Molecular Analysis of Samples Collected from Humans and Dogs in Jamaica, Giardia, Assemblage a Was the Only Genotype Found

Genotypic diversity and distribution of Giardia – Central Europe versus the Caribbean

Doctoral thesis at the Medical University of Vienna for obtaining the academic degree

Doctor of Philosophy

Submitted by

Ms. Mellesia Lee

Supervisor: Associate Professor Dr. Julia Walochnik, MSc, PhD Institute of Specific Prophylaxis and Tropical Medicine Center for Pathophysiology, Infectiology and Immunology, Medical University of Vienna Kinderspitalgasse 15 A-1090 Vienna, Austria

Vienna, March 2017

Table of Contents

TABLE OF CONTENTS ...... 3

I. DECLARATION ...... 7

II. LIST OF FIGURES ...... 8

III. LIST OF TABLES ...... 9

IV. SUMMARY ...... 10

V. ZUSAMMENFASSUNG ...... 12

VI. PUBLICATIONS ARISING FROM THE DOCTORAL THESIS ...... 14

VII. ABBREVIATIONS ...... 17

VIII. ACKNOWLEDGEMENTS ...... 19

CHAPTER ONE: INTRODUCTION...... 20

1.1. OVERVIEW ...... 20

1.2. DISEASE BURDEN ...... 20

1.3. GIARDIA TAXONOMY ...... 21

1.3.1. Generic and species names ...... 22

1.3.2. Classification by morphological characteristics ...... 23

1.3.3. Sub-classification of G. duodenalis ...... 24

1.4. BIOLOGICAL CHARACTERISTICS OF GIARDIA: MORPHOLOGICAL FORMS ...... 26

1.4.1. Interphasic nuclei and cellular features ...... 28

1.4.2. Cytoskeletal structure: flagella ...... 30

1.4.3. Cytoskeletal structure: adhesive disc ...... 30

1.4.4. Cytoskeletal structure: median body ...... 30

1.4.5. Cytoskeletal structure: microfilaments ...... 30

1.4.6. Trophozoite surface antigens: variant surface proteins ...... 31

1.4.7. Occurrence and significance of antigenic variation ...... 32

1.4.8. Surface antigens: Cyst Wall Proteins ...... 32

1.4.9. Enzymes and energy metabolism ...... 33

1.5. LIFE CYCLE ...... 34

1.6. TRANSMISSION CYCLES ...... 35

1.6.1. Foodborne transmission ...... 36

1.6.2. Waterborne outbreaks ...... 37

1.6.3. Assemblages associated with waterborne and foodborne outbreaks ...... 38

1.7. GIARDIA IN ENVIRONMENTAL WATER AND MATRICES ...... 38

1.8. PATHOGENESIS AND CLINICAL MANIFESTATION ...... 39

1.8.1. Giardia in children ...... 39

1.8.2. Giardia in immunocompromised persons ...... 40

1.9. HOST IMMUNE RESPONSE ...... 40

1.10. TARGET GENES FOR MOLECULAR ANALYSIS ...... 40

1.11. PREVALENCE OF GIARDIASIS IN HUMANS ...... 42

1.12. PREVALENCE OF GIARDIASIS IN ANIMALS ...... 43

1.13. GIARDIASIS IN AUSTRIA ...... 43

1.14. GIARDIASIS IN JAMAICA ...... 44

1.15. DISTRIBUTION OF ASSEMBLAGES IN HUMANS AND ANIMALS ...... 45

1.16. DISTRIBUTION OF THE A SUB-ASSEMBLAGES ...... 47

1.17. ZOONOTIC POTENTIAL ...... 49

1.18. TREATMENT OF GIARDIASIS ...... 49

1.19. VACCINATION AGAINST GIARDIA ...... 50

1.20. LABORATORY DIAGNOSIS OF INFECTIONS ...... 50

1.20.1. Microscopy ...... 51

1.20.2. Direct smear ...... 51

1.20.3. Concentration techniques ...... 51

1.20.4. Enzyme-Linked Immunosorbent Assay ...... 52

1.20.5. Immunofluorescence assay ...... 52

1.20.6. Nucleic acid detection ...... 52

1.21. DNA EXTRACTION AND PCR AMPLIFICATION OF GIARDIA CYSTS FIXED IN FORMALIN ...... 53

1.22. AIMS ...... 55

1.22.1. Publication I ...... 55

1.22.2. Publication II ...... 55

1.22.3. Publication III ...... 55

CHAPTER TWO: MATERIALS AND METHODS ...... 58

2.1. ETHICS STATEMENT ...... 58

2.2. SAFETY ...... 58

2.3. SAMPLE COLLECTION ...... 58

2.4. MICROSCOPIC ANALYSIS ...... 58

2.5. MOLECULAR INVESTIGATION ...... 59

2.5.1. DNA extraction ...... 59

2.5.2. Conventional PCR...... 59

2.5.3. Nested PCR ...... 59

2.5.4. Sequencing PCR ...... 62

2.6. PREPARATION OF SAMPLES STORED IN FORMALIN ...... 62

2.6.1. Pre-treatment and extraction protocols ...... 62

2.6.2. NanoDrop® spectrophotometry ...... 63

2.6.3. PCR protocol for formalin-fixed Giardia samples ...... 63

CHAPTER THREE: RESULTS ...... 66

3.1. MICROSCOPIC AND MOLECULAR SCREENING OF SAMPLES IN JAMAICA ...... 66

3.2. GENOTYPE AND SUBTYPE DISTRIBUTION OF GIARDIA IN JAMAICA ...... 66

3.3. GIARDIA GENOTYPE AND SUBTYPE DISTRIBUTION IN AUSTRIA ...... 67

3.4. MOLECULAR ANALYSIS ON FORMALIN-FIXED SAMPLES ...... 68

3.5. MODIFIED PCR: 4-FOLD CONCENTRATION OF DNA POLYMERASE ...... 68

3.6. CORRELATION BETWEEN AMPLIFICATION OF SHORT FRAGMENTS AND STORAGE ...... 68

3.7. DNA QUALITY AND ANALYSIS OF NUCLEOTIDE SEQUENCES ...... 68

CHAPTER FOUR: DISCUSSION ...... 70

4.1. PUBLICATION I AND II ...... 70

4.2. PUBLICATION III ...... 74

4.3. CONCLUSION ...... 76

PUBLICATIONS ...... 77

REFERENCES ...... 88

CURRICULUM VITAE ...... 110

I. Declaration

II. List of figures

Figure 1.Phylgram showing sub-classification of G. duodenalis at the gdh locus...... 26

Figure 2. Morphological features of Giardia trophozoites and a cyst ...... 27

Figure 3. Giardia in a stool samples of an infected patient in Austria ...... 29

Figure 4. Nuclei in trophozoites and cysts ...... 29

Figure 5. Deconvolving micrograph of the Giardia cyst wall proteins ...... 34

Figure 6. Giardia DNA and nuclear replication ...... 35

Figure 7. The Giardia transmission cycle ...... 36

Figure 8. Workflow of Giardia-positive formalin fixed samples ...... 64

Figure 9. Nested PCR at the tpi and bg genetic loci ...... 67

III. List of tables

Table 1. Taxonomic classification of Giardia...... 22

Table 2. Morphological characteristics of Giardia species based on light microscopy ...... 24

Table 3. Giardia assemblages and most commonly infected hosts ...... 25

Table 4. Distribution of documented waterborne outbreaks of giardiasis ...... 38

Table 5. Giardia genes frequently used in molecular phylogenetical studies ...... 41

Table 6. Prevalence of giardiasis in developing countries ...... 42

Table 7. Prevalence of giardiasis in industrial countries...... 45

Table 8. Prevalence of infection by assemblages ...... 46

Table 9. Distribution of the A sub-assemblages in humans and animals and the respective gene targets used ...... 48

Table 10. Gene target for molecular analysis and type of assay established...... 53

Table 11. Primers used in identification of Giardia at the tpi, gdh and bg loci ...... 61

IV. Summary

Giardia duodenalis (also referred to as G. lamblia and G. intestinalis) is a flagellated protozoan of worldwide distribution and a common colonizer of the gastrointestinal tract of a wide range of domestic and wild mammals, amphibians and birds. The infectious stage of Giardia is the cyst, transmissible through consumption of contaminated water, food and by direct fecal-oral contact. Giardia is classified into species, assemblages and sub-assemblages, each with a unique predilection to infect different groups of animals. Human giardiasis is typically caused by Giardia duodenalis assemblages A and B, the latter often also referred to as G. enterica. Both assemblages are further divided into several sub-assemblages, whereby sub-classification within assemblage B is not well supported. Both assemblages are zoonotic and are also very common in dogs and cats. Therefore, an in-depth analysis and comparison of genotypes isolated from humans and animals is essential for our understanding of species distribution and zoonotic potential of the various subtypes. As clinical specimens are often stored in formalin, which may lead to DNA cross-linking and fragmentation over time and provides a challenge for molecular analyses, another aim of this study was to develop a protocol to successfully extract and amplify Giardia DNA from positive samples stored in formalin.

This dissertation provides the first data on the molecular diversity of Giardia spp. in Austria and in Jamaica. Sixty five microscopically positive human samples form Austrian patients were subjected to nested PCR of the triosephosphate isomerase (tpi), glutamate dehydrogenase (gdh) and ß-giardin (bg) genes. Fifty two PCR-positive samples were successfully sequenced to assess the genotype distribution. From Jamaica, a total of 285 human and 225 dog stool samples were screened using conventional PCR targeting the SSU rDNA. This was followed by nested PCR on 63 positive samples targeting the tpi, gdh and bg genes. Moreover, forty five old Giardia-positive samples preserved in 1.5% formalin from a strain collection were used to establish a modified protocol to extract Giardia DNA from formalin-fixed samples and the effectiveness of this protocol was assessed by nested PCR amplification targeting the tpi, gdh and bg genes.

In Austria, assemblage B accounted for 65.4% (34/52) and assemblage A for 34.6% (18/52). A high level of genetic diversity was evident, with sub-assemblage AI being responsible for 9.6% (5/52), AII for 25% (13/52), BIII for 19.2% (10/52) and BIV for 46.2% (24/52) of the infections. There were no mixed infections. In Jamaica, 6.7% (19/285) of the human samples and 19.6% (44/225) of the canine samples were positive. On sequence analysis, assemblage A was predominant, with sub-assemblage AII accounting for the majority of infections in both, humans and dogs (79.0%; 15/19 and 70.5%; 31/44). Sub-assemblage AI was identified at much lower

10 rates, of 15.8% (3/19) and 29.5% (13/44) in humans and dogs, respectively. DNA was successfully amplified from more than half of the 45 formalin-fixed samples, whereby the tpi gene (64.4%) was far more successful than the bg (40%) and gdh (20%) locus in producing positive amplicons. All three loci provided the possibility of differentiating Giardia at the species level. Targeting the tpi gene, 43% (12/29) produced meaningful sequence data, at the bg and gdh loci, these were 50% (9/18) and 22.2% (2/9), respectively.

Altogether, the two distinct geographic locations showed clear differences in genotypes causing giardiasis in humans. While, the high genetic diversity at the sub-assemblage level in Austria corroborates the assumption that the majority of infections are imported, the samples from Jamaica indicate a zoonotic transmission cycle as evidenced by shared Giardia assemblages between humans and dogs. Moreover, it was shown that also Giardia cysts that had been stored in formalin for more than 10 years can be successfully genotyped.

V. Zusammenfassung

Giardia duodenalis (auch als G. Lamblia und G.Intestinalis) ist ein weltweit verbreiteter begeißelter Protist, der im Gastrointestinaltrakt des Menschen aber auch einer Reihe von Haus- und Wildtieren, Amphibien und Vögeln. Das infektiöse Stadium ist die Zyste, die durch kontaminiertes Wasser, Nahrungsmittel oder durch direkten fäko-oral Kontakt übertragen wird. Die Gattung Giardia wird in Arten, Assemblages und Sub-Assemblages, mit jeweils unterschiedlicher Wirtsspezifität unterteilt. Giardiose beim Menschen wird in erster Linie durch G. duodenalis Assemblage A und B verursacht, wobei Letztere auch als G. enterica bezeichnet wird. Beide Assemblages werden in jeweils mehrere Sub-Assemblages unterteilt, wobei zum Teil Inkonsistenzen in der Einteilung bestehen. Beide Assemblages sind zoonotisch und kommen auch bei Tieren, insbesondere bei Haustieren wie Hunden und Katzen, vor. Daher ist ein Vergleich von Isolaten aus Mensch und Tier und aus unterschiedlichen geographischen Regionen auf molekularer Ebene von großer Bedeutung, um die Verbreitung und das zoonotische Potential der unterschiedlichen Subtypen aufzuklären. Da klinische Proben oft in Formalin aufbewahrt werden, war ein weiteres Ziel dieser Arbeit die Entwicklung eines Protokolls zur erfolgreichen Extrahierung und Amplifizierung von Giardia DNA aus Formalin- fixierten Proben.

Diese Dissertation liefert die ersten Daten zur molekularen Epidemiologie von Giardia spp. in Österreich und Jamaika. In Österreich wurden 65 mikroskopisch positive humane Proben mittels Nested-PCR an drei genetischen Loci untersucht, nämlich den Genen für die Triosephosphate Isomerase (tpi), die Glutamat-Dehydrogenase (gdh) und das ß-Giardin (bg). Von 52 Proben konnten aussagekräftige Sequenzen für die Genotypisierung gewonnen werden. In Jamaika wurden 285 Human-Proben und 225 Proben von asymtomatischen und symptomatischen Hunden zunächst mit konventioneller PCR auf Giardien untersucht, und die 63 positiven Proben anschließend ebenfalls mittels Nested-PCR und Sequenzanalyse genotypisiert. Außerdem wurden 45 Formalin-fixierte positive humane Proben aus einer Stammsammlung in die Untersuchungen mit einbezogen und ein modifiziertes Protokoll zur Isolierung von Giardia DNA aus diesen Proben entwickelt.

Die Analyse der Proben aus Österreich ergab, dass Assemblage B mit 65,4% (34/52) den Großteil der Infektionen beim Menschen ausmachte, und Assemblage A lediglich 34,6% (18/52). Außerdem zeigte sich ein hohes Maß an genetischer Vielfalt, wobei Sub-Assemblage AI mit 9,6% (5/52), AII mit 25 % (13/52), BIII mit 19,2% (10/52) und BIV mit 46,2% (24/52) vertreten waren. Mischinfektionen traten nicht auf. In Jamaika ergab die molekulare Analyse eine Prävalenz von 6,7% beim Menschen und 19,6% beim Hund. Insgesamt war Assemblage A

12 vorherrschend, wobei Sub-Assemblage AII die Mehrzahl der Infektionen, sowohl beim Menschen (79,0%, 15/19) als auch beim Hund (70,5%, 31/44), ausmachte. Sub-Assemblage AI war mit nur 15,8% beim Menschen (3/19) und 29,5% beim Hund (13/44) vertreten. Aus mehr als der Hälfte der 45 Formalin-fixierten Giardia-Proben konnte erfolgreich DNA isoliert werden, wobei das tpi-Fragment mit 64,4% (29/45) am besten amplifziert wurde, während die Amplifikation des längeren bg-Fragments nur bei 40% (18/45) und die des gdh-Fragments nur bei 20% (9/45) funktioniert hat. Alle drei Loci ermöglichten eine Differenzierung der Giardia- Isolate auf Artniveau. Insgesamt konnten von 43% (12/29) der Proben aussagekräftige tpi-Gen- Sequenzen, von 50% (9/18) aussagekräftige bg-Sequenzen und von 22,2% (2/9) aussagekräftige gdh-Sequenzen ermittelt werden.

Es hat sich gezeigt, dass sich die beiden unterschiedlichen geografischen Regionen ganz deutlich in der Prävalenz der verschiedenen Genotypen unterscheiden. Während die hohe genetische Vielfalt der österreichischen Giardia-Isolate dafür spricht, dass es sich bei der Mehrzahl der Infektionen um importierte Infektionen handelt, spricht das Vorherrschen ein und desselben Genotyps bei Mensch und Hund in Jamaika für das Vorliegen eines zoonotischen Übertragungszyklus. Außerdem konnte gezeigt werden, dass mit dem modifizierten Protokoll zur DNA-Isolierung auch Giardia-Zysten, die über 10 Jahre in Formalin gelagert wurden, noch erfolgreich genotypisiert werden können.

VI. Publications arising from the doctoral thesis

Publication I

Multilocus sequence analysis of Giardia spp. isolated from patients with diarrhea in Austria

Authors: Mellesia F. Lee, Herbert Auer, John F. Lindo, Julia Walochnik

Journal: Parasitology Research. DOI:10.1007/s00436-016-5306-9

Contribution to publication I

Conceptualised and designed study; conducted all experiments, data analysis and interpretation and writing of the manuscript.

Publication II

Molecular epidemiology and multilocus sequence analysis of potentially zoonotic Giardia from humans and dogs in Jamaica

Authors: Mellesia F. Lee, Paul Cadogan, Sarah Eytle, Sonia Copeland, Julia Walochnik, John F. Lindo

Journal: Parasitology Research. DOI: 10.1007/s00436-016-5304-y

Contribution to publication II

Conceptualised and designed study; sample collection and shipment; conducted all experiments, data analysis and interpretation and writing of the manuscript.

Publication III

Successful extraction and PCR amplification of Giardia duodenalis DNA from formalin- fixed stool

Authors: Mellesia F. Lee, Herbert Auer, John F. Lindo and Julia Walochnik

Intended Journal: PLosOne

Contribution to publication III

Conceptualised and designed study; troubleshooting experiments for developing modified protocol for extraction and amplification, analysis and interpretation of data, drafting of manuscript.

VII. Abbreviations

BG: beta gardin cDNA: complementary deoxyribonucleic acid

CWP: cyst wall protein

CXXC: Cys-X-X-Cys

DAPI: 4',6-diamidino-2-phenylindole

DNA: deoxyribonucleic acid dNTP: deoxynucleotide triphosphate

EDTA: ethylene diamine tetra-acetic acid

EF 1α: elongation factor 1 alpha

ELISA: enzyme-linked Immunosorbent assay

ESV: excystation specific vesicle

FECT: formalin ether concentration technique

FMS: faculty of medical sciences

HSP: heat shock protein

IFA: immunofluorescent assay

IFN: interferon

IGS: Intergenic spacer

IL: interleukin

ISPTM: institute of specific prophylaxis and tropical medicine

ITS: internal transcribed spacer

MAb: monoclonal antibody

MedUni: medical university

MLG: multilocus genotypes

NADH: nicotinamide adenine dinucleotide

NPHL: national public health laboratory

ORF: open reading frame

PCR: polymerase chain reaction

PFOR: pyruvate ferredoxin oxidoreductase

RFLP: restriction fragment length polymorphism rRNA: ribosomal ribonucleic acid

RT-PCR: real time reverse transcription polymerase chain reaction

SSU rDNA: small subunit ribosomal deoxyribonucleic acid

Th: t-helper

TPI: triose phosphate isomerase

UHWI: university hospital of the West Indies

UWI: university of the West Indies

VSP: variant specific protein

WGA: wheat germ agglutinin

ZnSO4: zinc sulfate flotation

VIII. Acknowledgements

This dissertation reflects an accumulation of studies conducted over three years to obtain a Doctor of Philosophy in Molecular Parasitology. I extend gratitude to my supervisors who invested the time and effort necessary for the completion of degree. With many years of experience in the field, their exceptional level of knowledge, skills and experience are exactly what was required. I would like to thank all doctors, assistants and technicians from the various veterinary practices and hospitals who assisted in sample collection, storage and transportation.

The administrators at the University of the West Indies, Jamaica provided significant contributions in the form of financial and academic support making all aspects of this research possible. The most significant technical contribution came from Iveta Häfeli from the Institute of Specific Prophylaxis and Tropical Medicine Center for Pathophysiology, Infectiology and Immunology in Austria. Her many years of experience in DNA sequencing and analysis made the most important aspects of this research possible. I also thank the very smart and insightful Martina Koehsler who provided assistance in many vital aspects of this project.

CHAPTER ONE: INTRODUCTION

1.1. Overview

Giardia duodenalis is the earliest protozoan parasite discovered. The parasite was first described in 1681 by the Dutch textile manufacturer, lens grinder and scientist Antony van Leeuwenhoek who had performed microscopic examination of his own diarrheal stool (Cox, 2002). Members of the genus are common causes of gastrointestinal infections in humans and other vertebrate animals. Giardia has a worldwide distribution with more than 200 million people in Asia, Africa, Latin America, Europe and Australia having symptomatic infection and up to 500,000 new cases per year (Thompson et al., 2000). The parasite infects 2% of adults and 6- 8% of children in developed countries and nearly 33% of people in developing countries, contributing to 2.5 million deaths annually (Adam, 2001; CDC, 2012; Choy et al., 2014). Those most at risk of becoming infected are children in low-income countries, children in day-care settings, those in close contact with infected persons, persons who consume contaminated drinking water, especially campers and backpackers drinking water from lakes or rivers, travellers to endemic regions, persons who come in contact with infected animals and those participating in activities that involve anal-oral contact (CDC, 2012; CDC, 2015).

1.2. Disease burden

The occurrence or giardiasis in adults and children is significant from clinical and public health perspectives (Nikolic et al., 2011; Thompson et al., 2008a). The clinical presentation is influenced by the host’s physiological status and infection may range from asymptomatic to mild or fulminant giardiasis. Symptomatic giardiasis is believed to be underreported for a multitude of reasons and may be grossly underestimated (Yoder et al., 2010). The true burden in the United States is estimated to be as many as 2 million cases annually (Yoder et al., 2010). The total number of reported cases in the United States from 2006-2008 were 20,000 annually with an incidence of 7.4-7.6/ 100,000 (Yoder et al., 2010). Approximately 28.5 million cases are reported in China per year (Zheng et al., 2014). In countries such as Norway, Finland and Sweden, for every reported case of giardiasis, there were 254 to 867 cases undetected and/or unreported as such, suggesting a true incidence of symptomatic infections of 4,670 / 100,000 annually in the general population (Horman et al., 2004). One factor that may be responsible for such high numbers of unreported cases may include the fact that infections are usually self-limiting (Halliez & Buret, 2013). Therefore, a large number of persons with giardiasis usually do not present to a doctor. Another factor that should be taken into account is the mode in which the diagnostic parasitic products are excreted. The trophozoites and cysts may be excreted intermittently or

20 occasionally only, resulting in false negatives. As a result up to three consecutive samplings are recommended for diagnosis (Anderson et al., 2004).

1.3. Giardia taxonomy

The early taxonomy of Giardia was complicated by inconsistencies in species classification. Filice (1952) classified Giardia according to its host species rather than its morphological characteristics (Thompson et al., 2000). Then Giardia species were classified mainly based on morphological characteristics of trophozoites and cysts isolated from a variety of human and animal hosts. However, this method remained unsatisfactory because it does not correlate with current molecular approaches (Feng & Xiao, 2011). It is only recently that molecular tools have been developed to accurately classify Giardia species isolated from a wide range of mammalian hosts (Adam, 2001; Rayani et al., 2014). Until today, there are no well-defined criteria for species designation and classification beyond the genus level remains controversial (Adam, 2001).

Initial hierarchal classification placed Giardia in the phylum Metamonada (Nohýnková et al., 2006), the class Trepomonadea (Adam, 2001) the order of Diplomonadida, which is characterised by possessing two karyomastigonts with four flagella, two nuclei, no mitochondria, and no Golgi complex, and the family Hexamitidae (Adam, 2001). However, recent taxonomical classification assigned Giardia to the supergroup Excavata, second rank Giardiinae, subphylum Trichoza, superclass Eopharyngia, subclass Diplozoa and family Giardiidae (Table 1).

Table 1. Taxonomic classification of Giardia (Plutzer et al., 2014)

Taxonomic hierarchy Taxonomic rank

Higher rank Excavata

Second rank Giardiinae

Phylum Metamonada

Subphylum Trichozoa

Superclass Eopharyngia

Class Trepomonadea

Subclass Diplozoa

Order Giardiida

Family Giardiidae

Genus Giardia

1.3.1. Generic and species names

The generic name Giardia was first introduced by Kunstler in 1882 for a flagellate found in the intestines of tadpoles (Lebbad et al., 2010; Lujan & Svärd, 2011). In 1888, Blanchard suggested that the generic name should be in commemoration of Lambl who provided a detailed description of the parasite in 1859 and at that time referred to the organism as Cercomonas intestinalis (Adam, 2001; Lujan & Svärd, 2011). As a result, the parasite was subsequently referred to as Giardia lamblia. This suggestion was not challenged until 55 years later when Alexeieff argued that there was no taxonomic rationale to its reference. It was also well accepted that Cercomonas was not the correct generic name as evidenced by the description of the same flagellate in tadpoles by Kunstler in 1882 (Lujan & Svärd, 2011). Seven years prior to Kunstler’s discovery, Davaine described Giardia in rabbits which he called Hexamita duodenalis (Lujan & Svärd, 2011). Even though the ascribed generic name was criticized as inaccurate, Filice proposed that Hexamita duodenalis should be used (Lujan & Svärd, 2011). It was agreed that a single specific name was to be used for all forms of human Giardia infections and duodenalis, which has priority over intestinalis, was more fitting in accordance to the Rules of Zoological Nomenclature (Feng & Xiao, 2011; Lujan & Svärd, 2011). Although the specific name Giardia

22 duodenalis would appear to be correct according to such standards, intestinalis and lamblia are considered synonyms specifically for isolates of human origin (Lujan & Svärd, 2011).

1.3.2. Classification by morphological characteristics

The most commonly known Giardia species include G. duodenalis (primarily infecting humans and domestic mammals), G. muris (rodents), G. agilis (amphibians), and G. psittaci and G. ardeae (birds) (Thompson & Monis, 2004a). Initially, Giardia species were mainly classified based on morphology of cysts when Filice proposed that only morphological differences should be used as basis for classification (Lujan & Svärd, 2011). Species were classified based on the overall shape of the trophozoites and the median bodies which were considered most valuable (Erlandsen & Bemrick, 1987; Lujan & Svärd, 2011). Only three morphologically distinct groups existed: the first characterised by pyriform-shaped or pear-shaped trophozoites and “claw- hammer” median bodies designated as G. duodenalis (Table 2). The second, G. muris, was characterized by a more rounded trophozoite with rounded median bodies and primarily infects rodents. The third group consisted of trophozoites that were long or resembling a teardrop with a short adhesive disk and club-shaped median bodies, and were classified as G. agilis (Table 2) (Adam, 2001; Lujan & Svärd, 2011). In addition to the three morphological groups, isolates from birds with similar morphological features as G. duodenalis were characterized ultrastructurally. This warranted species recognition and they were referred to as G. psittaci and G. ardeae (Erlandsen & Bemrick, 1987). However, morphological features of G. ardeae were more in resemblance with G. muris (Erlandsen et al., 1990a; Lujan & Svärd, 2011). Eventually, such classification would cause great inconsistencies due to phenotypic variation which was especially reported within G.duodenalis (Lujan & Svärd, 2011). Such variations involved differences in dimensions and shape of trophozoites which questioned the value of morphologic classification (Thompson & Monis, 2004b).

Table 2. Morphological characteristics of Giardia species based on light microscopy (Adam, 2001)

Species Morphology Trophozoite Host (length/width in μm)

G. duodenalis Pear shaped 12-15/6-8 Humans, dogs, cats, wild animals

G. agilis Teardrop shaped 20-30/4-5 Amphibians

G. muris Short and rounded; small 9-12/5-7 Rodents rounded median body G. psittaci Pear shaped ~14/~6 Psittacine birds

G. ardeae Pear shaped ~10/~6.5 Herons

1.3.3. Sub-classification of G. duodenalis

There are eight assemblages (A-H) of G.duodelialis based on phenotypic and genetic analysis. Assemblage A and B have broad host ranges and infect predominantly humans and animals including dogs and cats (Table 3). Assemblages C, D, E, F, and G have narrow host ranges and strong specificities (Figure 1) (Feng & Xiao, 2011). Assemblages C and D are most commonly found in dogs, foxes and other canines (Table 3). Assemblage F and G are commonly found in cats and rodents, while H was identified in marine mammals (Feng & Xiao, 2011). Phylogenetic analysis of assemblage A and B resulted in sub-clustering within these assemblages; namely AI, AII, AIII and AIV which were derived from nucleotide sequence data from small subunit ribosomal deoxyribonucleic acid (SSU) rDNA and other house-keeping genes coding for glutamate dehydrogenase (gdh), β-gardin (bg), elongation factor 1 alpha (ef1α) and triose phosphate isomerase (tpi) (Feng & Xiao, 2011). Separation of assemblages AI and AII was done initially using a combination of allozyme and phylogenetic analysis of assemblage A at the gdh locus (Figure 1). Further analysis also supported this separation at the tpi and bg locus. A third classification into assemblages AIII was derived based on analysis of the same three gene loci. Classification of assemblage B into BIII and BIV was not supported by allozyme electrophoretic studies (Minetti et al., 2015).

Table 3. Giardia assemblages and most commonly infected hosts (Adam, 2001; Feng & Xiao, 2011)

Assemblage Species commonly infected Taxonomic names

AI Humans and animals (dogs, cats, beavers, livestock, deer, pigs) G. duodenalis

AII Humans G. duodenalis

AIII & AIV Animals G. duodenalis

B Humans and animals (livestock, rodents, beavers, chinchillas, marmosets) G. enterica

C & D Dogs & foxes G. canis

E Goats, cattle, pigs, sheep, alpacas G. bovis

F Cats G. cati/G.felis

G Rats G. samondi

H Marine mammals -

Figure 1.Phylgram showing sub-classification of G. duodenalis at the gdh locus (Feng & Xiao, 2011)

1.4. Biological characteristics of Giardia: morphological forms

Giardia exists in two morphological forms; as trophozoite and as cyst (Figure 3). Trophozoites are fragile and are found within the gastrointestinal tract of the infected hosts. They are approximately 10 to 15 μm in length and 7 to 10 μm in width (Table 2) (Adam, 2001). The cytoskeletal components of the trophozoite include median bodies, sucking disk on its ventral surface and four pairs of flagella (anterior, posterior, caudal and ventral) (Figure 2). Other organelles include two nuclei, lysosomal vacuoles, and ribosomal vacuoles (Adams et al., 2004). Organelles such as smooth endoplasmic reticulum, and nucleoli have not been identified (Adam, 2001). Due to its concave surface, the ventral disk is the site of attachment to the abdominal surface acting like a suction pad in the infected host and is presumed to be assisted by the

26 hydrodynamic movement of the flagella (House et al., 2011). The flagella, consist of nine pairs of outer microtubules and two inner ones (Adam, 2001). The rim of the ventral disk also contains contractile proteins, actin, myosin and tropomyosin which pharmacological agents such as metronidazole acts on to prevent adherence (Adam, 2001).

Figure 2. Morphological features of Giardia trophozoites (a, c, d) and a cyst (b) (Roberts et al., 2013)

1.4.1. Interphasic nuclei and cellular features

Most important cellular features of Giardia are the two bilaterally asymmetrical nuclei that are present within trophozoites (Figure 4). Both are located on the anterior aspect of the cell between the longitudinal axis (Lujan & Svärd, 2011). Giardia nuclei divide by binary fission, and after division, trophozoites encyst giving rise to four nuclei within each cyst (Figure 4 and 6) (Lujan & Svärd, 2011). Other important cellular components are glycogen granules which act as energy reserves for trophozoites and cysts (Lujan & Svärd, 2011). Organelles such as mitochondria and perioxiosomes present in higher eukaryotes were reported to be entirely lacking in Giardia (Adam, 2001). However, mitochondrial remnants in the form of mitosomes have been demonstrated, bounded by a double membrane which led to the conclusion that the organism is not primarily amitochondrial (Tovar et al., 2003). The two bilaterally asymmetrical nuclei are oval shaped with each measuring around 1 µm in diameter (Adam, 2001). Morphological studies have shown that the two nuclei do not have the same function (Lujan &

Svärd, 2011). DNA replication of each nucleus begins during the G1 phase of the interphase where protein synthesis takes place. This is followed by chromosomal replication during the S phase and eventually cytokinesis (Figure 6). After DNA replication, trophozoites divide by binary fission (McInally & Dawson, 2016).

Cysts are slightly smaller than trophozoites and range from 8 to 14 μm in length and 7 to 14 μm in width (Table 2). They are generally rigid and can survive in various environmental conditions. Cysts were shown to survive for as long as 56 days at 0°C to 4°C, and for 14 days at 17°C to 20°C (Erickson & Ortega, 2006). Longer survivals of up to 84 days were seen in river water at 0°C to 4°C (Erickson & Ortega, 2006).

A B

Figure 3. Giardia in a stool samples of an infected patient in Austria; A) trophozoite, B) cyst. Giemsa stain. Bar =10 μm (Walochnik & Aspock, 2014)

Figure 4. Nuclei in trophozoites and cysts (Ratner et al., 2008)

Left - Red stains (WGA) represents perinuclear regions and small vesicles of permeabilized trophozoites, nuclei are stained with DAPI. Middle - WGA binds in a punctate manner to the surface of a non- permeabilized trophozoite. Right- In cysts, WGA binds in a perinuclear pattern and densely labels membranes (arrowheads), these are closely opposed to the walls (arrows), and are stained green with anti-CWP1 antibodies.

1.4.2. Cytoskeletal structure: flagella

The four pairs of flagella begin in the basal bodies between the two nuclei and are named based on the positions in which they are located; caudal, ventral, anterolateral, posterior or posterolateral each assisting in propulsion of the trophozoite (Campanati et al., 2002; Lenaghan et al., 2011; McInally & Dawson, 2016). All flagella with the exception of the ventral ones are similar to those of other eukaryotic cells (Lujan & Svärd, 2011). The ventral flagella are anchored by a thin filament 30 kDa glycoprotein assumed to be its main component (Woessner & Dawson, 2012). Another flagellar protein; alpha 19-gardin was identified in the ventral flagellum and is associated with the microtubules (Saric et al., 2009; Vahrmann et al., 2008). Flagellar proteins anchoring to microtubules assist in motility (Nosala & Dawson, 2015).

1.4.3. Cytoskeletal structure: adhesive disc

The adhesive disk also known as ventral disk consists of microtubules anchored to the plasma membrane by microfilaments (Lourenço et al., 2012; Tumova et al., 2007). It is functionally involved in aiding the attachment of trophozoites to enterocytes as evidenced by dome-shaped imprints left behind in brush borders (Lujan & Svärd, 2011). The main components of the adhesive disk are gardins and tubulins (Lujan & Svärd, 2011; Piva & Benchimol, 2004).

1.4.4. Cytoskeletal structure: median body

Median bodies are composed of contractile proteins which are found in mitotic trophozoites (Lujan & Svärd, 2011; Piva & Benchimol, 2004). They are made of microtubules and have been used as a taxonomic tool for classification (Lujan & Svärd, 2011). Other proteins associated with the microtubules are similar to the beta-gardin gene (Hagen et al., 2011). The functional role of the median body remains unknown and is proposed to be the site of microtubule nucleation and interphase reserve for microtubule spindle assembly (Lujan & Svärd, 2011; Woessner & Dawson, 2012).

1.4.5. Cytoskeletal structure: microfilaments

Microfilaments have been seen and Giardia actin shows high resemblance to other eukaryotic species (58% amino acid identity) (Paredez et al., 2011). However, by laser scanning microscopy morphological changes can be observed after treatment with drugs that interfere with microfilaments, indicating their presence (Castillo-Romero et al., 2009; Correa & Benchimol, 2006).

1.4.6. Trophozoite surface antigens: variant surface proteins

The cell membrane of the trophozoites consists of major antigens known as variant surface proteins (VSPs) (Li et al., 2013). VSPs are a unique protein family that are rich in cysteine and exhibit four amino acid CXXC motifs and a hydrophobic membrane-spanning region at the extreme C-terminus (Mowatt et al., 1991; Nash & Mowatt, 1992). VSPs spontaneously undergo antigenic variation in vivo and in vitro and trophozoites have the ability to express up to 200 different VSPs, estimated based on analysis of vsp gene expression (Prucca & Lujan, 2009). However, only one vsp gene can be expressed on the surface of a trophozoite at a given time and a change occurs every 6-13 generations. Host antibodies are produced against VSP antigens to eliminate the parasite and frequent switching of VSPs may result in failure of the host immune system to effectively recognize and eliminate the pathogen. This antigenic property is effective in causing prolonged infections, with malabsorption of nutrients and weight loss seen in many Giardia cases (Faubert, 2000; Ish-Horowicz et al., 1989; Li et al., 2013; Pickering et al., 1984; Prado et al., 2005).

Much of the analyses of vsp gene expression were done on the G. duodenalis WB strain (assemblage A) which was first isolated from a patient in Afghanistan who had chronic symptomatic giardiasis. Another strain, the GS strain (assemblage B) was originally isolated from a patient in the US and the genome of both, WB and GS, were recently sequenced (Franzen et al., 2009; Jerlström-Hultqvist et al., 2010). The two genomes show 77% nucleotide and 78% amino acid identity in their protein coding regions (Franzen et al., 2009; Jerlström- Hultqvist et al., 2010). A comparison between both isolates identified 28 unique GS and 3 unique WB coding genes, and the vsp repertoires of the isolates were completely different (Franzen et al., 2009).

Six VSPs have shown to be localized to the cell membrane (Almirall et al., 2013), these are; VSPA6, TSA417, VSPH7, G3M, VSP1267 and VSP9B10A (Franzen et al., 2009; Jerlström- Hultqvist et al., 2010; Li et al., 2013). They were recognized using surface labelling and electron microscopic analysis of monoclonal antibodies (Mab) specific for the detection of each on the cell surface (Li et al., 2013; Serradell et al., 2016). It is possible that VSPs of different Giardia species may differ considerably especially for zoonotic assemblage A and B; one or more of these VSPs may predominate in specific subtypes or may not be present at all. A comparative analysis would be needed to make correlations in order to elucidate possible mechanisms of establishing and maintaining zoonotic infections.

1.4.7. Occurrence and significance of antigenic variation

Initial studies of surface antigens of Giardia species showed differences between strains detected by Enzyme-Linked Immunosorbent Assay (ELISA) and immunoelectrophoresis (Adam, 2001). Surface antigens varied in molecular mass from approximately 50 to 200-kDa in one study (Li et al., 2013). VSPA6 is a 170-kDa protein which was consistently expressed on trophozoites of strain WB as detected using MAb 6E7. VSP1267 is a 64-kDa surface antigen which reacts to tagged MAb 5C1 (Adam et al., 2010; Li et al., 2013). When human volunteers were infected with cloned GS trophozoites, four of four became infected with trophozoites expressing VSP H7 (72-kDa) (Adam, 2001). This finding indicated that VSP H7 is the most dominant surface antigen on GS trophozoites.

Investigation of antigenic variation using Giardia clone GS/M-83-H7 showed VSP H7 as its major VSP (Bienz et al., 2001). Therefore, the possibility exists that this may be the same for most Giardia species. Similarly, VSP9B10 is preferentially expressed on most Giardia trophozoites isolated from patients in Argentina (Bienz et al., 2001; Carranza et al., 2002). Analysis of vsp gene expression on the Giardia WB strain showed that VSP9B10 and VSP1267 may be expressed simultaneously (Carranza et al., 2002). The expression of VSP9B10 was reported in Giardia assemblage A (WB strain ) but not in assemblage B (GS strain) (Carranza et al., 2002). However, the expression patterns of the vsp genes are entirely unknown in many of the Giardia assemblages and sub-assemblages. Variations in the zoonotic potential of assemblages A and B may be linked to different expressions of VSPs.

1.4.8. Surface antigens: Cyst Wall Proteins

During encystation several Giardia genes are up-regulated which constitute the expression of surface proteins. These are known as Cyst Wall Proteins (CWPs) which are transported by encystation-specific secretory vesicles (ESVs) (Figure 5). The three main CWPs constituting the extracellular domain include CWP1, CWP2, and CWP3 (Figure 5). These proteins assemble mainly as extracellular matrix but also have portions that exist largely as carbohydrates such as N-acetylgalactosamine homopolymer (Chatterjee et al., 2010; Manning et al., 1992). The three CWPs also contain sites for N-linked glycosylation, C-terminal rich in cysteine (CRR) and N- terminal rich in leucine (LRRs). Cys-rich regions in CWP1 have been used as targets for diagnostic purposes while CWP2 possess an additional charge at its C-terminal which was previously used to reduce cyst formation in mice through immunization (Boone et al., 1999; Chatterjee et al., 2010; Lee et al., 2009; Meng et al., 1996; Sun et al., 2003). Other Cys-rich, non-VSP proteins have been categorized including EGF-like cyst proteins and High Cysteine

Membrane Proteins (HCNCp) which are targets in the wall of Giardia cysts (Chiu et al., 2010; Davids et al., 2006).

1.4.9. Enzymes and energy metabolism

Major enzymes acquired through lateral transfer for utilization of energy and other metabolic functions in Giardia include pyruvate oxidoreductase or pyruvate ferredoxin oxidoreductase (PFOR), alcohol dehydrogenase and pyrophosphate (Walochnik & Duchêne, 2016). PFOR is a membrane associated cytosolic enzyme which decarboxylates pyruvate in the presence of coenzymes to produce acetyl-CoA, CO2 and ferredoxin (Emelyanov & Goldberg, 2011). PORF identified in Giardia is different from the pyruvate dehydrogenase enzyme complexes that are found in the mitochondria of higher eukaryote and is a target for anti-giardial drugs (Walochnik & Duchêne, 2016). Four alcohol dehydrogenase enzyme-encoding genes have been identified in Giardia geneome. A 97-KDa has been expressed in trophozoites but downregulated during encystation. Others including the 44-kDa and 45-kDa proteins are usually upregulated during encystation (Walochnik & Duchêne, 2016). The mechanisms involved in oxidative stress protection in most eukaryotes using enzymes such as superoxide dismutase, peroxide and glutathione are lacking in Giardia trophozoites (Walochnik & Duchêne, 2016). A H2O-producing NADH oxidase, NADH peroxidase and other thioredoxin reductase-like disulphide reductase, cysteine, thioglycolate and other low molecular weight compounds are associated with oxidative stress protection (Walochnik & Duchêne, 2016).

For energy production, Giardia trophozoites under anerobic or microaerphilic conditions, use glycolysis to catabolize glucose by the Embden-Meyerhof-Panas and pentose phosphate pathways (Lujan & Svärd, 2011; Walochnik & Duchêne, 2016). Both pathways rely on pyrophospahate for phosphorylation producing catabolic end products such ethanol, acetate, alanine and CO2 (Lujan & Svärd, 2011).

Even though Giardia is well adapted to the environment, the organism has to scavenge thriving- flora for all its biosynthetic pre-cursor in surrounding environment (Lujan & Svärd, 2011). The organism lacks de novo lipid, purine and pyrimidine oxidative phosphorylation. Therefore, purine and pyrimidine bases are scavenged exogeniously and trophozoites relies on preformed lipids in the intestine (Yichoy et al., 2011).

Figure 5. Deconvolving micrograph of the Giardia cyst wall proteins (Chatterjee et

al., 2010)

A-Trophozoites stained for CWP1 (green) and plasma with alexafluor (red) showed that CWP1 was not expressed in trophozoites. B-Plasma membrane labelled with alexafluor. C- Shows vesicles labelled with CWPI (green). D-smaller and lesser clumps of CWP1– labelled vesicle. E-ESV staining CWP2. F-Bulk of CWP3 remains in ESV. G-ESV containing CWP3

1.5. Life cycle

Giardia infection is initiated when an appropriate host ingests infectious cysts. Cysts are stimulated by acidic pH in the stomach resulting in excystation in the proximal small intestine and the release of motile trophozoites (Figure 7) (Adam, 2001). Cysteine proteinases are necessary for encystation and excystation, while host proteases (trypsin and/or chymotrypsin) are required for excystation (Chatterjee et al., 2010; DuBois et al., 2008). After excystation, the four pairs of flagella emerge and trophozoites develop a ventral flange with protrusions extending from the cytoplasm (Adam, 2001; Hetsko et al., 1998; Lujan & Svärd, 2011). Trophozoites adhere to enterocytes and replicate by binary fission in the duodenum (Adam, 2001). After replication, they undergo encystation in the large intestine and cysts are excreted in

34 formed stool and survive well in the environment where they remain infectious for two weeks at 25°C and for up to 11, 7 and 1 week in water, soil and cattle faeces, respectively (Adam, 2001; Olson et al., 1999). In hosts experiencing diarrhea, trophozoites may be excreted unchanged and die shortly after (Adam, 2001).

Figure 6. Giardia DNA and nuclear replication (Svard et al., 2003)

1.6. Transmission cycles

Very low quantities of cysts are sufficient to initiate Giardia infections; the infectious dose has been reported to lie as low as 10 cysts (Thompson et al., 2008b). Based on epidemiological data, the main transmission cycles generally recognized are waterborne and environment, or foodborne and person to person (Lujan & Svärd, 2011). Waterborne transmission is often associated with sporadic outbreaks while person-to-person transmission is typified by high prevalence in institutions such as day-care centres. This is analogous to animal-to-animal

35 transmission where host-adopted assemblage E is frequently isolated from calves in pens (Geurden et al., 2008). Similarly, infections in dogs and cats may occur in crowded environments such as shelters or through visiting parks and locations previously contaminated by other pets (Dado et al., 2012). Giardia can also be transmitted sexually with many studies reporting high prevalence among men who have sex with men (Escobedo et al., 2014)

Figure 7. The Giardia transmission cycle (CDC, 2012).

1.6.1. Foodborne transmission

Foodborne outbreaks are less frequently reported than waterborne outbreaks because this mode of transmission accounts comparatively, for only a small number of infections (Lujan & Svärd, 2011). Further, this may be due to the fact that foodborne outbreaks are much harder to trace and offending food or product may long be discarded or used up before the onset of clinical manifestation.

Only nine cases of foodborne outbreaks have been documented and food products implicated have been inadequately cooked or raw (Iwamoto et al., 2010) (Lujan & Svärd, 2011). The number of cases reported from all the nine foodborne outbreaks involved a total of 196 persons. Food types such as sandwiches were responsible for 44% (88/196). Other food products included noodle salad, home-canned salmon, trippe soup, ice, raw sliced vegetables herbs, fruits, and shellfish and sprouted seeds, leafy vegetables were responsible for 55% (108/196) (Porter et al., 1990; Robertson, 2013).

1.6.2. Waterborne outbreaks

The public health significance concerning Giardia waterborne outbreaks is that large populations are typically affected (Efstratiou et al., 2017; Escobedo et al., 2010; Fletcher et al., 2012). This is particularly important in certain areas in developing countries where large waterbodies such as lakes, rivers or reservoirs are contaminated and sanitation is poor. When such waterbodies are the main source of water supply to large communities, waterborne outbreaks are likely (Escobedo et al., 2010; Fletcher et al., 2012). Most Giardia waterborne outbreaks are said to be point-source after direct exposure to contaminated water, while secondary transmission is less common (Lujan & Svärd, 2011). Factors facilitating waterborne transmission are the fact that large numbers of cysts are excreted from infected persons into the environment, such cysts are environmentally stable and can survive for long periods of time in different waterbodies while maintaining infectivity (Craun et al., 2010; Feng & Xiao, 2011; Nygård et al., 2006). Up to 2 x 106 cysts per gram of faeces can be excreted in human faeces and 1.5 x 106 per gram from cattle (Geurden et al., 2010). Another factor facilitating waterborne outbreaks is the relatively low infectious dose for transmission of infection (Zmirou-Navier et al., 2006).

A comprehensive review on the impact of infectious diseases affecting humans from the period of WWI until 2003 reported that waterborne outbreaks with G. duodenalis were responsible for 41% (132/325) of such cases (Karanis et al., 2007). Of 132 outbreaks, 78% (103) were caused by consumption of contaminated drinking water (Karanis et al., 2007). Other outbreaks have been reported in the US, Norway and Finland (Table 4). In one of the earliest reported incidences, an estimated 50,000 people became infected in 1954 (Lujan & Svärd, 2011). Decades later, other outbreaks were reported during the late 1970s and 1980s. Since the year 2000, large outbreaks have been reported in Norway involving 48,000 people exposed to contaminated drinking water (Nygård et al., 2006; Robertson et al., 2006a). Since then, the vast majority of outbreaks have been reported in the US, Canada, UK, Germany and Sweden (Table 4).

Table 4. Distribution of documented waterborne outbreaks of giardiasis (modified from (Lujan and Svärd, 2011; Batt and Robinson, 2014)

Country No. of waterborne outbreaks Percentage

USA 82 77.4%

Canada 14 13.2%

UK 3 2.8%

Sweden 2 1.9%

Finland 1 0.9%

New Zealand 1 0.9%

Norway 1 0.9%

Germany 2 1.9%

1.6.3. Assemblages associated with waterborne and foodborne outbreaks

Assemblages A and B were reported in cases of waterborne giardiasis; however, assemblage A was more commonly identified (Prystajecky et al., 2015). Travel-related outbreaks have also been reported (Feng & Xiao, 2011; Prystajecky et al., 2015). Nursery outbreaks and sporadic cases have been reported in the UK, and North Wales (Appelbee et al., 2003). Small cases of foodborne outbreaks occurring in school children and members of a church who had food from the same restaurant in San Francisco also have been reported; however, assemblages were not specified (Sulaiman et al., 2003). Waterborne outbreaks have also been reported in 1,500 patients in Norway in which assemblage A and B were both responsible (Farid & Amir, 2015; Robertson et al., 2006b).

1.7. Giardia in environmental water and matrices

Different concentrations of Giardia cysts have been reported in drinking, recreational and raw water (Hashimoto et al., 2002; Richard et al., 2016). In water matrices in Europe, Asia, North and South America, occurrence rates of 30%-100% (cysts concentration; <0.01 to <100) have been reported in sampled water (Anceno et al., 2007; Schets et al., 2008). However, concentration is generally higher in developing countries; 12,780 cysts were present in one litre of surface water and 32,400 in a canal in Thailand (Anceno et al., 2007; Lim et al., 2008).

1.8. Pathogenesis and clinical manifestation

As previously described, trophozoites adhere to the brush border epithelium using their adhesive disk. This structure along with the flagella and various enzymes assist in colonization (House et al., 2011). While adhering to the enterocytes, the trophozoites cause alteration in sodium chloride iron excretion (Buret, 2007) (Di Genova & Tonelli, 2016).

The clinical symptoms of giardiasis may range from asymptomatic to symptomatic and from self- limiting to chronic disease. However, some hosts may be symptomatic with severe gastroenteritis and nutrient malabsorption (Fraser et al., 2000; Thompson et al., 2008b). The onset of signs and symptoms usually follows an incubation period of one week and may include abdominal cramps, bloating, foul-smelling greasy diarrheal stool, nausea and decreased appetite which may persist for several days or also much longer (Begaydorova et al., 2014; Kappagoda et al., 2011). Histological examination reveals trophozoites on mucosal surfaces or invading the brush border of epithelial cells (Yakoob et al., 2005). Infections that persist for weeks may result in lymphatic hyperplasia and bacterial proliferation. Extra-intestinal symptoms are rare; however, urticaria, reactive arthritis and retinitis were previously reported (Adam, 2001; Pietrzak et al., 2005). The severity of clinical manifestation is also said to be correlated with specific strains of G. duodenalis in humans. Assemblage B is associated with more severe cases of diarrhea and the severity of infection is also influenced by the immune, clinical and nutritional status of the host (Ignatius et al., 2012; Lujan & Svärd, 2011).

1.8.1. Giardia in children

Poor sanitation and poverty are typically associated with giardiasis in children especially in developing countries. Most cases of Giardia infections are reported from children between ages 1-4 and 5-9 years (Lujan & Svärd, 2011). In children, infections may also be asymptomatic or symptomatic. Chronic infections are associated with malnutrition, micronutrient deficiency, iron deficiency anaemia, poor cognitive function and failure to thrive (Bartelt & Sartor, 2015). However, a study in Peru followed 220 children and found that the status of infection did not correlate with weight or height for age (Hollm-Delgado et al., 2008).

1.8.2. Giardia in immunocompromised persons

Immunocompromised individuals include those infected with HIV, experiencing malignancy, undergoing chemotherapy, suffering from malnutrition or malabsorption or undergoing immunosuppressive therapy (Evering & Weiss, 2006). Such conditions decrease the host’s ability to effectively eliminate pathogens and are at increased risk of prolonged infections. Even though Giardia is not considered a major cause of enteritis in HIV-infected persons, the parasite is more likely to cause diarrheal disease in these persons (Stark et al., 2009). However, the clinical presentation is not different from giardiasis occurring in HIV-negative persons; occasionally HIV-infected persons may also be asymptomatic (Faubert, 2000).

1.9. Host immune response

As previously discussed, most Giardia infections are self-limiting and typically resolve in less than a week (Halliez & Buret, 2013). Production of secretory IgA antibodies is the primary mechanism by which the immune system maintains control of infections (Faubert, 2000; Solaymani-Mohammadi & Singer, 2010). Low levels of IgG have been reported and are especially associated with prolonged infections (El-Gebaly et al., 2012). Several antigens are responsible for stimulating immune response such as VSPs present on trophozoites and CWPs present on cysts (Lujan & Svärd, 2011). CD4 T cells are assisting in the production of IgA responses. Cytokines such as interferon (IFN) gamma, IL-2 (Interleukin-2), IL-5 and IL-6 produced by Th1 (T-helper-1) and Th2 are upregulated to induce the production of such antibodies against Giardia antigens (Bayraktar et al., 2005; Matowicka-Karna et al., 2009). One study correlated very high levels of INF-gamma, IL-4 and IL-5 with prolonged infections while elevated IL-8 were associated with infections of shorter duration (Long et al., 2010).

1.10. Target genes for molecular analysis

Several Giardia genomes have become available in the past years, including the ones of the WB and the GS strain (Franzen et al., 2009; Morrison et al., 2007). These are candidates primarily allowing the assessment of Giardia sequence variability and gene function. Multilocus sequencing approaches have been used to target a variety of Giardia genes, some of which have been used to establish molecular phylogeny and have been proposed to be informative for genotyping (Caccio & Ryan, 2008; Feng & Xiao, 2011; Lebbad et al., 2010). The most common gene targets are housekeeping genes such as the tpi and gdh genes, or for structural protein such as the bg, and ribosomal genes (Feng & Xiao, 2011). Other functional genes have also been used (Table 5).

Table 5. Giardia genes frequently used in molecular phylogenetical studies (Feng & Xiao, 2011; Lujan & Svärd, 2011)

Genes Function Species sequence available

MIh1 DNA repair G.duodenalis

Ef1α Translocational apparatus G.duodenalis, G.muris

Feredoxin Electron transfer G.duodenalis

Histone H2B Nucleosomal protein G.duodenalis

Histone H4 Nucleosome protein G.duodenalis

Alpha tubulin Structural protein G.duodenalis

Actin Structural protein G.duodenalis

ITS Non-coding ribosomal G.duodenalis

Chaperonin 60 Heat Shock Protein (HSP) G.duodenalis

ORF C4 Hypothetical (HSP) G.duodenalis

18S rDNA Ribosomal G.duodenalis, G.muris, G.agilis, G.microti, G.ardeae

ITS regions 5.8s Ribosomal G.duodenalis, G.muris, G.agilis, G.ardeae rDNA

Ribosomal protein Ribosomal G.duodenalis L7a

TPI Housekeeping G.duodenalis, G.muris, G.agilis, G.ardeae

GDH Housekeeping G.duodenalis, G.muris, G.agilis, G.ardeae

BG Structural protein G.duodenalis

1.11. Prevalence of giardiasis in humans

Giardiasis in humans is widespread; with infection rates of e.g. 1.4% in the US, 1.3% in the UK , 1.5 % in Germany, 2.5% in South Korea and 1.6% in Australia (Table 7) (Church et al., 2010; Huh et al., 2009; Read et al., 2002; Sagebiel et al., 2009). Most of these studies focused on asymptomatic children at day-cares and kindergarten, and subsets of symptomatic and asymptomatic adult patients presenting to hospitals or outpatient clinics. As described previously, infection rates are reported to be much higher in developing countries and can be as high as >30% (Table 6).

Table 6. Prevalence of giardiasis in developing countries

Country No. of samples tested Prevalence (%) Reference

Albania 125 17.6% (Berrilli et al., 2006)

Poland 232 1.3% (Solarczyk & Majewska, 2010)

Brazil 366 23.8% (Volotao et al., 2007)

Peru 845 23.8% (Perez Cordon et al., 2008)

Bangladesh 2,534 12.7% (Haque et al., 2005)

Nepal 1,096 4.1% (Singh et al., 2009)

Malaysia 321 23.7% (Mohammed Mahdy et al., 2009)

Thailand 204 23.7% (Traub et al., 2009)

Egypt 52 34.6% (Foronda et al., 2008)

Sahrawi 120 34.2% (Lalle et al., 2009) Republic

Saudi Arabia 1,500 6.5% (Adams et al., 2004)

Uganda 62 5% (Graczyk et al., 2002)

1.12. Prevalence of giardiasis in animals

The prevalence of giardiasis in domestic animals may be very high, especially in dogs and cats and there is strong evidence suggesting that these animals are responsible for cross transmission to humans (Feng & Xiao, 2011; Ryan & Caccio, 2013). In the US, infections were reported to be 19% and 11%, in symptomatic dogs and cats, respectively, based on detection of parasite antigen in stool (Carlin et al., 2006). In Europe, 24.78% of dogs and 20.3% of cats presenting with gastrointestinal symptoms were reported to be infected (Epe et al., 2010). Beavers were also implicated in zoonotic transmission of waterborne outbreaks in the US because hikers and campers reporting with giardiasis most often recall drinking water from streams or lakes (Erlandsen et al., 1990b; Feng & Xiao, 2011). Infectious Giardia cysts were also recovered from drinking water at treatment systems and storage reservoirs which suggest that contamination of water supplies may result from humans, domestic animals, and wildlife (Heitman et al., 2002).

The prevalence of giardiasis in farm animals such as cattle, sheep, goat and pig is also known (Feng & Xiao, 2011). Infections in cattle are well reported with rates up to 60 and 82% from 50 randomly selected Danish pigs and cattles, 38% in 4 week old calves in Germany and 49% in 6 month old calves in Norway (Hamnes et al., 2006; Jager et al., 2005; Langkjaer et al., 2007; Maddox-Hyttel et al., 2006). Giardiasis in sheep and goats have also been reported in many countries while infections in pigs are less frequently reported (Feng & Xiao, 2011). In the small Caribbean island of Trinidad and Tobago the prevalence of giardiasis in dogs was 25% in one study (Mark-Carew et al., 2013).

1.13. Giardiasis in Austria

Giardia infections are not well studied in Austria although giardiasis was reported as the most common parasitic infection among travellers returning from endemic areas (Reinthaler et al., 1998). Among patients in three sentinel clinics, infections with Giardia spp. were recorded at low prevalence (2.7%) (Reinthaler et al., 1998); however, there are no data on the genetic characterization of isolates from humans. Uncharacterized Giardia infections have been reported from calves and infections with assemblage A have been reported from cats (Haschek et al., 2006; Hinney et al., 2015). Pet ownership is high in many European countries with an average of 14.9% (range 7.2% to 35.0%) cat ownership and 12.0% (range 5.4% to 35.0%) dog ownership (Anyo et al., 2002). Further, in Germany, almost 50% of cases of giardiasis were suspected to be acquired indigenously (Espelage et al., 2010). Small outbreaks and occasionally complicated cases of giardiasis have been reported from rural areas and from the young and elderly

43 population in Austria, but molecular data on strains involved in human infections are largely lacking (Fortunat et al., 2004).

1.14. Giardiasis in Jamaica

Cases of human giardiasis have been reported in Jamaica and other Caribbean islands, but data from these regions are limited to surveys and screening studies. Asymptomatic and symptomatic giardiasis was reported at 57% and 43%, respectively, from 95 primary school children in Cuba (Pelayo et al., 2008). The prevalence among a hospital cross-sectional hospital population in Jamaica was 5.1% (17 of 328) using antigen and microscopic detection (Lindo et al., 1998). Further, a prospective community based study in Jamaica reported a prevalence of 6.3% (Rawlins et al., 1991). Therefore, Jamaica is a typical endemic region for giardiasis and infection rates in humans are assumed to be high (Lindo et al., 1998). Moreover, there also is a large population of stray dogs and cats in cities and towns, which might significantly contribute to contamination of the environment. There have been no studies of the parasite in dogs and cats in Jamaica and the zoonotic importance of the parasite has not been established.

Table 7. Prevalence of giardiasis in industrial countries.

Country No. of samples tested Prevalence (%) Reference

Germany 202 1.5% (Sagebiel et al., 2009)

Italy 1,989 0.4% (Crotti et al., 2005)

Netherlands 892 2% (Homan & Mank, 2001)

Portugal 190 3.7% (Almeida et al., 2006)

Australia 353 7.6% (Read et al., 2002)

New Zealand 66 7.6% (Learmonth et al., 2003)

1.15. Distribution of assemblages in humans and animals

Molecular analysis on giardiasis in humans and animals shows that assemblage A and B are generally distributed in developed and developing countries, such as Australia, Germany, Belgium, Italy, Malaysia, Thailand and Egypt (Table 8). However, assemblage A was the only genotype reported in Portugal and Brazil in adults (Table 8). In dogs and cats, infections are caused mainly by assemblage C and D (Lebbad et al., 2008; Traub et al., 2004). Occasionally infections occurring in cats are caused by assemblage F (Table 3). Despite the predilection of these genotypes to infect dogs and cats, infections with assemblage A and B are also frequently reported in these animals (Berrilli et al., 2004; Claerebout et al., 2009; Rimhanen-Finne et al., 2007; Ryan & Caccio, 2013). Host-specific assemblage E has also been reported in farm animals, primarily in cattle, sheep, goats and pigs (Table 3) (Appelbee et al., 2003; Armson et al., 2009; Geurden et al., 2008; Ryan & Caccio, 2013). Along with assemblage E, assemblage A is also regularly identified in these animals (Table 3). Such assemblages have also been reported in wild animals such as gorilla, chimpanzee, mandrill, moose, reindeer and coyote (Caccio & Ryan, 2008; Graczyk et al., 2002; Lebbad et al., 2010; Robertson et al., 2007a; Thompson et al., 2009).

Table 8. Prevalence of infection by assemblages

Country Assemblage/genotype Infection rates (%) Reference

Australia A&B 7.6 (Read et al., 2002)

Belgium A&B 4 (Sprong et al., 2009)

Germany A&B 1.5 (Sagebiel et al., 2009)

Italy A&B 0.4 (Crotti et al., 2005)

Netherlands A&B 2 (Van der Giessen et al., 2006)

Portugal A 3.7 (Robertson et al., 2007a)

New Zealand A&B 7.6 (Learmonth et al., 2003)

Albania A&B 17.6 (Berrilli et al., 2006)

Poland A&B 1.3 (Solarczyk et al., 2010)

Brazil A 23.8 (Volotao et al., 2007)

Peru A&B 20.4 (Cooper et al., 2007)

Bangladesh A&B 12.7 (Haque et al., 2005)

Nepal A&B 4.1 (Singh et al., 2009)

Malaysia A&B 23.7 (Mohammed Mahdy et al., 2009)

Thailand A&B 20.3 (Traub et al., 2009)

Egypt A&B 34.6 (Foronda et al., 2008)

1.16. Distribution of the A sub-assemblages

The genetic distribution of Giardia sub-assemblages in humans and animals differs according to geographic region (Table 9). Sub-assemblage AI and AII also differ in host specificity, sub- assemblage AII was more commonly reported in human infections while other animals were more commonly infected with sub-assemblage AI (Feng & Xiao, 2011; Thompson & Monis, 2004a). In other studies, sub-assemblage AI and AII are predominantly identified in humans and animals around the world (Table 9). Sub-assemblage AI has been predominantly isolated from humans in Italy and Portugal (Table 9), sub-assemblage AII has been isolated from patients in France, Peru and Australia (Table 9), and both assemblages were identified together in studies conducted in Brazil, Bangladesh and the Philippines. A recent study in the UK examined 35 human samples and 64% were Assemblage B and 27% were sub-assemblage AII (Amar et al., 2002). In Australia, infections with the genotypes from assemblage B were also more prevalent (70%) in humans and dogs than with those of assemblage A (sub-assemblage AII) (30%) (Thompson et al., 2008b; Traub et al., 2004). Despite the mode of transmission or origin of specific assemblages, person-person transfer in localized endemic communities or institutional settings is common.

Table 9. Distribution of the A sub-assemblages in humans and animals and the respective gene targets used

Country Host Subtype Gene Reference

France Human AII tpi, bg (Bertrand et al., 2005)

Italy Human AI bg (Marangi et al., 2010)

Portugal Human AI bg (Sousa et al., 2006)

UK Human AII tpi (Amar et al., 2002)

Brazil Human AI & AII bg (Volotao et al., 2007)

Peru Human AII tpi (Cooper et al., 2010)

Bangladesh Human AI &AII tpi (Almerie et al., 2008)

Philippines Human AI &AII tpi (Yason & Rivera, 2007)

Australia Human AII gdh (Yang et al., 2010)

Germany Dog AI gdh (Leonhard et al., 2007)

Italy Dog AI bg (Marangi et al., 2010)

Mexico Dog AI & AII bg (Lalle et al., 2005b)

Brazil Dog AI bg (Volotao et al., 2007)

US Cat AI gdh (Vasilopulos et al., 2007)

Brazil Cat AI bg (Volotao et al., 2007)

Japan Cat AI tpi, bg (Suzuki et al., 2011)

Norway Moose AI bg, gdh (Robertson et al., 2007b)

US Herring AI tpi, gdh (Lasek-Nesselquist et al., 2008)

1.17. Zoonotic potential

Giardia infections in companion animals such as dogs and cats are usually asymptomatic and are potentially transmissible to humans (Caccio & Ryan, 2008; Thompson & Monis, 2004a; Thompson et al., 2008b). Recent studies have reported different assemblages to have different zoonotic potentials (Caccio et al., 2005; Palmer et al., 2008; Thompson & Monis, 2004a; Thompson et al., 2008b). Zoonotic assemblage A is frequently found in livestock and companion animals such as dogs, cat, and horses (Feng & Xiao, 2011). Assemblage B is also frequently reported but is less commonly reported in dogs (Feng & Xiao, 2011). Mixed infections also occur with assemblage B which suggests that the infection could have been passed on from dogs to humans and vice versa (Thompson & Monis, 2004a).

The same assemblages and sub-assemblages have been implicated repeatedly in cases of giardiasis in dogs, cats, humans and other animals. Most interests in zoonotic transmission focus of assemblage A because this is less host-specific than all other zoonotic assemblages (Thompson & Monis, 2004a). Sub-assemblage AI has been isolated from dogs in Germany, Italy, Mexico and Brazil (Table 9). The same subtype has also been identified in cats in the US, Brazil and Japan (Table 9). Other animals such as moose and herrings have also been found to be infected (Table 9). While assemblage B is associated with prolonged and severe cases of diarrhoea and is considered more pathogenic, it is less suspected to be zoonotic (Feng & Xiao, 2011). The general consensus is that sub-assemblage AI and AII are associated with zoonotic infections in humans, while assemblage B predominantly infects humans (Plutzer et al., 2010; Wang et al., 2014). Although reported less often, human infections with assemblages C, D, E and F have occurred, providing evidence of zoonotic transmission (Ryan & Caccio, 2013).

1.18. Treatment of giardiasis

Several chemotherapeutic compounds belonging to the nitroimidazole group such as metronidazole, quinacrine, tinidazole and furazolide are useful in treating human giardiasis (Gardner & Hill, 2001; Lujan & Svärd, 2011). Nevertheless, resistance to metronidazole and furazolidine have been described in vitro (Leitsch, 2015). Metronidazole is also being used in veterinary medicine (Lujan & Svärd, 2011). Other drugs such as the benzimidazoles also remain a treatment option for giardiasis in companion animals. A pyrantel-febantel-praziquantel combination has been used in cats and dogs and has been found to be more efficacious than the standard treatment; significantly reducing the number of cysts (Bowman et al., 2009; Montoya et al., 2008; Payne et al., 2002; Scorza et al., 2006).

1.19. Vaccination against Giardia

As previously described, secretory IgA is produced in the lamina propria of the enterocytes and the antigen-specific CD4 plus Th cells response provides the most important immunological protection against Giardia (Abdul-Wahid & Faubert, 2008; Faubert, 2000; Roxstrom-Lindquist et al., 2006). Although mucosal antibodies are produced during active infection, only 1% of infections stimulate polyclonal antibodies. For that reason, antibodies directed against epitopes of the parasite play a significant role in elimination (Lujan & Svärd, 2011). As such, the development of effective vaccines requires characterization of epitopes that are effective in stimulating protective antibodies (Lujan & Svärd, 2011). The challenge that exists for the production of an anti-Giardia vaccine is that systemic administration may be inappropriate because Giardia infections are typically localized to the mucosa. As a result, local intestinal antibodies are required to successfully eliminate the parasite. Candidate antigens for a vaccine against Giardia have been derived through extensive study of the Giardia WB strain. These include antigens against HSPs, lectins, gardins, tubulins and VSPs (Lujan & Svärd, 2011). Induction of immunity against Giardia has been successful experimentally. Protective immune responses have been initiated by DNA vaccines introduced at the intestinal mucosa (Lujan & Svärd, 2011; Weiss, 2003). Other experimental vaccines involved administration of plasmids, micro-encapsulated DNA, and bactofection vehicles such as Salmonella typhimurium (Weiss, 2003).

GiardiaVax is a vaccine established for dogs experiencing chronic symptomatic infection. The vaccine which has been first licensed in Canada consists of chemically inactivated trophozoites which have been demonstrated to significantly reduce shedding of Giardia cysts in young pups and dogs with symptomatic giardiasis when therapy has been unsuccessful (Anderson et al., 2004). Despite being highly effective at reducing the quantity of cysts in infected animals, GiardiaVax does not provide strain-specific protection and prevent the recurrence of infections. Because the vaccine was also licensed as an adjunct aid in treatment, it is helpful in reducing the environment contamination during outbreaks, but provides no benefit when administered to infection free dogs. As of 2006, the Giardia vaccine has been listed by the American Animal Hospital Association as “not recommended” (Kahn & Line, 2007).

1.20. Laboratory diagnosis of infections

There are several microscopic, antigen detection and molecular methods available for identifying Giardia infections in humans and animals (Gotfred-Rasmussen et al., 2016). Some routine diagnostic laboratories will use microscopic methods while others may use a combination of antigen or molecular detection. Antigen detection is also available to identify antigens expressed

50 on the surfaces of trophozoites and cysts in dogs and cats. The most common antigen detection test that is used to diagnose infections in companion animals is the IDEXX SNAP Giardia rapid test (Rishniw et al., 2010). Molecular methods are well developed and are commonly used, not only in routine diagnostics, but also in molecular epidemiological studies (Thompson & Ash, 2016).

1.20.1. Microscopy

Detection of Giardia trophozoites and cysts by microscopy can be tedious. Trophozoites are usually present only in diarrheal stool while cysts may be shed intermittently in formed stools and it is generally recommended that three consecutive stool samples are examined by (Anderson et al., 2004; McAllister et al., 2005). For microscopy the most commonly used methods are direct smear, formalin ether concentration, with or without subsequent staining, e.g. with trichrome (McHardy et al., 2014).The advantage of using microscopy is that it is simple, and that eventually other parasites (other protozoa cysts or helminth eggs and larvae) may be detected at the same time. Further, microscopy will allow for the discovery of new infectious causes of diseases. Although, microscopy remains the mainstay of diagnosis in many developing countries, it is time-consuming and thus increasingly replaced by molecular techniques in high-income countries.

1.20.2. Direct smear

For diarrheal stool samples, a direct smear may be performed to detect motile trophozoites containing two nuclei and four pairs of flagella on the dorsal surface. Despite being the most reliable method for diagnostic purposes, trophozoites are fragile and may die quickly after passage if conditions in the environment are not feasible for survival. For the method to be effective stools must be examined within 30 minutes of being passed by the patient which is very rarely possible (Cama & Mathison, 2015) .

1.20.3. Concentration techniques

Formalin-ether concentration is the most widely used microscopic method in the diagnosis of intestinal parasites (Cama & Mathison, 2015). The method was developed in 1948 by Ritchie and has undergone several modifications. Essentially, stool samples are subjected to centrifugation after they are homogenised in formalin and fat emulsified using an organic solvent such as diethyl-ether or ethyl acetate (Becker et al., 2011). The parasite stages are trapped in the sediment after centrifugation and this is examined after staining using iodine (Becker et al., 2011).

Flotation techniques are also used for diagnostic and epidemiologic investigations. The aim of this technique is to float parasite stages on the surface of a high specific gravity solution such as table sugar, zinc sulfate (ZnSO4) or sodium chloride ((Becker et al., 2011). Sheather’s and ZnSO4 flotation techniques are the most commonly used flotation methods used to detect intestinal parasites in faecal samples of animals (Dryden et al., 2005b). Sheather’s solution has a specific gravity of 1.27 to 1.33 which is considered the Gold Standard for flotation of parasitic products (Dryden et al., 2005b). With this technique, all eggs and protozoan cysts and oocysts are expected to float while maintaining their morphologic structure while the fecal debris sinks.

1.20.4. Enzyme-Linked Immunosorbent Assay

Various ELISA kits detecting Giardia surface antigen in stool are commercially available and they are usually highly sensitive and also adequately specific. They can be used for routine diagnosis, but also to screen large batches of samples. This is especially useful when microscopic detection fails due to low parasite densities (Lujan & Svärd, 2011). Immunochromatographic assays are also commercially available and are usually more rapid than ELISA (Abdel Hameed et al., 2008) (Johnston et al., 2003).

1.20.5. Immunofluorescence assay

The Giardia IFA uses fluorescent-labeled antibodies specific for Giardia cyst antigens. Detection antibodies conjugated to fluorescein isothiocyanate under blue filter block microscope fluoresces green when antibodies are bound to antigenic epitope (Gotfred-Rasmussen et al., 2016). Indirect IFA is available and is widely used and is especially useful in detecting Giardia cysts in food and environmental samples. Instead, two antibodies are used for detection and is more time consuming as a result (Keserue et al., 2011).

1.20.6. Nucleic acid detection

Nucleic acid detection methods are more sensitive than microscopic and immunological approaches for detecting Giardia infections (Tavares et al., 2011). Giardia DNA can be extracted also from a small number of cysts. However, there is no standardized approach (Hawash, 2014). Many genes have been targeted for molecular analysis (Table 10) (Feng & Xiao, 2011; Minetti et al., 2015). A multilocus approach targeting 2 or more gene fragments is recommended for characterizing assemblages A and B (Zheng et al., 2014). Molecular analysis includes conventional PCR targeting short gene fragments such as SSU rDNA for detection of the parasite (Table 10). Traditionally, RFLP (Restriction Fragment Length Polymorphism) has been used. However, nested and RT-PCR (Reverse Transcription-Polymerase Chain Reaction) is

52 increasingly being used to target longer gene segments such as the tpi, gdh and bg which are more reliable targets for identification, particularly at the sub-assemblage level. Multiplex and RT-PCR also offer the possibility to detect different species in the same reaction (Stark et al., 2011). Mixed infections can be problematic in endemic settings and especially when only one sample is analysed. RT-PCR is highly sensitive; however, the quantitative nature of such a method is invariable in estimating the parasitic DNA load (Lujan & Svärd, 2011; Stark et al., 2011). Despite well-developed molecular methods to identify different species of Giardia, molecular detection methods are difficult to implement in molecular diagnostic labs because to date, standardized methods do not exist (Stark et al., 2011). Nonetheless, some multiplex RT- PCR methods have been implemented for simultaneous detection Giardia and other protozoan parasite which demonstrated high specificity (Stark et al., 2011).

Table 10. Gene target for molecular analysis and type of assay established

Gene Assay (PCR) Characterization level

TPI Conventional, nested, sequencing, RFLP, RT, microarray Genotype, subtype

GDH Conventional, nested, sequencing, RFLP Genotype, subtype

BG Conventional, nested, sequencing, RFLP Genotype, subtype

Ef1α Conventional, Nested, sequencing Genotype, subtype

SSU-rDNA Conventional , Nested, microarray Genotype

GLORF-C4 Conventional, sequencing, RFLP Genotype

IGS Conventional, nested, sequencing Genotype

1.21. DNA extraction and PCR amplification of Giardia cysts fixed in formalin

Intestinal protozoans have emerged with important public health concerns causing widespread infections in humans and animals. Microscopic analysis guided by morphological characteristics of parasites remains the gold standard for diagnosing such infections, e.g. caused by representatives of the genera Giardia, Entamoeba or Cryptosporidium. Formalin-fixed samples can be stored for long periods of time (Dietrich et al., 2013). However, because the morphological characteristics of parasites provide no information at the species level, microscopy alone is of minimal use for epidemiological studies (Libman et al., 2008; McHardy et al., 2014).

Formalin fixation has been in use for more than 100 years to preserve biological materials and other specimens (Dietrich et al., 2013). Molecular biological studies on Giardia isolates from humans and animals are providing useful information on genotypic distribution. This allows for better understanding of the epidemiology, the predominant infection routes, and the evolutionary and phylogenetic relationships. Like in other biological materials, preservation of Giardia cysts in formalin leads to modification, degradation and cross-linking of DNA which makes it difficult to extract DNA for molecular analyses (Gilbert et al., 2007). Targets used in molecular analysis are susceptible to mechanical stress due to DNA cross-linking, as a result, their access to enzymes such as polymerase used for molecular detection is decreased (Dietrich et al., 2013). Despite cross-linking and destruction DNA, modified protocols have successfully extracted DNA from formalin-fixed samples (Tang et al., 2009). Nevertheless, successful amplification remains difficult and obtaining high quality DNA for molecular analysis remains a challenge. To optimize the yield, studies have focused primarily on extraction protocols and optimized PCR amplification methods. Such treatments prior to extraction include pre-heating, washing, treatment with proteinase K and EDTA (Masuda et al., 1999; Shi et al., 2002). Important variables to consider when optimizing PCR protocols include the annealing temperature, Mg2+ concentrations and buffer and cycling conditions (Lorenz, 2012). Independent variables such a PCR inhibitors presenting as ionic detergents should also be taken into account (Hoppe et al., 1992; Weyant et al., 1990). To further increase the likelihood of successful amplification, nested PCR is effective at reducing spurious products by including two sets of primer pairs (Lorenz, 2012).

1.22. AIMS

1.22.1. Publication I

Multilocus sequence analysis of Giardia spp. isolated from patients with diarrhea in Austria.

The first aim of this dissertation was to use molecular tools to characterize samples identified from humans in Austria to compare them with isolates from Jamaica. Investigating and comparing a region with assumed low endemicity (Austria) with a region with high endemicity (Jamaica) will not only for the first time delivers molecular data on strains/ genotypes involved in human infections in these regions, but also provide data which is of molecular epidemiological significance. This focuses on characterization of sub-assemblages (AI, AII, AIII, AIV, BI, BII, BIII and BIV) involved in human infections in both populations to better understand the transmission dynamics.

1.22.2. Publication II

Molecular epidemiology and multilocus sequence analysis of potentially zoonotic Giardia spp. from humans and dogs in Jamaica

The second aim was to identify and compare the host distribution of Giardia assemblages and sub-assemblages from humans and dogs in Jamaica. While there have been studies on the prevalence of Giardia assemblages in humans and animals from several countries, there have been no reports from Jamaica. Jamaica presents an ideal study site because of the absence of enforcement of animal control laws and a large population of stray dogs which increase the potential for zoonotic transmission. Also, the prevalence of giardiasis in developing countries is known to be significantly higher than in industrialized countries. Identifying and comparing the distribution of Giardia assemblages among hosts will lead to an understanding of the level of zoonotic and anthroponotic transmission in this geographical region.

1.22.3. Publication III

Successful extraction and PCR amplification of G. duodenalis DNA from formalin fixed stool.

The third aim was to assess whether sample pre-treatment, and modified DNA extraction protocols followed by PCR amplification were useful in identifying Giardia isolates preserved in

55 formalin at the assemblage and sub-assemblage level, respectively. Such protocol focused primarily on pre-treatment in EDTA, proteinase K digestion and a PCR using commonly targeted genes to see whether such samples could be useful in future molecular characterization studies.

CHAPTER TWO: MATERIALS AND METHODS

2.1. Ethics statement

The study protocol was approved by the University Hospital of the West Indies/University of the West Indies / Faculty of Medical Sciences (UHWI/UWI/FMS) Ethics Committee and the Medical University of Vienna (Ethics permit number: 1070/2015). It was conducted according to NIH, guidelines, ISO15189 standard for medical laboratories and the Austrian Gene Technology Law.

2.2. Safety

Safety regulations using biological and chemically hazardous material by international standards, the Austrian Gene Technology Law and the ISO 15189 standard for medical laboratories were strictly followed.

2.3. Sample collection

Fecal samples collected from humans and dogs (285 and 225, respectively) were obtained from the National Public Health Laboratory (NPHL) in Kingston, the diagnostic laboratory of Department of Microbiology, UHWI and 20 veterinary practices across Jamaica. Samples were collected from symptomatic patients presenting at the UHWI and the NPHL. Samples collected for each veterinary practice included dogs presenting with or without evidence of gastrointestinal disturbances. Only human samples were collected from Austria. Isolated Giardia cysts from Austrian patients were stored at the parasitology laboratory at the Institute of Specific Prophylaxis and Tropical Medicine (ISPTM), Medical University of Vienna (MedUni Vienna). Stool samples from these patients were processed using microscopy and positive samples were stored for molecular analysis. A total of 65 Giardia positive samples were stored at the institute in 85% alcohol. Samples were stored Samples were stored in a sodium acetate and ethanol solution until use. Moreover, a set of old, formalin-fixed Giardia-positive stool samples from the sample collection at the ISPTM were included in the study

2.4. Microscopic analysis

Identification of Giardia trophozoites from fresh diarrheal stool collected from the Jamaican patient population was done using the direct smear method. Formalin or ethanol ether concentration technique (F/EECT) was used to identify Giardia cysts in all formed human stool samples (Becker et al., 2011; Won et al., 2015). ZnSO4 flotation (specific gravity; 1.2-1.25) was done on stool samples collected from dogs (Dryden et al., 2005a). Giardia trophozoites and cysts were identified based on their morphological characteristics and comparison with reference

58 specimens. Due to suspected cases of low parasite density in Jamaica, all samples negative on microscopic analysis were also concentrated and stored for DNA extraction.

2.5. Molecular investigation

2.5.1. DNA extraction

A total of 510 fecal specimens collected from humans and dogs in Jamaica were included in the study and stored in 85% ethanol until DNA extraction was performed. From the Austrian study cohort, sixty five Giardia-positive samples were selected for molecular investigation. Subsamples (~ 3g) of each were concentrated and used for molecular analysis. DNA was extracted by adding 20 mg/200 µl of fecal specimen in a 1.5ml tube. Stool samples were added to 1000 ml stool lysis buffer, heated at 70ºC for 10 minutes and vortexed for 3 minutes. The protocol outlined in QIAamp Fast DNA Stool Kit® (Qiagen, Hilden, Germany) was closely followed for completion of extraction. The final extracted products were stored at -20ºC until PCR was performed.

2.5.2. Conventional PCR

All 510 samples collected from humans and dogs in Jamaica were subjected to screening using conventional PCR by amplifying small fragments of the SSU rDNA. This was done using primers

RH11 and RH 4 (Table 11). PCR was run in 50 µl reactions consisting of molecular grade H2O,

25mM MgCl2, HOT FIREPol® (5U/µl), 10 x Buffer B20 mM dNTP mix, and 10 pmol/µl of forward and reverse primer dilutions (Solis BioDyne®, Estonia). Cycle conditions for amplification were as follow; 95ºC for 15 min, 35 cycles (95ºC for 1 min; 52ºC for 2 min and 72ºC for 2 min and a final extension 72ºC for 7 min). PCR products were stained and visualized using gel electrophoresis with a GelRed™ stain (BioTrend, Cologne, Germany).

2.5.3. Nested PCR

After first molecular screening, nested PCR was done on all samples, including also the 65 Giardia-positive samples from the Austrian patient population. Nested PCR targeted three loci (tpi , gdh, and bg) (Table 11). These were used as genetic markers for genotyping and subtyping as previously described (Feng & Xiao, 2011; Gelanew et al., 2007). Cycle conditions for both, primary and nested amplifications, were identical: 35 cycles (94ºC for 1 min, 45ºC for 30 s and 72ºC for 45 sec). Primary primer pairs amplify external products which were subsequently used as templates in nested reactions. Species-specific primers were used for analysis of mixed infections with assemblage A and B targeting the tpi gene (Anuar et al., 2014) among Austrian patients. All reactions were done using an Eppendorf® thermocycler with an initial preheating at

95ºC for 10 min and a final extension at 72ºC for 7 min. The final PCR amplicons were stained with GelRed™ stain (BioTrend, Cologne, Germany) using 2% agarose for analysis under a UV transilluminator box after gel electrophoresis.

Table 11. Primers used in identification of Giardia at the tpi, gdh and bg loci (Feng & Xiao, 2011; Zheng et al., 2014)

Primer Oligonucleotide 5’-3’ Amplicon Assay

TPI1 For ( 5’-AAC GCA ATC ACT GTA TCT-3’) 650bp Nested

TPI2 Rev (5’-CAA TGA CAA CCT CCT TCC-3’)

TPI3 For (5’-CTT CAT CGG CGG TAA CTT-3’) 334bp

TPI4 Rev (5’-GGC ACG CTT AGC CTT CTT-3’)

GDH1 For (5’-TTC CGT GTT CAG TAC AAC TC-3’) 754bp Nested

GDH2 Rev (5’-ACC TCG TCC TGA GTG GCG CA-3’)

GDH3 For (5’-ATG ACT GAG CTT CAG AGG CAC GT-3’) 530bp

GDH4 Rev (5’-GTG GCG CAA GGC ATG ATG CA-3’)

G7 For (5’-AGG CCC GAC CTC ACC CGC AGT GC-3’) 753bp Nested

G759 Rev(5’-GAG GCC GCC CTG GAT CTT CGA GAC GAC-3’)

GiarF For(5’-GAA CGA ACG AGA TCG AGG TCC G-3’) 511bp

GiarR Rev (5’-CTC GAC GAG CTT CGT GTT-3’)

AssAF For (5′-CGC CGT ACA CCT GTC-3′) 332bp Conventional

AssAR Rev (5′-AGC AAT GAC AAC CTC CTT CC-3′)

AssBF For (5'-GTT GTT GTT GCT CCC TCC TTT-3') 500bp Conventional

AssBR Rev (5'-CCG GCT CAT AGG CAA TTA CA-3')

RH11 For(5’-ATC TTC GAG AGG ATG CTT GAG-3’) 150 Conventional

RH4 Rev (5’-AGT ACG CGA CGC TGG GAT ACT-3’)

2.5.4. Sequencing PCR

Positive amplicons were cut from the respective gels and purified using the illustra GFX PCR DNA and Gel Band Purification Kit® (illustra GFX, GE Healthcare, Austria). If sequencing PCR was delayed, PCR products were purified, eluted and then stored at -20ºC until sequencing PCR was performed. Purified products were sequenced using the ABI BigDye® Terminator V3.1 sequencing kit and an ABI automated sequencer (both Applied Biosystems, Vienna, Austria). For comparison, published Giardia sequences were obtained from GenBank®. Multiple alignments of nucleotide sequences were conducted using the CLUSTAL W® alignment program and analyzed using GeneDoc (version 2.7.000). Genotypes and subtypes were identified according to (Feng & Xiao, 2011; Thompson, 2000).

2.6. Preparation of samples stored in formalin

Forty five old, formalin-fixed (1.5% formalin) Giardia-positive samples from the sample collection at the parasitology laboratory at the ISPTM, MedUni Vienna, Austria were included in the study and analysed by molecular methods during 2013-2015. After microscopic re-analysis to assess the overall condition of the samples, they were subjected to pre-treatment, DNA extraction and PCR amplification. Positive controls included known Giardia positive samples stored in 85% ethanol. The effectiveness of the molecular analysis on formalin fixed samples was subsequently tested in nested PCR experiments.

2.6.1. Pre-treatment and extraction protocols

Prior to DNA extraction, all samples were subjected to pre-treatment:

1) Washing 500 µl of sample in 400 µl of 100% alcohol

2) Centrifugation at 16,000x g for 10 minutes

3) Discarding supernatant to remove formalin residue

4) Air-drying to remove alcohol residue

5) Washing in 400 µl of 70% alcohol

6) Centrifugation for 16,000x g for 10 minutes

7) Discarding supernatant followed by air-drying

8) Adding 500 µl of 0.5M EDTA to sample residue

9) Incubation for 1 hrs at 55°C

10) Adding 100 µl of proteinase K to the mix

11) Incubation for 2hr at 55°C

12) Discarding supernatant

13) Adding 1000 µl of stool lysis buffer

14) Incubation at 85°C for 10 minutes in a thermomixer with mechanical shaking at 350rpm

15) Homogenization by mechanical shaking in a PowerBead Tube at 5000rpm for 20s.

Extraction procedure was done following the instructions of the QIAamp Fast DNA Stool Kit (Qiagen, Hilden, Germany. After extraction, eluted DNA from each samples were stored at -20ºC until PCR was performed.

2.6.2. NanoDrop® spectrophotometry

After DNA extraction, the DNA concentrations of every eluate were measured using a NanoDrop™ (ThermoScientific, Vienna, Austria).

2.6.3. PCR protocol for formalin-fixed Giardia samples

After DNA extraction, nested PCR was done targeting the tpi, bg and gdh loci (Gelanew et al., 2007; Zheng et al., 2014). Conditions for primary and nested PCR included: 35 cycles (94ºC for 10 min, 45ºC for 30 sec and 72ºC for 45 sec) including an initial preheating at 95ºC for 10 min and a final extension at 72ºC for 7 min. PCR reactions (50 µl) included 10 µl of master mix reaction containing molecular grade water, 25mM MgCl2, 20mM dNTP mix, HOT FIREPol® (5U),5 µl 10 x Buffer B and 10 pmol/µl of forward and reverse primer dilution (Solis BioDyne, Estonia). End products were visualized using gel electrophoresis.

1. The same PCR protocol was repeated on samples that had given no amplicons after PCR, despite being microscopically positive. Eluates of samples were increased to 6 µl and 9 µl and PCR repeated under the same conditions (Figure 8). 2. If amplification remained unsuccessful after the first step, PCR was repeated using 4-fold concentration of polymerase to ensure that the carry-over of proteinase K was not responsible for false negative (Figure 8).

Positive amplicons were purified using the Gel Band Purification kit (illustra GFX, GE Healthcare, Austria) and directly sequenced (BigDye Termin ator V3.1, Applied Biosystems, Austria).

Figure 8. Workflow of Giardia-positive formalin fixed samples

CHAPTER THREE: RESULTS

3.1. Microscopic and molecular screening of samples in Jamaica

By microscopy, 1.6% (8/510) of samples collected from humans and dogs in Jamaica were positive on investigation. FECT conducted on all human samples detected only two samples positive with cysts from 285 (0.7%). Of 225 dog samples, six (2.7%) were positive in the ZnSO4 flotation technique.

Molecular screening by conventional PCR of all 510 samples targeting the SSU rDNA locus yielded amplicons of the appropriate size in sixty three samples. Giardia positive samples were identified at a rate of 6.7% (19/285) in humans and notably higher, 19.5% (44/225) in dogs based on conventional PCR.

3.2. Genotype and subtype distribution of Giardia in Jamaica

Multilocus sequence analysis by nested PCR targeting three genetic loci showed that the tpi gene is highly sensitive for molecular detection of Giardia genotypes with 100% (63/63) positive on amplification. A slightly lower detection rate was seen at the bg locus, with 70% (44/63) while 55.6% (35/63) were positive at the gdh locus. Sequencing PCR on all 63 positive samples only identified Giardia assemblage A from both, humans and dogs, in Jamaica. Subtyping analysis revealed that sub-assemblage AII was most prevalent in both, humans and dogs, accounting for 84.2% (16/19) and 70.5% (31/44), respectively. Sub-assemblage AI was responsible for fewer cases of giardiasis in both populations at 15.8% (3/19) in humans and 29.5% (13/44) in dogs, respectively. Four samples gave inconsistent results when results from the SSU rDNA were compared at the sub-assemblage level. Sub-assemblage AI (three samples) and assemblage B (one sample) were identified. However, when these samples were compared at all other loci, sub-assemblage AII was consistently identified while AI was identified a single time at the bg locus. The results provided show that sub-assemblage AII was most prevalent among humans and dogs with a clear evidence of genetic polymorphism within the tpi gene.

A B

600BP 600BP 450BP 450BP 350BP 350BP 250BP 250BP 150BP 150BP

50BP 50BP PC 1 2 3 4 5 NC PC PC 1 2 3 4 NC

Figure 9. Nested PCR at the tpi (A) and bg (B) genetic loci

PC: positive control, NC: negative control

3.3. Giardia genotype and subtype distribution in Austria

Of 65 Giardia positive samples on initial microscopic analysis, DNA was successfully extracted from 52 which was followed by nested PCR amplification at the tpi, bg (Figure 9) and gdh loci. Assemblage B accounted for most cases of giardiasis at 65.4% (34/52). Assemblage A identified in fewer cases of human giardiasis at 34.6% (18/52). Analysis using species-specific primers for assemblage A and B showed no evidence of mixed infections. Subtyping analysis showed a comparably high level of species diversity whereas sub-assemblage BIV was responsible for most cases 46.2% (24/52). Sub-assemblage AII was identified at a slightly lower rate of 25% (13/52), followed by BIII and AI in for 19.2% (10/52) and 9.6% (5/52) of patients, respectively. The tpi gene was successfully amplified 100% of all samples. However, amplification rates at the bg and gdh loci were slightly lower; 86.5%; 45/52 and 55.8%; 29/52, respectively. Genotype inconsistencies were observed at the bg locus in up to 44% (15/34) of samples which were non- concordant to other genetic loci. Seven samples were identified as sub-assemblage BIII at the tpi but were subsequently identified as sub-assemblage BIV at the bg and gdh loci. In similar analyses, sub-assemblage AI was identified at the gdh when results from three samples were analysed but were identified as sub-assemblage AII when the tpi locus was used.

3.4. Molecular analysis on formalin-fixed samples

DNA quantification from formalin fixed samples after NanoDrop® spectrophotometry showed that samples fixed in formalin generally had a lower concentration of DNA (mean 5.96 ng/ul) compared to non-formalin fixed samples. Positive controls were fixed in ethanol and generally contained twice the DNA concentration (mean 10.44 ng/ul).

Standard PCR conducted on all 45 samples showed a 62.2 %( 28/45) amplification yield when the tpi gene was used. A lower amplification yield was seen at the bg and gdh loci with amplification rates of 33.3% (15/45) and 18.0% (8/45), respectively. Ethanol-fixed Giardia samples used as positive controls showed successful amplification at all three genetic loci. To assess whether low DNA concentration contributed to unsuccessful amplifications, PCR was repeated on all samples that were unsuccessfully amplified using higher concentrations of DNA which however, resulted in no difference in amplification yield.

3.5. Modified PCR: 4-fold concentration of DNA polymerase

A 4-fold concentration of DNA polymerase was included in the master mix to reduce PCR inhibition of longer fragments, e.g. the bg and gdh genes. PCR repeated on samples that were unsuccessfully amplified with 4-fold polymerase resulted in two additional positives, one at the tpi and the other at the bg locus. Therefore, pre-treatment, modified DNA extraction along with modified PCR amplification shows slightly higher rates of amplification.

3.6. Correlation between amplification of short fragments and storage

There was a correlation between the length of storage and amplification of fragments of 334bp at the tpi versus the bg and gdh genes which were both longer than 500 bp. Samples fixed for more than 10 years were all positive at the tpi locus. Those that were stored for longer showed up to 46.7% (7/15) positive. None of the samples stored for more than years were positive at the bg or gdh loci. However, only a few samples that were stored for shorter duration were positive at these two loci.

3.7. DNA quality and analysis of nucleotide sequences

In a genotyping experiment, analysis of sequences obtained showed that also with formalin-fixed samples it was possible to differentiate Giardia genotypes at the assemblage and sub- assemblage level. Despite evidence of sequencing artifacts, 50% (9/18) of samples produced high quality readable sequences at the bg locus and identified assemblage B; subtype BIV. In

68 addition to the high rates of amplification seen at the tpi locus, high quality sequences were also produced for this locus, in up to 41.2% (12/29). Such readable sequences identified assemblage A and B; one sub-assemblage AII, ten sub-assemblages BIII and one BIV. The gdh locus produced mostly sequences of poor quality. However, 22.2% (2/9) were able to differentiate Giardia at the species level and identified sub-assemblage BIII.

CHAPTER FOUR: DISCUSSION

4.1. Publication I and II

The overall prevalence of giardiasis on microscopic analysis was 1.6% in humans and dog in Jamaica. Infection rates were higher in asymptomatic dogs than in symptomatic humans (2.7% and 0.7%, respectively). Molecular analyses using conventional PCR showed a higher prevalence of 6.7% in humans and 19.5% in dogs. Multilocus sequence analysis at the tpi, bg and gdh loci showed that assemblage A was the only genotype found in humans and dogs in Jamaica.

Low detection rates in microscopic analysis were not unexpected due to known low sensitivity of the microscopy associated with intermittent shedding of Giardia cysts; shedding of fewer cysts in asymptomatic cases and distortion of cysts during flotation due to changes in osmotic pressure (Zajac et al., 2002). The findings from this study were comparable to results obtained from a study previously at the outpatient clinic in Jamaica UHWI which reported a prevalence of 1.8% using microscopy and 5.2% using antigen detection (Lindo et al., 1998). A decline in intestinal parasitic infection has been reported by Rawlins (1982), which was attributed to improved sanitation and living standards. However, in a prospective community based study the prevalence of human giardiasis in Jamaica was estimated to be 6.3% (Rawlins et al., 1991). Molecular investigation showed a significantly higher prevalence and is reflective of previous estimates based on antigen detection (Lindo et al., 1998). Findings from similar studies reported infection rates to be twice as high when comparing microscopic and molecular detection techniques (Scaramozzino et al., 2009).

Comparison of findings from molecular analysis from this study to developed countries showed similar prevalence rates. Infection rates in humans were lowest in asymptomatic children in the day-care in the UK (1.3%), symptomatic and asymptomatic patients from the Rocky Mountain region is the US (6.4%%), five kindergartens in Germany (1.5%), and among community based asymptomatic individuals in Australia (1.6%) (Church et al., 2010; Davies et al., 2009; Huh et al., 2009; Read et al., 2002; Sagebiel et al., 2009). In developing countries, the parasite accounts for between 8 and 35% of infections in children 2-5 years old in South and Far East Asia, 120 children in Sahrawi, individual from three villages in Western Uganda, and 61 patients in rural Bangkok (Dib et al., 2008; Johnston et al., 2010; Lalle et al., 2009; Sánchez-Vega et al., 2006; Solarczyk et al., 2010; Tungtrongchitr et al., 2010; Usluca et al., 2010). The prevalence of Giardia was estimated between 6.02% and 9.34% and can be as high as 54.8% in day-care settings in Cuba (Cañete et al., 2012; Jerez Puebla et al., 2015; Puebla et al., 2014). The

70 frequency of asymptomatic and symptomatic infections in Cuba was much higher in the children and accounted for 57 and 45%, respectively, (Pelayo et al., 2008). A slightly lower prevalence is often reported in the adult population; infection rates were 25.1% and up to 14% in pregnant women in Mexico and African migrants to the US (Manzardo et al., 2008; Robertson et al., 2010). However, it worth noting that steps in molecular analysis process; from DNA extraction, PCR amplification, target gene selected (multi copy or single copy) etc. - can significantly affect the sensitivity and specificity of molecular methods, thereby impacting the evaluated prevalence of giardiasis (Thompson & Ash, 2016). Low prevalence as reported from many countries may also be the result from infections being underreported, e.g. while Germany, where giardiasis is a reportable disease, reports around 4,000 cases every year, Austria, where giardiasis is not a reportable disease, reports only around 50 cases, although it has approximately 1/10 of the population of Germany (ECDC annual reports).

On molecular analysis of samples collected from humans and dogs in Jamaica, Giardia, assemblage A was the only genotype found. The finding was similar to four studies conducted in Mexico which reported a predominance of assemblage A in humans and animals including dogs (Cedillo-Rivera et al., 2003; Eligio-Garcia et al., 2008; Lalle et al., 2005a; Ponce-Macotela et al., 2002). Assemblage A was responsible for up to 75% of cases among 41 patients in Egypt and in 100% of samples tested from another study conducted in Mexico (Anuar et al., 2014; Helmy et al., 2009). However, In La Habana, Brazil, assemblage B was the most common cause of giardiasis in children with a prevalence of 55% (Jerez Puebla et al., 2015). Four genotypic studies conducted on isolates from Cuba showed that assemblage B was more common than assemblage A (Puebla et al., 2016). Assemblage B was also most dominant in humans in Leon, Nicaragua while host-specific assemblage C and D were associated with infections in dogs (Lebbad et al., 2008; Puebla et al., 2016; Puebla et al., 2014). Despite being a common cause of giardiasis in humans in many countries, assemblage B has yet to be identified in humans in Jamaica. While such comparisons may provide information on infection dynamics in human and animals, they should be approached with caution as many studies conducted are based on analysis of a single gene (Eligio-Garcia et al., 2008).

A different picture was seen on molecular characterization of isolates from patients in Austria where assemblage B accounted for the vast majority (65.4%) of infections while Assemblage A occurred in 34.6%.Comparing these results to other European countries, e.g. assemblage B was identified in 61.1% (127 of 208) of patients from Sweden, and from 60% (21 of 35) of sporadic cases in England and Wales (Amar et al., 2002; Lebbad et al., 2011). In German travellers, higher prevalence of assemblage A was seen over B (Broglia et al., 2013). In poor communities in Rome and symptomatic patients in Hungary and Slovakia, assemblage B was predominantly

71 identified based on analysis of the tpi gene (Marangi et al., 2010; Plutzer et al., 2014; Strkolcova et al., 2016).

Generally, assemblage B was most often responsible for food and waterborne outbreaks being most commonly identified Beavers which is considered an important reservoir for distributing infections cysts in fresh water (Amar et al., 2002; Daly et al., 2010; Karon et al., 2011; Minetti et al., 2015; Robertson et al., 2006b; Sulaiman et al., 2003). This was suspected to be due to increased virulence and which resulted in a more active transmission cycle (Lebbad et al., 2008; Lebbad et al., 2011). It was also reported that children under 15 years old are at higher risk of contracting infections with assemblage B (Anuar et al., 2014). Although assemblage B was identified as the most common cause of giardiasis in most countries around the world, assemblage A is also frequently responsible for cases of human giardiasis (Feng & Xiao, 2011; Thompson & Monis, 2004a).

Further subtyping on all genotypes showed that sub-assemblage AII was the most common cause of giardiasis in humans and dogs in Jamaica (84.2% and 70.5%, respectively). This was followed by assemblage AI which was reported at a rate of 19.8% and 29.5% in humans and dogs, respectively. In Austria, sub-assemblage BIV (46.2%) was the most common cause of giardiasis. This was followed by sub-assemblage AII, identified in 25% (13/52), BIII (19.2%) and then AI (9.6%). These findings show clearly that variations exist in different geographic locations. Giardia subtypes AI and AII show differences in host preferences, with AI commonly implicated in animal infections while AII typically in human infections (Sprong et al., 2009). However, it is worth noting that sub-assemblage AII is most common in dogs in Jamaica. These results could suggest that human infections could be acquired from dogs directly or indirectly. Such evidence suggests that dogs may act as reservoirs in maintaining and transmitting infections to humans further facilitating a zoonotic cycle. As such, a high active zoonotic-anthroponotic cycle is suspected. The opposite is also possible where humans may be responsible for a establishing a anthrophonic-zoonotic cycle. Human to human transmission will result in the same genotype being transferred from one person to the next. As such, the high prevalence of sub-assemblage BIV and AII in humans in Austria suggests that infections were likely acquired through anthroponotic transmission. However, sub-assemblages AI, AII, BIII and BIV have also all been isolated from other hosts including cattle, monkey, horses, dogs and cats (Ballweber et al., 2010; Caccio & Ryan, 2008; Sprong et al., 2009).

Molecular analysis of all samples isolated from dogs provided no evidence of host-specific genotypes such as assemblage C and D in Jamaica. Such genotypes are reportedly the most common cause of giardiasis in dogs and cats and are designated host-specific because they

(almost) only occur in dogs and cats. Assemblage C and D accounted for up to 76.5% of cases in one study (Berrilli et al., 2004). Another study conducted from the Caribbean islands of Trinidad and Tobago showed that assemblage C and D were identified in dogs at 15.4% and 80.8%, respectively (Mark-Carew et al., 2013). However, high genotypes diversity was seen in dogs in Germany where assemblage A, B, C, and D were identified. Host specific assemblages C and D were more frequently detected, namely in 75.6% (93/123) of samples tested (Pallant et al., 2015). Another study conducted in Germany included asymptomatic dogs, which also were the focus of our study, and reported very few host-specific assemblages (assemblage C at 9% and assemblage D at 0%) while assemblage A was identified at a much higher rate of 60% (Leonhard et al., 2007). In this study, these genotypes may have been undetected due to a high prevalence of mixed infections and species-specific primers may be required for such detection. The predominance of assemblage A seen in dogs in this study may also be due to the cross- sectional nature of sample collection from asymptomatic animals. As a result, this high prevalence of assemblage A may indicate that dogs are asymptomatic carriers of the assemblage. The fact that the study population consisted mainly of adult dogs could also play a role in the lack of identification of host-specific assemblage C and D which are most commonly reported in young pups (Alves & Santos, 2016).

The predominance of one circulating genotype in both humans and dogs in Jamaica provides evidence of zoonotic transmission. Zoonotic transmission is possible due to the large number of free roaming dogs on the island which has no leash laws. As such, dogs may act as a source of contamination of the environment and eventually food and water thereby facilitating transmission to humans. However, more robust molecular epidemiological methods are needed to establish a direct connection. Also necessary to be taken into account is “reverse zoonotic” transmissions which have been demonstrated in Uganda and Canada in non-human primates and muskoxen (Johnston et al., 2010; Kutz et al., 2008; Thompson, 2013; Thompson & Ash, 2016).

There were no cases of mixed infections in humans in Austria. Also in a French study, no mixed infections were found; all isolates were assemblage A (Bertrand, Albertini et al. 2005). Mixed infections have been reported with sub-assemblage AII and assemblage B from patients in England (Amar et al., 2002). Mixed infections with assemblages A and B have been reported in up to 41% in another study (Sprong et al., 2009). Cases of mixed infection have also been reported from India, Italy, and Australia where assemblage A and B are most commonly responsible (Carvalho-Costa et al., 2007; Epe et al., 2010; Feng & Xiao, 2011). Mixed infections might be more likely detected with a larger sample population; however, mixed infections could very well simply be infrequent in areas of low endemicity (Anuar et al., 2014).

Using the tpi gene combined with nested PCR, high sensitivity and specificity of detection and identification was achieved with stool samples from both, Austria and Jamaica. However, when the tpi and gdh genes were used as targets, much lower amplification rates were seen previously reported (Thompson & Ash, 2016). Primers targeting the tpi locus have shown detection limits of 1000 pg/ µl while PCR conducted on the SSU rRNA had limits of 10 pg of DNA/microliter (Jaros et al., 2011). High successful amplification rate at the tpi may also be associated with shorter amplification length. There is a correlation between high amplification yield when the tpi gene was targeted which was well suited for studies that assessed the evolutionary relationship within Giardia (Feng & Xiao, 2011). Similar results were seen when a multilocus approach was taken with rates of amplification at the tpi, bg and gdh locus at 70%, 45% and 33% (Huey et al., 2013).

4.2. Publication III

The third study that constitutes this thesis revealed that that a modified extraction protocol involving an initial pre-treatment step followed by standard amplification can successfully amplify DNA from Giardia cysts stored in formalin for more than 10 years. DNA was successfully extracted from a total of 64.4% of analysed samples (29/45). Other studies extracting genomic DNA from formalin-fixed and paraffin-embedded samples used similar protocols employed in this study with proven amplification success (Pikor et al., 2011; Sengüven et al., 2014).

The results of this study have shown high amplification rates which differ with each genetic locus. The tpi gene amplifying fragments of 334 bp showed the highest amplification rate of 64.4% (29/45), while the bg locus of 511 bp length and the gdh locus of 530 bp in nested reactions showed amplification rates of 40% (18/45) and 20% (9/45), respectively. This difference between loci was also hypothesized in other studies which proposed that PCR amplification on samples extracted from formalin show an increase amplification yield when fragments below 350 bp were targeted (Frank et al., 1996; Lin et al., 2009; Takano et al., 2010). The bg and gdh gene targets are significantly longer. Another study assessing the performance of DNA extraction on formalin-fixed tissues also showed that it is less likely to amplify gene fragments longer than 300bp (Dietrich et al., 2013). Molecular epidemiological studies conducted on giardiasis in humans and animals using samples fixed in ethanol revealed a similar trend. In our own study, also the tpi gene showed the highest amplification rate, namely of 100% (63/63) while the bg and gdh loci showed significantly lower amplification rates (70%; 44/63 and 55.6; 35/63) (Lee et al., 2017). One study on extracting genomic DNA from formalin-fixed samples also showed that amplification rates were highest when the gene fragment is short; fragments of

152 bp, 268 bp, 676 bp, yielded amplicons in 69%, 17% and 5% , respectively (Gillio-Tos et al., 2007).

When samples that gave no amplicons were subjected to re-amplification using higher DNA concentrations, amplification still remained unsuccessful. This may be as a result of severe DNA degradation, fragmentation or lack of DNA with sufficient integrity. This could also be as a result of inhibitors which may prevent attachment of polymerase to the initiation sites.

Along with modified extraction, a robust PCR protocol was also necessary to optimize amplification. PCR was repeated on false negative samples using the 4-fold concentration of DNA polymerase to antagonize DNA debris formed on fixation which may act as inhibitor preventing fragment elongation (Dietrich et al., 2013; Frank et al., 1996). Even though the concentration of polymerase was increased, only 2 samples produced positive results at the tpi and bg locus. Because there are multiple factors which may contribute to samples producing false negative results, a single conclusion cannot be drawn. Notwithstanding, higher concentrations of DNA polymerase were useful to reduce PCR inhibitors

There was a clear correlation between the lengths of storage and successful amplication. Also here, significant differences were seen between the different loci. Even samples stored for more than 10 years showed 100% amplification using the tpi gene. However, at the bg and gdh loci only samples stored for shorter periods of time showed positive results. This suggests that the longer samples are stored in formalin does not affect the amplification yield when very short fragments, such as the tpi gene are used as diagnostic tool. In other studies, the correlation between the length of storage and successful amplification was not so evident. Samples stored over a three year period showed more or less the same amplification pattern (Lin et al., 2009; Niland et al., 2012). Even DNA fixed for as long as 20-25 years can be successfully amplified using phenol and xylene purification (Gillio-Tos et al., 2007; Santos et al., 2008). Another study demonstrated that DNA from tissue fixed in formalin for 8 years with targeted fragments of up to 536 bp and 989 bp could generally be amplified, although with very low success rates. The same gene fragments were targeted from samples fixed in 95% ethanol which showed very high amplification success rates even in fragments that were much longer (1327 bp) (Greer et al., 1991).

In analysing the DNA quality in genotyping, it was shown that high quality sequences can be obtained from samples fixed in formalin. This allowed for genotyping and even subtyping of around half of the samples investigated. Sequencing errors were observed at all three loci. Sequencing errors of the G/A and C/T type from samples fixed in formalin were also seen in other studies and were presumably caused by failure of the polymerase to recognize cytosine

75 and guanine which leads to the substitution for another nucleotide (Astolfi et al., 2015; Do & Dobrovic, 2012). One sequencing error per 500 bp can be expected in formalin-fixed material, the majority of the errors concerning the cytosine or guanine position (Williams et al., 1999). Such artifacts usually occur as a result of damaged or cross-linking nucleotides, however, Taq DNA polymerase is also responsible for falsely incorporating adenosine residues (Williams et al., 1999).

4.3. Conclusion

The current molecular epidemiological investigation suggests that giardiasis is a significant public health concern in Jamaica. The same genotype has been implicated in infections in both, humans and dogs. The significantly higher prevalence in dogs shows that they are more likely infected and are suspected key players in the zoonotic cycle. Molecular analysis of Giardia strains from patient samples in Austria revealed a comparably high level of genetic diversity which may support the assumption that most cases of human giardiasis in Austria are imported. Findings also suggest that assemblage B is more likely to be diagnosed as cause of human giardiasis because it is assumed to be more virulent and thus infections with this assemblage are more likely to become clinically manifest. Multilocus sequence analysis targeting the tpi, gdh and bg locus is necessary for understanding the distribution dynamics of the various Giardia subtypes. Thus, the findings from this dissertation should contribute to a better understanding of Giardia epidemiology, transmission dynamics and zoonotic potential. This dissertation also showed that DNA can be successfully amplified form Giardia cysts fixed in formalin. Amplified products produced sequences which were able to differentiate Giardia at the subtype level. The adapted and optimized protocols established during this thesis may be valuable for phylogenetic studies on old and historic samples, not only of Giardia spp., but also on other protozoan parasites. Therefore, this method provides great promise for the conduct of retrospective molecular diagnostics and epidemiological studies.

Publications

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Curriculum Vitae

Mellesia F. Lee

Marlene Hamilton Hall University of The West Indies Kingston, Jamaica +867 861 3151 [email protected] EDUCATION

2014-present Doctor of Philosophy Medical University Vienna, Austria

2010-2013 Master of Science in Medical Microbiology University Of The West Indies, Kingston, Jamaica topic: “A Retrospective Analysis of Pathogens Associated With Blood Stream Infections and their Antimicrobial Resistance Pattern at the UHWI during September 2011- December 2011”

2007 – 2010 Bachelor of Science in Nursing University Of The West Indies, Kingston, Jamaica

2001 – 2007 CXC (Caribbean Examination Council) CAPE (Caribbean Advanced Proficiency Examinations) St. Andrew High School for Girls, Kingston, Jamaica Major fields of study: Biology, Physics, Chemistry, Mathematics, English Language, Religious Education and Social Studies

PUBLICATIONS

1. Lee M, Auer H, Lindo J, Walochnik J. 2017. Multilocus Sequence Analysis of Giardia Spp. Isolated from Patients with Diarrhea in Austria. Parasitology Research 116(2): 477-481.

2. Lee M, Cadogan P, Eytle S, Copeland S, Walochnik J, Lindo J. 2017. Molecular Epidemiology and Multilocus Sequence Analysis of Potentially Zoonotic Giardia Spp. From Humans and Dogs in Jamaica. Parasitology Research 116(1): 409-414.

PROFESSIONAL AND LABORATORY EXPERIENCE

2010- present Research Technologist/Research Laboratory Assistant Department Of Microbiology, University Of The West Indies, Kingston, Jamaica

Processing and storage of blood and stool samples from:  Bustamante Hospital for Children, Kingston, Jamaica  National Public Health Laboratory, Kingston, Jamaica  University Hospital of the West Indies, Kingston, Jamaica Sample Management and Storage

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 Processing of samples for serological testing  Conduct rapid testing techniques such as HIV, Hepatitis, etc.  Data entry and appropriate sample storage Diagnosis in the discipline of Parasitology, Bacteriology, Virology  Detection of malaria antigen in blood for rapid diagnosis of disease  Diagnosis of Trichomona vaginalis at the point of care at the UHWI  Conduct concentration and staining techniques for intestinal parasites  Conduct serological testing of blood samples for Malaria, Leptospirosis, EBV,HTLV, VZV, HSV (1&2) antibodies using ELSIA  Direct and indirect IFA on stool and swabs, and tissue culture processing, staining techniques for bacterias  Laboratory work-up on GPC, GPB, GNB, anaerobes  Data entry and sample arrangement for VITEK 2 system (bioMérieux,Durham and BacT/ALERT® 3D (bioMérieux, Durham, North Carolina) machines Molecular testing  Data entry and appropriate sample storage  Conduct DNA extraction and amplification of parasite DNA  Molecular characterization; Giardia and Ancylostoma  DNA sequencing; Giardia and Ancylostoma genes  Sequence analysis (CLUSTAL X® and GeneDoc® version 2.7.000 Data management  Accurate electronic record keeping  Responsible for statistical analysis of data  Any other duties deemed necessary by the Principal Investigator Reporting  Reports to the Principal Investigator  Monthly report to the Ministry of Health (ELISA IgG-Dengue) 2007–2010 Clinical Practicum (Various Hospitals, Health Centers and Mental Institutions in Jamaica)

Medical practical experiences:  Physical assessment, mental status examination of hospitalized patients  Assist in diagnosis and treatment of medical, surgical, obstetrics, gynaecologic, paediatric, psychiatric, geriatric and critically ill patients in intensive care units  Holistic patient care: venepuncture, medication administration (IV, IM), assist in passing tracheostomy, nasogastric tube and chest tube

LEADERSHIP

2008-2009 CEAC (Culture and Entertainment Affairs Chairperson) Mary Seacole Student Hall, University Of The West Indies, Kingston, Jamaica

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2007-2009 Sports Chair Person Mary Seacole Student Hall, University Of The West Indies, Kingston, Jamaica

2004-2005 Student Council St. Andrew High School For Girls, Kingston, Jamaica

AWARDS

2007-2013 Several certificates for outstanding academic work University Of The West Indies, Kingston, Jamaica

2006 Platinum for outstanding work in CXC Top performance in class Association Of Principals And Vice Principals, Kingston, Jamaica

CONFERENCES AND INTERNATIONAL EXPERIENCE

2010 – 2013 Attendance at various International Conferences held at the University of the West Indies

2013 15th International Meeting On the Biology And Pathogenicity Of Free Living Amoebae (FLAM) Conference in Vienna, Austria

2012 Graduate Cultural Exchange Work & Travel Wisconsin USA

2011 14th International Meeting On the Biology And Pathogenicity Of Free Living Amoebae (FLAM) Conference In Montego Bay, Jamaica

CONFERENCES AND INTERNATIONAL EXPERIENCE

Languages English: Native German: Basic Computer skills MS office Experienced SPSS: Experienced Professional LaTex Experienced

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