1 Spectrum of gastrointestinal helminths determined using 60- 67 conventional coprology IV.1.1 Macroscopic examination 60 IV.1.2 Coprological microscopic examination 60- 61 IV.1.2.1 Modified Wisconsin Sugar Flotation (MWSF) 60 IV.1.2.2 Sodium Nitrate Flotation and Sedimentation 60 IV.1.2.3 Harada-Mori Fecal Cultures 61 Contents IV.1.3 Overall spectrum of detected parasitic helminth genera and 61- 67 morphotypes IV.2 Morphometrics and morphological description of 67- 72 helminth developmental stages IV.2.1 NEMATODES 67- 72 IV.2.1.1 Strongyle nematodes (Superfamily Strongyloidea) 70- 73 IV.2.1.1.1 Genus Ternidens 69 IV.2.1.1.2 Genus Necator 69 IV.2.1.1.3 Genus Oesophagostomum 69- 70 IV.2.1.1.4 Family Trichostronglidae 70 IV.2.1.2 Genus Strongyloides (Superfamily Strongyloidoidea) 70- 71 IV.2.1.3 Genus Trichuris (Family Trichuridae) 71 IV.2.1.4 Genus Capillaria (Family Capillariidae) 72 IV.2.1.5 Genus Enterobius (Family Oxyuridae) 72 IV.2.2 CESTODES 72 IV.2.3 TREMATODES 72 IV.3 Parameters of helminth infection 73- 90 IV.3.1 Overall frequency of helminth infection 73 IV.3.2 Helminth prevalence 73- 83 IV.3.2.1 NEMATODES 73- 76 IV.3.2.2 CESTODES 76 IV.3.2.3 TREMATODES 76 IV.3.2.4 Influence of sampling intensity on nematode and cestode 76- 77 prevalence IV.3.2.5 Variations in helminth prevalence between male and female 77- 80 chimpanzees IV.3.2.6 Method comparison 80- 82 IV.3.2.6.1 Comparison of nematode detection rates between MWSF and 80- 81 Harada-Mori fecal cultures IV.3.2.6.2 Comparison of helminth detection rates between fresh and 81- 82 formalin- preserved samples IV.3.3 Helminth morphotype richness and prevalence of multiple 83- 85 infections IV.3.3.1 Overall sample helminth morphotype richness 83 IV.3.3.2 Cumulative individual helminth morphotype richness 83- 84 Contents IV.3.3.3 Influence of host-intrinsic factors on cumulative individual 84- 85 helminth morphotype richness IV.3.3.4 Prevalence of multiple infections 85 IV.3.4 Propagule output (ppg-values) 85- 86 IV.3.5 Influence of seasonality and host intrinsic factors on fecal 87- 90 propagule output IV.4 Identification and characterization of cultured strongyle 91- 114 L3-larvae and Strongyloides specimens using genetic markers & comparison with sequences of helminth developmental stages from sympatric monkeys IV.4.1 Identification of strongyle L3-larvae by ITS-2 and 12s RNA 92- 110 mtDNA sequencing IV.4.1.1 Ternidens L3-larvae (Subfam. Chabertiinae) 92- 95 IV.4.1.1.1 Overview of examined larvae and success rates of DNA 92 extraction and PCR amplification IV.4.1.1.2 Analysis of ITS-2 sequences 92- 93 IV.4.1.1.3 Analysis of 12s RNA mtDNA sequences 93 IV.4.1.1.4 Comparison with Ternidens sequences from sympatric 93- 94 monkeys IV.4.1.1.5 Comparison with Ternidens sequences from other host 94- 95 species IV.4.1.1.6 Phylogenetic analysis 95 IV.4.1.2 Oesophagostomum L3-larvae (Subfam. Chabertiinae) 95- 101 IV.4.1.2.1 Overview of examined larvae and success rates of DNA 95 extraction and PCR amplification IV.4.1.2.2 Analysis of ITS-2 sequences 96 IV.4.1.2.3 Analysis of 12s RNA mtDNA sequences 97 IV.4.1.2.4 Comparison with Oesophagostomum sequences from 97- 99 sympatric monkeys IV.4.1.2.5 Phylogenetic analysis of Oesophagostomum and Ternidens 99- 101 sequences IV.4.1.3 Necator L3-larvae (Fam. Ancylostomatidae) 102- 106 IV.4.1.3.1 Overview of examined larvae and success rates of DNA 102 extraction and PCR amplification IV.4.1.3.2 Analysis of ITS-2 sequences 102- 103 IV.4.1.3.3 Analysis of 12s RNA mtDNA sequences 103- 104 IV.4.1.3.4 Comparison with Necator sequences from sympatric 104 monkeys Contents IV.4.1.3.5 Phylogenetic analysis 104- 106 IV.4.1.4 Trichostrongylid L3-larvae (Fam. Trichostrongylidae) 107- 110 IV.4.1.4.1 Overview of examined larvae and success rates of DNA 107 extraction and PCR amplification IV.4.1.4.2 Analysis of ITS-2 sequences 107 IV.4.1.4.3 Analysis of 12s RNA mtDNA sequences 107- 108 IV.4.1.4.4 Comparison with sequences of trichostrongylid larvae from 108 sympatric monkeys IV.4.1.4.5 Phylogenetic analysis 108- 110 IV.4.2 Identification and characterization of L3-larvae & free-living 111- 114 adults of Strongyloides spp.
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    A STUDY OF THE NEMATODE CAPILLARIA BOEHM! (SUPPERER, 1953): A PARASITE IN THE NASAL PASSAGES OF THE DOG By CAROLEE. MUCHMORE Bachelor of Science Oklahoma State University Stillwater, Oklahoma 1982 Master of Science Oklahoma State University Stillwater, Oklahoma 1986 Submitted to the Faculty of the Graduate College of the Oklahoma State University, in partial fulfillment of the requirements for the Degree of DOCTOR OF PHILOSOPHY May, 1998 1ht>I~ l qq ~ 1) t-11 q lf). $ COPYRIGHT By Carole E. Muchmore May, 1998 A STUDY OF THE NEMATODE CAPILLARIA BOEHM!. (SUPPERER, 1953): APARASITE IN THE NASAL PASSAGES OF THE DOG Thesis Appro~ed: - cl ~v .L-. ii ACKNOWLEDGMENTS My first and most grateful thanks go to Dr. Helen Jordan, my major adviser, without whose encouragement and vision this study would never have been completed. Dr. Jordan is an exceptional individual, a dedicated parasitologist, indefatigable and with limitless integrity. Additional committee members to whom I owe many thanks are Dr. Carl Fox, Dr. John Homer, Dr. Ulrich Melcher, Dr. Charlie Russell. - Dr. Fox for assistance in photographing specimens. - Dr. Homer for his realistic outlook and down-to-earth common sense approach. - Dr. Melcher for his willingness to help in the intricate world of DNA technology. - Dr. Charlie Russell, recruited from plant nematology, for fresh perspectives. Thanks go to Dr. Robert Fulton, department head, for his gracious support; Dr. Sidney Ewing who was always able to provide the final word on scientific correctness; Dr. Alan Kocan for his help in locating and obtaining specimens. Special appreciation is in order for Dr. Roger Panciera for his help with pathology examinations, slide preparation and camera operation and to Sandi Mullins for egg counts and helping collect capillarids from the greyhounds following necropsy.
  • Trypanoxyuris

    Trypanoxyuris

    Trypanoxyuris (Trypanoxyuris) minutus associated with the death of wild southern brown howler monkey,SCIENTIFIC Alouatta guariba COMMUNICATION clamitans, in Rio Grande do Sul, Brazil. 99 TRYPANOXYURIS (TRYPANOXYURIS) MINUTUS ASSOCIATED WITH THE DEATH OF A WILD SOUTHERN BROWN HOWLER MONKEY, ALOUATTA GUARIBA CLAMITANS, IN RIO GRANDE DO SUL, BRAZIL* J.F.R. Amato1, S.B. Amato1, C.Calegaro-Marques1, J.C. Bicca-Marques2 1Departamento de Zoologia, Instituto de Biociências, Universidade Federal do Rio Grande do Sul, CP 700, CEP 90001-970, Porto Alegre, RS, Brasil. E-mails: [email protected] ou [email protected] ABSTRACT This paper reports the death of a wild, subadult male of a southern brown howler monkey (bugio-ruivo), Alouatta guariba clamitans. The animal was found dead by the owner of a 60 ha. farm (Fazenda São Maximiano), located along the interstate road BR-116, km 308, Guaíba, State of Rio Grande do Sul, southern Brazil, 30º10'46,74"S, 51º23'30,78"W, in August 2000. The paper also describes the specimens of Trypanoxyuris (Trypanoxyuris) minutus found in the cecum. All organs were examined for helminths but were negative, except the cecum, which was full of macerated leaf litter and nematodes. The cecum wall was hyperemic, very thin, and distended, possibly by the large volume of material present. All the cecum contents were suspended in 5 liters of 0.85% saline physiological solution, from which a sample of 10% was taken and thoroughly examined. Six thousand one hundred and eighty-seven nematodes were counted in the sample (males + females). A total of 61,870 helminths were estimated in the entire cecal infrapopulation.