CHARACTERISATION OF NATIVE

TRYPANOSOMES AND OTHER

PROTOZOANS IN THE AUSTRALIAN

MARSUPIALS THE QUOKKA (SETONIX

BRACHYURUS) AND THE GILBERT’S

POTOROO (POTOROUS GILBERTII)

Thesis presented by

Jill Austen

Bachelor of Science with First Class Honours

in Biomedical Science

for the degree of

Doctor of Philosophy

School of Veterinary and Life Sciences

Murdoch University

2015

i

DECLARATION

I declare that this thesis is my own account of my research and contains as its main content, work that has not previously been submitted for a degree at any tertiary institution.

Jill Austen

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ACKNOWLEDGEMENTS

First and foremost, I would like to express my sincere gratitude and appreciation to my supervisors: Professor Una Ryan, Associate Professor Simon Reid, Dr Tony

Friend and Dr William Ditcham. Una you introduced me into the wonderful world of parasites over a decade ago and since then my fascination for microscopic organisms has grown experientially. Your exceptional knowledge, guidance, belief in my research and kindness have never failed to impress me and I hope that one day I may strive to walk in your footsteps. If earth angels do exist I truly believe you are one. Simon, or should I say trypanosome guru. Thank you so much for initially introducing me to your parasite of choice ‘Trypanosomes”. These blood borne parasites have won my heart and curiosity and trypanosomes are now my preferred parasite of choice. I will always remember your brain storming sessions and exceptional advice and for that I am truly grateful. Tony, thank you so much for your continuous support, wildlife expertise and early morning walks through the bush. This PhD would not have been possible without your collaboration and dedication and our adventures on Rottnest Island will never be forgotten. William, thank you so much for your exceptional laboratory intellect and for always being my go to man for when I had endless questions. You always managed to enlighten me with your novel ideas and boost my self-esteem in the process and for that

I am truly grateful.

I would like to acknowledge the support and friendship extended by the staff and students of the Vector and Waterborne Pathogens Group and staff at Murdoch

University. Particular thanks go to Professor Peter Irwin (with his famous quote ‘You need more evidence’), Frances Brigg, Gordon Thompson, Peter Fallon, Dr Kirsty

Townsend, Russell Hobbs, Aileen Elliot, Dr Scott Edwards, Dr Rongchang Yang, Garry

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Allen, Ryne Charsley, Gavin D’Mello, Michael Slaven and Gerard Spoelstra for their skills and assistance in facets of this study.

Thank you to my fellow colleagues who have now become close personal friends, particular thanks goes to Tegan McNab and Linda Davies for their help on

Rottnest Island, my office buddies, Andrea Ducki, Ahmed (aka the King,), and Emilija

Filipovska-Naumovska for all the laughs over the years as well as Bong Sze How, Ryan

Jefferies, Joanne McCoubrie, Annika Estcourt, Mark O’Dea, and Amanda Barbosa for making my PhD student days memories to treasure.

Family and friends without your continuous love and support I would not have been able to have accomplished so much and fulfilled my dream of studying Australian trypanosomes. I would like to thank my mum (Margaret Smith) and dad (Chris

Meinema) for always believing in me and encouraging me to live my dreams and to never give up. I have learnt that a successful PhD relies mostly on the support you get from home. I am therefore truly indebted to my husband Jeff who has continuously supported me throughout my studies and has been the sole carer of our three gorgeous boys on the weekends. Jeff you have sacrificed so much, therefore I would like to dedicate this thesis to you and our three beautiful boys Jacob, Dylan and Brodie.

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SYMBOLS AND ABBREVIATIONS

Symbols

~ approximately

= equals

> greater than

< less than

- minus

% percent

x times

± plus-minus sign

Abbreviations

µL Microlitre ACP Alternative complement pathway AIDS Acquired immunodeficiency syndrome AMP Ampicillin ANOVA Analysis of variance ApoLi Apolipoprotein Li BI Bald Island BIIT Blood incubation infectivity test B Breadth measurment bp Base pair C° Degrees Celsius CI Confidence interval CM Cunninghams liquid medium

CO2 Carbon dioxide CO1 Cytochrome c oxidase subunit 1 CRISPs Cysteine-rich secretory proteins DA Diminazene aceturate DAPI 4’6-diamidino-2-phenylindole DNA Deoxyribonucleic acid dNTP Deoxynucleoside triphosphate v

EDTA Ethylenediamine tetra acetic acid Eg or e.g. Exempli gratia – for example et al. et alia: and others. EIDs Emerging infectious diseases FAM 6-carboxy fluorescein, acronym FCS Foetal calf serum FISH Fluorescent in situ hybridisation FITC Fluorescein isothiocyanate FF Free measurement GAPDH glyceraldehyde-3-phosphate dehydrogenase g relative centrifugal force HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HDL High density lipoprotein HPR Human haptoglobin-related protein HSS Human serum sensitive HuMSEM Human modified sloppy Evan’s medium HoMSEM Horse modified sloppy Evan’s medium

H2O Water i.e. id est – in other words ICN Pharmaceutical company IgG Immunoglobulin G IPTG Isopropyl β-D-1-thiogalactopyranoside IU International units k Kinetoplast K+ Potassium kb Kilobase kDNA Kinetoplast deoxyribonucleic acid Km Kilometers KN Kinetoplast to nucleus measurement L Liter LB Luria-Bertani L8C4 Anti-paraflagellar rod antibody M Molar concentration MAC Membrane attack complex mg Milligram

MgCl2 Magnesium chloride min Minute ML Maximum likelihood

vi mL Millilitre mM Millimolar mm Millimetre MP Maximum parsimony mtDNA Mitochondrial deoxyribonucleic acid MSEM Modified sloppy Evan’s medium n Number NA Nucleus to anterior measurement Na+ Sodium NCBI National Center for Biotechnology Information ng Nanograms NHS Normal human serum NSW New South Wales p Probability of an event due to chance alone PBS Phosphate-buffered-saline PCR Polymerase chain reaction PCV Packed cell volume pers. comm. Personal communication PH Negative logarithm of hydrogen ion concentration PI Post inoculation PK Posterior to the kinetoplast measurement pmol Picomoles PN Posterior to nucleus measurement Qld Queensland rDNA Ribosomal deoxyribonucleic acid RNA Ribonucleic acid RI Rottnest Island RPM Revolutions per minute SD Standard deviation SE Standard error SEM Scanning electron microscopy SOC Super Optimal Broth sp. Unknown species (singular) sp. n Novel species spp. Several species SPSS Statistics Package for Social Studies SRA Serum resistance associated gene 18S rRNA 18S ribosomal RNA

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Taq Thermus aquaticus deoxyribonucleic acid polymerase TEM Transmission electron microscopy TgsGP T. b. gambiense specific gene TL Total length TLFs Trypanosome lytic factors TPB Two Peoples Bay USA United States of America UV Ultra violet light VSG Variable surface glycoprotein WA Western Australia WHO World Health Organisation YT Yeast extract and tryptone X-GAL 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside

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ABSTRACT

Trypanosomes are blood-borne parasites that can cause severe disease in both humans and animals, resulting in very large economic losses worldwide. In contrast to the wealth of information on pathogenic species such as cruzi and

Trypanosoma brucei, little is known of the pathogenicity, prevalence and life-cycles of trypanosomes in native Australian mammals. The aim of this thesis was to characterise trypanosomes and other protozoans from the critically endangered Gilbert’s potoroo

(Potorous gilbertii) and the quokka (Setonix brachyurus) from Western Australia using morphological and molecular analysis.

A novel Trypanosoma species, Trypanosoma copemani was identified in

Gilbert’s potoroos and quokkas using molecular and morphological analysis. Further molecular characterisation of T. copemani in quokkas at both the 18S rRNA and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) loci identified multiple T. copemani genotypes revealing that the parasite is genetically variable. Given the phylogenetic grouping of T. copemani as a stercorarian (requiring faecal transmission by vector), the fact that are common ectoparasites of Gilbert’s potoroos and quokkas and the previous identification of trypanosomes in ticks, ticks were examined as potential vectors of T. copemani. Motile trypanosomes were detected in both the haemolymph and midgut sections of Ixodes australiensis ticks removed from quokkas and Gilbert’s potoroos and stained trypanosomes were detected within a faecal smear.

Morphologically, the trypanosomes resembled in vitro forms of T. copemani, representing epimastigotes and slender trypomastigote stages, with dividing stages detected within the midgut region. Molecular analysis of the tick isolates, showed 100% sequence identity to T. copemani at the 18S rRNA locus, suggesting that the tick is a putative vector for T. copemani.

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Investigation of the life-cycle stages of native Australian trypanosomes using microscopy and in vitro culture of blood from quokkas and Gilbert’s potoroos revealed that native Australian trypanosomes are highly polymorphic, with three different trypomastigote blood stream forms detected within blood films, resembling slender, medium and broad stages. In addition, promastigote, sphaeromastigote and amastigote stages were observed directly within blood films and this is the first report of these stages in the circulatory system of Australian . Three novel trypanosome life- cycle forms representing an oval form, an extremely thin form and an adherent form were also identified both in vitro and in vivo, while a novel tiny form and a novel circular form were only detected in culture. Trypanosoma vegrandis was also detected for the first time in one quokka isolate (Q1340) and confirmed using species-specific primers.

As tourists and quokkas on Rottnest Island have a close relationship, the zoonotic potential of T. copemani was investigated by determining the relative susceptibility of T. copemani to human serum using the blood incubation infectivity test. Trypanosoma copemani was observed by microscopy in all human blood cultures from day 5 to day 14 post inoculation. The mechanism for normal human serum resistance in T. copemani is not known. The results of this thesis show that at least one native Australian trypanosome species may have the potential to infect humans.

In addition to blood, quokka faecal samples were also collected to investigate the prevalence of Eimeria by PCR screening of faecal samples from three quokka populations, Two Peoples Bay, Bald Island and Rottnest Island, respectively. The PCR prevalence of Eimeria was 62.5%, 85.0% and 78.3% for these three locations respectively. Two Eimeria species were identified based on morphometric analysis from sporulated oocysts, Eimeria quokka and Eimeria setonicis, with the majority of quokkas co-infected with both species. Singular infections, however, were identified in a few

x individuals allowing molecular analysis to be performed using both the 18S rRNA and cytochrome c oxidase subunit 1 genes. Phylogenetic analysis grouped E. quokka and E. setonicis within the Eimeria clade. This study is the first one to characterise

E. quokka and E. setonicis by molecular analyses, enabling more extensive resolution of evolutionary relationships among marsupial-derived Eimeria species.

Overall this study has characterised the novel T. copemani in both the quokka and the Gilbert’s potoroo and has shown T. copemani to be both morphologically and genetically variable. This is a stercorarian trypanosome, transmitted through vector faecal contamination and may potentially be infectious to humans. The findings of this thesis highlight the complexity of Australian trypanosomes and provide insights into the prevalence, potential pathogenicity and human infectivity of this novel parasite. Further research is required to assess the role of trypanosome infection on marsupial population dynamics and the implications for management and conservation.

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PUBLICATIONS AND CONFERENCES

Publications arising from this thesis:

Austen, J.M., Jefferies, R., Friend, J.A., Ryan, U., Adams, P., Reid, S.A., 2009.

Morphological and molecular characterisation of Trypanosoma copemani n. sp.

(Trypanosomatidae) isolated from Gilbert's potoroo (Potorous gilbertii) and quokka

(Setonix brachyurus). Parasitology 136, 783-792.

Austen, J.M., Ryan, U.M., Friend, J.A., Ditcham, W.G.F., Reid, S.A., 2011. Vector of

Trypanosoma copemani identified as Ixodes sp. Parasitology, 138, 866-872.

Austen, J.M., Friend J.A., Yang, R., Ryan, U.M. 2014. Further characterisation of two

Eimeria species (Eimeria quokka and Eimeria setonicis) in quokkas (Setonix brachyurus). Experimental Parasitology 138, 48-54.

Austen, J.M., Ryan, U. M., Ditcham, W. G. F., Friend, J. A. and Reid, S. A. 2015. The innate resistance of Trypanosoma copemani to human serum. Experimental

Parasitology. 153, 105-110.

Austen, J.M., Reid, S. A., Robinson, D. R., Friend, J. A., Ditcham, W. G. F., Irwin P.

J., Ryan,U. 2015. Investigation of the morphological diversity of the potentially zoonotic Trypanosoma copemani in quokkas and Gilbert’s potoroos. Parasitology, 142,

11, 1443-52.

Austen, J.M., Paparini, A., Reid, S. A., Friend, J. A., Ditcham,W. G. F., Una Ryan, U.

2015. Molecular characterisation of native Australian trypanosomes in quokka (Setonix brachyurus) populations from Western Australia. Parasitology International, 65, (3),

205-208.

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Abstracts in Conference proceedings

Austen, J.M., Ditcham, W. G. F., Ryan, U.M., Friend, J.A., Reid, S. A. 2010. Human serum resistance in a native Australian trypanosome: is it zoonotic? XIIth International

Congress of Parasitology (ICOPA) conference 15-20 August, Melbourne, Australia.

Austen, J.M., Ryan U.M., Friend, J.A., Ditcham, W. G. F., Reid, S. A. 2013.

Identification of the tick as the vector for T. copemani. ASP and WAAVP conference

25-29 August, Western, Australia.

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TABLE OF CONTENTS

DECLARATION ...... ii ACKNOWLEDGEMENTS ...... iii SYMBOLS AND ABBREVIATIONS ...... v ABSTRACT ...... ix PUBLICATIONS AND CONFERENCES ...... xii TABLE OF CONTENTS ...... xiv LIST OF FIGURES ...... xix LIST OF TABLES ...... xxvii CHAPTER 1 - LITERATURE REVIEW ...... 1 1.1 General history of trypanosomes ...... 2 1.2 and phylogeny ...... 3 1.3 General morphology of trypanosomes ...... 5 1.4 Salivarian and stercorarian mammalian trypanosomes ...... 7 1.4.1 The salivarian trypanosomes ...... 8 1.4.2 The stercorarian trypanosomes ...... 9 1.5 Vectors of trypanosomes ...... 12 1.6 Life-cycle of trypanosomes ...... 14 1.6.1 Salivarian life-cycle ...... 15 1.6.2 Stercorarian life-cycle ...... 16 1.6.2.1 Development within the mammalian host ...... 16 1.6.2.2 Development within the vector ...... 17 1.7 Clinical signs and pathogenesis of trypanosomiasis ...... 18 1.7.1 Pathogenesis of salivarian trypanosomes ...... 18 1.7.2 Pathogenesis of stercorarian trypanosomes ...... 20 1.7.3 Human pathogenic trypanosomes ...... 21 1.7.4 Drug therapies and vaccines...... 24 1.8 History of Australian trypanosomes ...... 25 1.9 Trypanosomiasis in Australian marsupials ...... 35 1.10 Characterisation of trypanosomes ...... 36 1.11 Molecular characterisation of trypanosomes ...... 37 1.12 Phylogenetic analysis ...... 38 1.13 Western Australian marsupials the Gilbert’s potoroo and quokka ...... 40 CHAPTER 2 - GENERAL MATERIALS AND METHODS ...... 44 2.1 Sample collection and geographical locations ...... 45 2.2 Collection of ectoparasites ...... 46 2.3 Detection of trypanosomes ...... 46 xiv

2.3.1 Preparation of blood film ...... 46 2.3.2 In vitro cultivation of trypanosomes ...... 46 2.4 Immunofluorescence ...... 47 2.5 Scanning electron microscopy (SEM) ...... 48 2.6 Morphological analysis of trypanosomes ...... 49 2.7 DNA analysis of trypanosomes ...... 50 2.7.1 DNA extraction ...... 50 2.7.2 DNA amplification of the 18S rRNA gene...... 50 2.7.3 DNA amplification of the GAPDH gene ...... 51 2.7.4 PCR controls and agarose gel electrophoresis ...... 51 2.7.5 Purification of PCR products ...... 52 2.8 Cloning ...... 52 2.9 Sequencing ...... 54 2.10 Phylogenetic Analysis ...... 55 CHAPTER 3 - MORPHOLOGICAL AND MOLECULAR CHARACTERISATION OF TRYPANOSOMA COPEMANI (TRYPANOSOMATIDAE) ISOLATED FROM GILBERT’S POTOROO (POTOROUS GILBERTII ) AND QUOKKA (SETONIX BRACHYURUS) ...... 56 3.1 Introduction ...... 57 3.2 Materials and Methods ...... 58 3.2.1 Study site and sample collection ...... 58 3.2.2 Detection of trypanosomes in whole blood ...... 58 3.2.2.1 Preparation of blood films ...... 58 3.2.3 Genetic characterisation and phylogenetic analysis ...... 59 3.3 Results ...... 60 3.3.1 Microscopy ...... 60 3.3.2 In vitro culture ...... 62 3.3.3 Statistical analysis of morphological measurements ...... 63 3.3.4 Sequence analysis of Trypanosoma copemani isolated from the Gilbert’s potoroo and quokka ...... 64 3.3.5 Phylogenetic relationships of the Trypanosoma spp. from Gilbert’s potoroo and quokka ...... 65 3.4 Discussion ...... 68 3.4.1 Infection and pathogenesis ...... 69 3.4.2 Morphology ...... 70 3.4.3 Evolutionary and taxonomic relationships ...... 72 CHAPTER 4 - VECTOR OF TRYPANOSOMA COPEMANI PUTATIVELY IDENTIFIED AS IXODES SP...... 74 4.1 Introduction ...... 75 4.2 Materials and Methods ...... 76 xv

4.2.1 Study site and sample collection...... 76 4.2.2 Detection of trypanosomes in ectoparasites ...... 77 4.2.3 Detection of trypanosomes in tick faeces ...... 77 4.2.4 Morphological measurements ...... 78 4.2.5 DNA extraction ...... 78 4.2.6 DNA amplification and sequencing...... 78 4.3. Results ...... 78 4.3.1 Identification of ectoparasites ...... 78 4.3.2 Trypanosomes present in tick faeces ...... 79 4.3.3 Trypanosomes present in ectoparasites ...... 81 4.3.4 Comparative analysis of morphological measurements of trypanosomes from quokka blood and from ticks...... 82 4.3.5 Characterization of T. copemani isolated from blood and ticks ...... 83 4.4 Discussion ...... 84 CHAPTER 5 - INVESTIGATION OF THE MORPHOLOGICAL DIVERSITY OF NATIVE AUSTRALIAN TRYPANOSOMES IN QUOKKAS AND GILBERT’S POTOROOS ...... 88 5.1 Introduction ...... 89 5.2 Materials and Methods ...... 90 5.2.1 Study site and sample collection...... 90 5.2.2 Microscopy ...... 92 5.2.3 In vitro cultivation of trypanosomes ...... 92 5.2.3.1 BIIT medium ...... 92 5.2.4 Fluorescence in situ hybridisation (FISH) ...... 93 5.2.5 Molecular characterisation...... 93 5.3 Results ...... 95 5.3.1 Trypanosome morphology ...... 95 5.3.1.1 Promastigote stage ...... 96 5.3.1.2 Sphaeromastigote stage ...... 98 5.3.1.3 Amastigote stage ...... 99 5.3.1.4 Oval form ...... 99 5.3.1.5 Extremely thin form ...... 100 5.3.1.6 Adherent and free moving forms ...... 101 5.3.1.7 Tiny form ...... 102 5.3.1.8 Circular form ...... 103 5.3.4 Molecular confirmation of trypanosomes and piroplasms ...... 103 5.4. Discussion ...... 104 CHAPTER 6 - MOLECULAR CHARACTERISATION OF NATIVE AUSTRALIAN TRYPANOSOMES IN QUOKKA (SETONIX BRACHYURUS) POPULATIONS FROM WESTERN AUSTRALIA ...... 110 xvi

6.1 Introduction ...... 111 6.2 Materials and Methods ...... 112 6.2.1 Study site and sample collection...... 112 6.2.2 PCR amplification and sequencing of trypanosomes ...... 113 6.2.3 Phylogenetic analysis ...... 114 6.2.4 Statistical analysis ...... 114 6.3 Results ...... 115 6.3.1 Prevalence of trypanosomes in quokka populations ...... 115 6.3.2. Molecular characterisation of trypanosomes in quokka isolates at the 18S rRNA locus ...... 115 6.3.3 Phylogenetic analysis of trypanosome isolates from quokkas at the GAPDH gene ...... 119 6.4 Discussion ...... 122 CHAPTER 7 - THE INNATE RESISTANCE OF TRYPANOSOMA COPEMANI TO HUMAN SERUM ...... 125 7.1 Introduction ...... 126 7.2 Materials and Methods ...... 127 7.2.1 Study site and sample collection...... 127 7.2.2 DNA extraction ...... 127 7.2.3 Molecular characterisation of T. copemani ...... 127 7.2.4 In vitro human serum resistance: blood incubation infectivity test ...... 128 7.2.5 Statistical analysis ...... 129 7.3 Results ...... 130 7.3.1 Microscopy ...... 130 7.3.2 Molecular characterisation of T. copemani ...... 130 7.3.3 Blood incubation infectivity test (BIIT) ...... 130 7.4 Discussion ...... 133 CHAPTER 8 - FURTHER CHARACTERISATION OF TWO EIMERIA SPECIES (EIMERIA QUOKKA AND EIMERIA SETONICIS) IN QUOKKAS (SETONIX BRACHYURUS) ...... 139 8.1. Introduction ...... 140 8.2. Materials and Methods ...... 141 8.2.1 Sample collection...... 141 8.2.2 Screening and morphological analysis of oocysts ...... 142 8.2.3 DNA isolation and PCR amplification ...... 143 8.2.4 Sequencing and phylogenetic analysis ...... 143 8.2.5 Statistical analysis ...... 143 8.3. Results ...... 144 8.3.1 Eimeria prevalence ...... 144 8.3.2 Morphological characterisation of Eimeria oocysts ...... 145 xvii

8.3.3 Phylogenetic analysis of E. quokka, E. setonicis and Eimeria spp. from the quokka at the 18S rRNA gene ...... 148 8.3.4. Phylogenetic analysis of E. quokka and E. setonicis from the quokka at the COI gene ...... 151 8.4 Discussion ...... 153 CHAPTER 9 - GENERAL DISCUSSION ...... 158 9.1 Blood-borne and enteric parasites in marsupials and management implications 159 9.2 Morphology versus molecular characterisation ...... 163 9.3 Transmission of T. copemani ...... 165 9.4 The zoonotic potential of Australian trypanosomes ...... 165 9.5 Future studies ...... 166 9.6 Conclusions ...... 168 REFERENCES ...... 170

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LIST OF FIGURES

Figure 1.1 Morphological characteristics of the genus Trypanosoma.

Figure 1.2 Morphological stages described in Trypanosomatidae: [1] amastigote

stage; [2a] & [2b] sphaeromastigote stages; [3] promastigote; [4]

opisthomastigote; [5] epimastigote stage; [6] trypomastigote stage; [7]

choanomastigote stage.

Figure 1.3 Micrograph of T. brucei in a blood film representing a typical salavarian

trypanosome.

Figure 1.4 Micrograph of T. cruzi in a blood film, representing a typical stercorarian

trypanosome.

Figure 1.5 Diagrammatical representation of a typical salavarian life-cycle.

Figure 1.6 Diagrammatical representation of a typical stercorarian life-cycle.

Figure 1.7 Micrograph of T. lewisi in a blood film.

Figure 1.8 Micrograph of T. binneyi in a blood film isolated from the Platypus.

Figure 1.9 Micrograph of a Trypanosoma sp. in a blood film isolated from the

southern brown bandicoot.

Figure 1.10 Micrographs of Trypanosoma species isolated from Australian animals.

(A) Wombat AAP isolate. (B) Wallaby ABF isolate. (C) Currawong

isolate.

Figure 1.11 Micrograph of T. irwini in a blood film isolated from the .

Figure 1.12 Micrograph of the novel trypanosome isolate in a blood film isolated

from the common brushtail possum.

Figure 1.13 Micrograph of T. vegrandis in blood films isolated from the woylie.

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Figure 1.14 Eukaryotic rDNA array showing the 18S rRNA gene in relation to the

other RNA subunits. Two internal transcribed spacers (i.e. ITS-1 and

ITS-2) separate the 18S, 5.8S and 28S genes.

Figure 1.15a Photograph of the Gilbert’s potoroo (Potorous gilbertii).

Figure 1.15b Photograph of a quokka (Setonix brachyurus).

Figure 2.1 Map of Western Australia showing the geographical location of the three

marsupial sampling sites.

Figure 2.2 Measurable morphological diagnostic parameters of trypanosomes.

Figure 3.1 Light micrographs of Trypanosoma copemani isolated from the blood of

a Gilbert’s potoroo. (A) Slender trypomastigote form in a Modified

Wright stained blood film. (B) Medium trypomastigote form in a

Modified Wright stained blood film. (C) Broad trypomastigote form in a

Modified Wright stained blood film. (D) Dividing trypanosome form in a

Modified Wright stained blood film. Scale bar represents 10 µm.

Figure 3.2 Light Micrographs of Trypanosoma copemani isolated from the blood of

a quokka and haemoparasites isolated from the Gilbert’s potoroo and

quokka. (A) Trypomastigote form in a Modified Wright stained blood

film from a quokka and a piroplasm (indicated by the arrow) in an

erythrocyte from the quokka (B) Microfilaria in a Modified Wright

stained blood film from a Gilbert’s potoroo. Scale bars represent 10 µm.

Figure 3.3 Light micrographs of Trypanosoma copemani from in vitro cultures of

blood from a Gilbert’s potoroo. (A) Trypomastigote and epimastigote

forms in a Modified Wright stained film. (B) A dividing

sphaeromastigote form in a Modified Wright stained film. (C) Dividing

epimastigote form with two nuclei and two kinetoplasts and

sphaeromastigote form in a Modified Wright stained film. (D)

xx

Promastigote form in a Modified Wright stained blood film. Scale bar

represents 10 µm.

Figure 3.4 Distance-based phylogenetic tree of 32 Trypanosoma species inferred

using partial 18S rRNA gene sequences (790 positions). Relationships

were determined using Neighbor-joining and Maximum Composite

Likelihood methods. Bootstrap values are shown as percentages of 1,000

replicates and branches corresponding to partitions reproduced in less

than 50% bootstrap replicates are collapsed. Values shown in bold

represent support for each clade using distance, maximum parsimony

and maximum likelihood algorithms respectively. The scale bar

represents the proportion of base substitutions per site. Species from

Australian marsupials are shown with an asterisk.

Figure 3.5 Linearized distance-based phylogenetic tree inferred using partial 18S

rRNA gene sequences (503 positions) showing the relationship of T.

copemani to other closely related species. (B) Subsection of a distance-

based phylogenetic tree inferred using partial 18S rRNA gene sequences

(959 positions) revealing the 3 distinct genotypes within the species T.

copemani (A, B and C). Relationships were determined using Neighbor-

joining and Maximum Composite Likelihood methods. Bootstrap values

are shown as percentages of 1,000 replicates and units are in number of

base substitutions per site. New sequences are shown in bold.

Figure 4.1 Light micrographs of trypanosomes detected in the faeces of the tick (I.

australiensis) using a Modified Wright stain.

Figure 4.2 Light micrograph of T. copemani in Modified Wright stain detected in

the haemolymph of the tick collected from the Gilbert’s potoroo P170.

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Trypomastigote form (t), epimastigote form (e). Scale bar represents 10

µm.

Figure 4.3 Light micrographs of trypanosomes in Modified Wright stain from the

midgut region of a tick collected from quokka Q2917-2918. (A)

Trypomastigote form (t). (B) Dividing epimastigote form (e). (C)

Dividing trypomastigote form (d). Scale bar represents 10 μm.

Figure 4.4 Light micrographs of T. copemani in Modified Wright stain detected in

the blood of quokka 4489 captured from Bald Island. Scale bar

represents 10 µm.

Figure 5.1 Light micrographs of trypanosomes in a Modified Wright-Giemsa

stained blood film. (A) The slender stage in a blood film from quokka

Q1369 -1355 from Two Peoples Bay. (B) The medium stage in a blood

film from quokka Q3355-3284 from Two Peoples Bay. (C) The broad

stage in a blood film from quokka Q3355-3284 from Two Peoples Bay.

Scale bars represent 10 µm.

Figure 5.2 (A) Light micrograph of the trypanosome promastigote stage in a

Modified Wright-Giemsa stained blood film from quokka Q1340 from

Two Peoples Bay. (B) Scanning electron micrograph of the promastigote

trypanosome stage from in vitro culture originally isolated from Gilbert’s

potoroo P83. (C) Unstained trypanosome using differential interference

contrast (DIC) from Gilbert’s potoroo P83. (D) Immunofluorescent

staining of the paraflagellar rod of a trypanosome isolated from Gilbert’s

potoroo P83 using monoclonal antibody (L8C4). (E) DAPI staining of

nuclear DNA from Gilbert’s potoroo P83. (F) Combined images of C, D

and E. Scale bars represent 10 µm.

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Figure 5.3 Micrographs of sphaeromastigote trypanosomes. (A) Light micrograph

of a sphaeromastigote stage (indicated by arrow) in a Modified Wright-

Giemsa stained blood film from quokka Q1342-1338 from Two Peoples

Bay. (B) Sphaeromastigote stage from in vitro culture originally isolated

from Gilbert’s potoroo P94. (C) Immunofluorescent staining using

monoclonal antibody (L8C4) of a sphaeromastigote in vitro isolated from

a quokka Q1 from Bald Island. Scale bars represent 10 µm.

Figure 5.4 (A) Light micrograph of the trypanosome amastigote form in vivo from

quokka QRS3. (B) FISH analysis of amastigote in vivo from quokka

Q4908-4846. (C) Light micrograph of amastigote in vitro stained with

Giemsa from quokka Q1342-1338 N = nucleus, K = kinetoplast. (D)

FISH analysis of the same amastigote in in vitro culture. Scale bar

represents 10 µm.

Figure 5.5 (A) Light micrograph of the oval form trypanosomes from quokka

Q1051 from Two Peoples Bay. (B) Oval shape form trypanosome from

in vitro culture originally isolated from Gilbert’s potoroo P94. (C) Oval

form trypanosome in vitro culture from quokka Q2088-2050 stained with

Modified Wright-Giemsa stain. Scale bars represent 10 µm.

Figure 5.6 (A) Extremely thin trypanosome form in a Modified Wright-Giemsa

stained blood film from quokka Q3343-3304 from Bald Island. (B) Light

micrograph of the extremely thin trypanosome form in BIIT in vitro

culture, originally isolated from Q2088-2050. Scale bars represent 10

µm.

Figure 5.7 Light micrographs of (A) Initial adherent trypanosome form on the

surface of the erythrocyte (indicated by arrow) from quokka Q2088-

2050. (B) A more developed adherent trypanosome form on an

xxiii

erythrocyte in MSEM in vitro culture originally isolated from quokka

Q3336-3325 from Bald Island. (C) Immunofluorescent staining using

L8C4 monoclonal antibody of the adherent trypanosome form showing

green fluorescence of the flagellum and co-staining of nuclei (blue

fluorescence) using DAPI from quokka Q1. (D) FISH analysis of two

initial adherent trypanosome forms showing green fluorescence (white

arrows) adhered to the surface of an erythrocyte in vitro (Q1). (E) FISH

analysis of the more advanced adherent trypanosome form (white arrow)

on the surface of an erythrocyte in vivo isolated from quokka Q4633-

4613 from Two Peoples Bay. Scale bars represent 10 µm.

Figure 5.8 (A) The tiny trypanosome from in vitro culture, originally isolated from

Gilbert’s potoroo P83. Scale bars represent 10 µm. (B) SEM of the tiny

trypanosome form (indicated by arrow) from in vitro culture originally

isolated from Gilbert’s potoroo P83. Scale bars represent 2.5 µm.

Figure 5.9 (A) Immunofluorescent staining (L8C4) of the circular trypanosome

form in vitro showing green fluorescence (white arrows) of the flagellum

around the circumference of the parasite (from quokka Q1). (B) The

same form showing the nuclear region within the centre stained with

DAPI. Scale bars represent 10 µm.

Figure 6.1 Phylogenetic relationships of T. copemani genotype A and B quokka

isolates (underlined this study) from Two Peoples Bay (TPB), Bald

Island (BI) and Rottnest Island (RI) using the Bayesian Inference method

of partial (841 bp) 18S rRNA sequences. Posterior probabilities are

indicated on the main branches. Quokka isolates representing T.

copemani genotype A (#) and T. copemani genotype B (*) are listed.

Isolates designated ** were mixed T. copemani genotype A and B

xxiv

infections. The scale bar represents the proportion of base substitutions

per site.

Figure 6.2 Phylogenetic relationships of T. copemani genotype A and B quokka

isolates (underlined this study) from Two Peoples Bay (TPB), Bald

Island (BI) and Rottnest Island (RI) using the Bayesian Inference method

based on partial (641 bp) GAPDH sequences. Posterior probabilities are

indicated on the main branches. Quokka isolates representing T.

copemani genotype A (#) and T. copemani genotype B (*) are listed.

Isolates designated ** were mixed T. copemani genotype A and B

infections. The scale bar represents the proportion of base substitutions

per site.

Figure 7.1 Light micrograph of T. copemani in a Modified Wright stained blood

film from quokka Q2088. Scale bar represents 10 µm.

Figure 7.2 Mean log10-transformed count (plus half minimum detection level) of T.

copemani grown in cultures containing serum from 3 humans (HA, HB,

HC) and one horse (H0C). The trend line represents the mean log-

transformed count and markers () represent individual counts.

Figure 7.3 Numbers of trypanosomes / mL measured at several time points after

direct inoculation of infected quokka blood into HuMSEM and

HoMSEM.

Figure 7.4 Light micrographs of T. copemani grown in vitro, in a culture of blood

from the quokka, which had undergone the blood incubation infectivity

test. (A) Epimastigote (e) and trypomastigote (t) forms at day 10, in a

Modified Wright stained smear from HuMSEM. (B) Sphaeromastigotes

(s), epimastigotes and trypomastigote forms at day 10, in a Modified

Wright stained smear from HuMSEM. (C) Trypomastigote form of T.

xxv

copemani at day 18, grown in HuMSEM overlayered with RPMI at

37ºC. Scale bar represents 10 µm.

Figure 8.1 Nomarski interference-contrast micrographs of Eimeria oocysts isolated

from quokka faeces (quokka isolate Q2929-4443) from Two Peoples

Bay, Western Australia, resembling E. quokka based on morphological

characteristics (A and B) and (quokka isolate Q4124-4119) from Bald

Island, Western Australia, resembling E. setonicis based on

morphological characteristics (C).

Figure 8.2 Evolutionary relationships of Eimeria quokka, Eimeria setonicis and

Eimeria sp. inferred by distance analysis of partial (481bp) 18S rRNA

sequences. Percentage support (>50%) from 1,000 pseudoreplicates from

neighbor-joining analyses is indicated at the left of the support node.

Shaded isolates are from this study.

Figure 8.3 Evolutionary relationships of Eimeria quokka and Eimeria setonicis from

the quokka inferred by distance analysis of the mitochrondial

cytochrome oxidase gene (CO1) (426 bp). Percentage support (>50%)

from 1,000 pseudoreplicates from neighbor-joining analyses is indicated

at the left of the support node. Shaded isolates are from this study.

xxvi

LIST OF TABLES

Table 1.1 Taxonomic classification of Trypanosoma.

Table 2.1 Nucleotide sequences of primers used in PCR assays.

Table 3.1 A list of Gilbert’s potoroo and quokka isolates used in this study.

Table 3.2 The mean dimensions and standard error (SE) of morphological features

of Trypanosoma copemani isolated from blood stream forms from the

Gilbert’s potoroo (Potorous gilbertii) and quokka (Setonix brachyurus).

Table 4.1 A list of animal isolates and isolated tick species captured from Bald

Island and Two Peoples Bay used in this study.

Table 4.2 Mean dimensions and standard error (S.E.) of morphological features of

trypanosomes isolated from I. australiensis tick faeces from quokka

Q2322-2351, I. australiensis tick midgut from quokka Q2917-2918 and

T. copemani blood stream trypanosomes from quokka blood.

Table 5.1 A list of quokka and Gilbert’s potoroo isolates captured from the three

different geographical locations used in this study.

Table 6.1 A list of quokka isolates captured from the three different geographical

locations used in this study.

Table 6.2 Prevalence of T. copemani genotypes in quokka isolates from Two

Peoples Bay (TPB), Bald Island (BI) and Rottnest Island (RI) as

determined by PCR and sequencing of partial 18S rRNA fragments. The

95% confidence intervals are given in parenthesis. Mixed infections were

determined by cloning and sequencing of amplicons producing mixed

sequencing chromatograms.

Table 8.1 Quokka isolates in which faecal samples were collected from, for this

study.

xxvii

Table 8.2 The prevalence (and 95% confidence intervals) of Eimeria in quokkas

detected by both microscopy and PCR from three geographical locations.

Table 8.3 Morphometric (µm) comparison of Eimeria oocysts from quokkas* from

Two Peoples Bay, and from quokkas● from Bald Island with previously

published morphometrics for E. quokka and E. setonicis from Rottnest

Island quokkas (Barker et al., 1988). *(Q4125 - 4169), (Q4177 - 4185),

(Q4124 - 4119), ●(Q4546 - 4547), (Q2367-2332), (Q4527 - 4578),

(Q4547 - 4546), (Q2929 - 4443).

xxviii

CHAPTER 1: LITERATURE REVIEW

1

1.1 General history of trypanosomes

Fagellates of the genus Trypanosoma are ubiquitous haemoprotozoans that infect a wide range of animals and are the causative agents of several diseases of major social and economic impact around the world. The first described observation of a trypanosome was made by a German physician, G. Valentin, who in 1841 observed elongated motile organisms in the blood of a trout (Valentin, 1841). It has been speculated, however, that Antony Van Leeuwenhock, termed the father of protozoology, may have observed trypanosomes in 1680, with the discovery of organisms in the guts of horse-, believed to be the developing stages of Trypanosoma theileri (Hoare,

1972). Forty years after the discovery of trypanosomes, the pathogenic potential of these parasites was noted by veterinary officer Griffith Evans who discovered an organism in horses and camels from India suffering from a disease, surra. This disease- causing parasite, named Trypanosoma evansi in honour of G. Evans, led to subsequent trypanosome investigations, with Trypanosoma brucei brucei discovered as the causative agent of nagana in livestock and in wildlife by David Bruce in 1895 (Bruce,

1895). It was not until the early 1900s that trypanosomes infecting humans were discovered. Trypanosoma brucei gambiense was first described by Dutton in 1902, in the cerebrospinal fluid of a patient suffering from sleeping sickness (Dutton, 1902), while Trypanosoma brucei rhodesiense was identified as the aetiological agent of an acute form of sleeping sickness in Africa (Stephens and Fantham, 1910). Another human infective trypanosome, named Trypanosoma cruzi, which was distinct from its

African relatives, was identified in South America by a Brazilian scientist Carlos

Chagas in 1909 (Chagas, 1909). Chagas’ disease is a potentially life-threatening illness affecting around 6–7 million people worldwide, with most cases within Latin America

(WHO, 2015). Since the 1900s, numerous other trypanosome species have been identified worldwide, infecting all classes of including amphibians, reptiles,

2 fish, birds and mammals, with the majority appearing to cause no disease to their hosts.

Such examples include Trypanosoma rotatorium from frogs, Trypanosoma chelodina from tortoises, Trypanosoma sinipercae from fish, Trypanosoma avium from birds,

Trypanosoma lewisi from rodents, Trypanosoma pestanai from badgers, Trypanosoma thylacis from bandicoots and Trypanosoma binneyi from the platypus to mention only a few. To date relatively few Trypanosoma species have been isolated from native

Australian fauna and little is known of their epidemiology and impact. It has been suggested by Noyes et al. (1999) that the low detection rate of trypanosomes in native

Australian mammals may be contributed to the low parasitaemia typically observed in

Australian animals.

1.2 Taxonomy and Phylogeny

The term trypanosome is used when discussing the members of the genus

Trypanosoma, a genus of haemoflagellated protozoan parasites of the family

Trypanosomatidae, within the order (Levine et al., 1980; Stevens and

Brisse, 2004). The systematics of Trypanosoma and the revised classification of mammalian trypanosomes (Moreira et al., 2004) is outlined in Table 1.1.

Table 1.1 Taxonomic classification of Trypanosoma

Kingdom Eukaryota Phylum Euglenoza Class Kinetoplastida Order Family Trypanosomatidae Genus Trypanosoma

3

The kinetoplastids are a widespread group of flagellated protozoa known to parasitise virtually all animal taxa as well as plants and insects (Hoare, 1972).

Kinetoplastids include the trypanosomatid parasites, Trypanosoma brucei,

Trypanosoma cruzi and Leishmania, which are the causative agents of major human diseases (Hamilton et al., 2004). Kinetoplastids are and structurally possess conventional eukaryotic organelles, including endomembranes (nuclear membranes

Golgi apparatus, endoplasmic reticulum), cytoskeletal microtubules and a membrane- bound nucleus and mitochondria (Bastin et al., 2000). Unlike other eukaryotes, kinetoplastids only contain one extremely large and generally elongated mitochondrion per cell (Bastin et al., 2000; Stevens and Rambaut, 2001; De Souza, 2002). The mitochondrial DNA (mtDNA), which is distinct from the nuclear DNA, is condensed into one sub-structure, called the kinetoplast. This structure is the distinguishing feature of the order Kinetoplastida and is always found near the basal body located at the base of the flagellum (Bastin et al., 2000; Stevens and Rambaut, 2001; De Souza, 2002). The size and position of the kinetoplast in relation to the nucleus can vary among species and is of diagnostic value (Hoare, 1972). In T. cruzi for example, the kinetoplast appears as a slightly concave disk of 1.0 µm in length (De Souza, 2002), whereas in T. lewisi it appears elongated (Hoare, 1972). This unique organelle contains mtDNA termed kinetoplast DNA (kDNA) and is unlike that of any other DNA in nature (Morris et al., 2001). The kDNA is a network of thousands of interlocking DNA circles consisting of 25-50 maxicircles and 5000-27,000 minicircles (Morris et al., 2001).

Maxicircles are around 20-38 kb in length and encode ribosomal RNAs and proteins for mitochondrial bioenergetic processes, while minicircles (0.46-2.5 kb) encode guide

RNAs utilised in RNA editing (Vickerman, 1994).

4

1.3 General morphology of trypanosomes

Trypanosomes have been described as being polymorphic with various trypomastigote stages associated within the same species, for example, stumpy, slender and broad trypomastigote forms have been reported for T. cruzi (Brener, 1973). For ease of characterisation, Hoare (1972) described the trypomastigote form as the true trypanosome, on which all morphometrics are based. This stage is typically characterised as being lanceolate in shape with a flattened body, and tapers into a point towards the anterior end. The posterior end of the parasite can vary in shape but is generally broader than the anterior end and tapers to a round or blunt tip (Hoare, 1972).

A flagellum arising from near the kinetoplast and basal body is associated with an undulating membrane along the length of the cell and where it detaches from the anterior end is termed the free flagellum as characterised by Hoare (1972) and represented in Figure 1.1.

Figure 1.1. Morphological characteristics of the genus Trypanosoma.

In order for trypanosomes to move through the circulatory system, they must have a functional paraflagellar rod (PFR) within their flagellum. This complex lattice- like structure that runs alongside the flagellar sheath, enables the flagellum to move and is found only in free-living dinoflagellates, euglenoids and kinetoplastids (Bastin et al.,

1998; Kohl and Gull, 1998). The presence of this unique structure enables 5 trypanosomes to swim via their flagellum while dragging their cell bodies behind

(Bastin et al., 2000). The point of emergence of the flagellum is often utilized as a morphological distinguishing feature. This is made possible because trypanosomes pass through a variety of developmental stages during their life-cycle with the origin of the flagellum indicated by the position of the kinetoplast.

Seven distinct stages of Trypanosomatidae have been characterised by Hoare,

(1972) and are represented diagrammatically in Figure 1.2 and summarised as follows:

The amastigote stage [1] is described as a non-flagellated form, with a round or elongated appearance (Hoare, 1972). The sphaeromastigote [2], first described by Brack in 1968 (Brack, 1968), is typically rounded in appearance with a flagellum and represents a transitional stage between the amastigote and the epimastigote stage. The promastigote [3] is represented by an elongated body and typically is characterised by its anterior kinetoplast and an arising anterior flagellum. The opisthomastigote [4] is a stage only found in the subgenus Herpetomonas and is represented by an elongated form with a post-nuclear kinetoplast and a flagellum, which passes through the entire body and emerges from the anterior end. The epimastigote stage [5] is typically characterised by its juxtanuclear kinetoplast and has a flagellum that arises close to the kinetoplast and emerges from the side of the body. The trypomastigote [6] is known as the ‘true trypanosome’, which is generally observed within the circulatory system. This stage is elongated and has a post-nuclear kinetoplast and flagellum that emerges from the side of the body running along its surface or along an undulating membrane. The choanomastigote [7] resembles the shape of a barley corn, with a funnel shaped anterior reservoir from which emerges the flagellum. The kinetoplast is ante nuclear in this form and is restricted to the genus Crithidia (Hoare, 1972).

6

1

2a

7 2b 3 4 5 6

Figure 1.2. Morphological stages described in Trypanosomatidae: [1] amastigote stage; [2a] & [2b] sphaeromastigote stages; [3] promastigote stage; [4] opisthomastigote stage; [5] epimastigote stage; [6] trypomastigote stage; [7] choanomastigote stage (Hoare, 1972).

1.4 Salivarian and stercorarian mammalian trypanosomes

Species of Trypanosoma infecting mammals are subdivided into two groups; the salivarians and stercorarians and the groupings are dependent on where the infective stages develop within the vectors (Hoare, 1972). For ease of description, the African trypanosomes T. brucei ssp. group within the salivarian trypanosomes. This grouping is based on the metacyclic trypanosome’s anterior station development in the salivary glands of the vector with transmission of these parasites occurring via inoculation.

Trypanosoma cruzi, the South American trypanosome is typically stercorarian with the posterior station developmental cycle in the vector completed within the hindgut, with transmission being via faecal contamination. As with most classification systems however, there are always some species that show exceptions to the rules. Much debate on the taxonomic and evolutionary status of Trypanosoma rangeli (which is infective to humans but unlike T. cruzi is not pathogenic in humans), has confounded many parasitologists. Trypanosoma rangeli can be transmitted by both the anterior and

7 posterior end of a triatomine bug, with metacyclic trypanosomes developing at both sites. Trypanosoma rangeli is thought to be salivarian by some scientists and stercorarian by others; however it has been shown to have more characteristics similar to T. cruzi and phylogenetic analysis of the 18S ribosomal RNA (rRNA) gene showed that is closely related to South American isolates of T. cruzi and Trypanosoma dionisii, with both belonging to the stercorarian subgenus Schizotrypanum (Stevens et al.,

1999a).

1.4.1 The Salivarian trypanosomes

The salivarian trypanosomes (subgenera Duttonella, Nannomonas, Trypanozoon), include T. vivax, Trypanosoma congolense, and the T. brucei ssp. group. In general these trypanosomes are described as having a blunt posterior end with a terminal or subterminal kinetoplast with or without a free flagellum (Figure 1.3). Trypanosoma vivax is considered a Duttonella trypanosome, which is short in length ranging from 21-

25.4 µm with distinguishing features that include a ‘club shaped’ posterior end and a large terminally placed kinetoplast. These features alone are considered sufficient to separate Duttonella from other salivarian trypanosomes (Stevens and Rambaut, 2001).

The subgenus Nannomonas contains three recognised Trypanosoma species, T.simiae,

T. congolense and T. godfreyi. Trypanosomes within this subgenus typically lack a free flagellum and their undulating membranes are often inconspicuous (Hoare, 1972;

Stevens and Rambaut, 2001). Morphologically these species are indistinguishable, thus requiring molecular-based analysis to confirm the identity of these species (Hoare,

1972; Stevens et al., 1999a; Stevens and Rambaut, 2001). The subgenus Trypanozoon contains three species; the T. brucei group, T. equiperdum and T. evansi. These species are usually distinguished by differences in their epidemiology and pathology, as like the members of the Nannomonas group, all three species are morphologically indistinguishable. Infection with T. b. rhodesiense produces swollen lymph nodes

8 known as Winterbottom’s sign, while infection associated with T. b gambiense causes no glandular enlargement (Pentreath and Kennedy, 2004). Trypanosoma evansi is indistinguishable from the intermediate forms of T. brucei occurring as long-slender forms, ranging in length from 15-36 µm (Stevens and Brisse, 2004). Trypanosoma equiperdum is morphologicall indistinguishable from T. evansi, so diagnosis of this species is generally based on acute clinical signs which inclued subcutaneous swelling of the genital organs and presence of trypanosomes in the seminal fluid and viginal mucus of horses (Brun et al., 1998).

Figure 1.3. Micrograph of T. brucei in a blood film representing a typical salivarian trypanosome (http://www.cdc.gov/dpdx/).

1.4.2 The stercorarian trypanosomes

The stercorarian trypanosomes (subgenera Schizotrypanum, Megatrypanum,

Herpetosoma) are characterised by having typical pointed posterior ends with large non terminal kinetoplasts, and are always in possession of a free flagellum (Figure 1.4)

(Hoare, 1972). Reproduction of stercorarian trypanosomes within their mammalian hosts has been shown to be discontinuous (blood stream trypomastigotes do not divide) with division taking place in the amastigote or epimastigote stages as classically demonstrated in the T. cruzi life-cycle (Hoare, 1972; Vickerman, 1985). Trypanosoma cruzi is the most recognized and studied trypanosome species grouped within the

Schizotrypanum. To date T. cruzi is the only pathogenic species among the known

9 stercorarians and was responsible for infecting approximately 17-20 million people in

Central and South America in the 1990s (Fernandes and Andrews, 2012). Other recognised Schizotrypanum taxa also include species that appear to be restricted to bats such as T. vespertilionis, T. dionisii, T. hedricki and T. myoti. Morphologically

Schizotrypanum comprises small bloodstream forms with a mean size range between 14 to 24 µm, including the flagellum (Hoare, 1972). The bloodstream forms are distinctively curved with large kinetoplasts that have been described in detail by Hoare

(1972). In addition to humans, T. cruzi is known to parasitise a wide variety of mammalian hosts and has been identified in hundreds of animals from eight different orders including Artiodactyla, Carnivora, Chiroptera, Didelphimorphia, Perissodactyla,

Primates, Rodentia, and Xenarthra (Teixeira et al., 2011). Marsupials of the genus

Didelphis are considered the earliest hosts of T. cruzi in South America with the opossum (Dipelphis sp.) the most common reservoir of T. cruzi. Opossums are capable of maintaining amastigotes in their tissues and epimastigotes in their anal gland secretions, enabling environment contamination (Teixeira et al., 2011). Wild mammals such as the armadillo (Dasypus sp.), raccoon (Procyon lotor) and the wood rat

(Neotoma floridana), have also been reported to act as occasional reservoir hosts

(Herrera et al., 2015). With the advance of DNA technology, a phylogenetic study by

Stevens et al. (2001) demonstrated the close phylogenetic associations between T. cruzi,

T. rangeli, bat trypanosomes, trypanosomes from the South American mammals and a trypanosome from an Australian kangaroo (Macropus giganteus) (H25) (Stevens et al.,

2001). Based on this data, Stevens et al. (2001) suggested that the T. cruzi clade may have originated and evolved in isolated mammals from the South American and

Australian continents (Stevens and Rambaut, 2001). A recent study by Hamilton et al.

(2012) hypothesised that T. cruzi may have evolved from a broad range of bat

10 trypanosomes and that these trypanosomes successfully made the switch to other mammalian hosts in both the New and Old Worlds.

Figure 1.4. Micrograph of T. cruzi in a blood film, representing a typical stercorarian trypanosome (http://www.cdc.gov/dpdx/).

Hoare (1972) described the subgenus Megatrypanum as having typically large trypanosomes that multiply as epimastigotes within their mammalian hosts.

Morphologically their kinetoplast is situated close to the nucleus and far from the posterior end of the body (Hoare, 1972). The most common Megatrypanum is

Trypanosoma theileri, which infects domestic and wild ruminants worldwide. The mean length of T. theileri is approximately 60-70 µm, but it may occasionally reach 100 µm in length (Stevens and Brisse, 2004). The validity of the subgenera Herpetosoma and

Megatrypanum has been questioned and shown to be lacking in evolutionary and taxonomic relevance. (Stevens et al., 1999b), demonstrated that Megatrypanum infecting deer were more closely related to Trypanosoma cyclops isolated from a monkey (Macaca sp.) compared to other Megatrypanum spp. However, no new classification system has been defined to date and therefore the current terminology is still in use.

The subgenus Herpetosoma contains trypanosomes that are smaller than

Megatrypanum and are able to multiply as either epimastigote or amastigote forms

(Stevens et al., 2001). These trypanosomes are medium in size, have a slender curved 11 body with a pointed drawn out posterior end and a pronounced free flagellum. The kinetoplast is large and rod-shaped and lies nearer to the posterior extremity than to the nucleus (Hoare, 1972). The majority of Herpetosoma spp. parasitise rodents and all known species to date are said to be non-pathogenic (Hoare, 1972). Trypanosoma lewisi is grouped into this taxon and parasitises murids worldwide. Morphologically the body of T. lewisi is characteristically curved, with the posterior end drawn out to a point and a total mean length of 21-37 µm. It has also been described as having a slightly developed undulating membrane and a well-developed free flagellum (Hoare, 1972). The vector of

T. lewisi is the rat (Nosopsyllus fasciatus) (Molyneux, 1969b, a). Infections with T. lewisi are generally not of medical or veterinary significance. However, T. lewisi was isolated from a sick human infant in Malaysia by Johnson in 1933, where it was believed that rat inhabiting the child’s dwellings were responsible for transmission

(Stevens and Brisse, 2004). Since then, T. lewisi has been reported to infect humans on seven occasions (Lun et al., 2009).

1.5 Vectors of Trypanosomes

The epidemiology of diseases caused by Trypanosoma species are determined in part, by the ecology of the vector and the host-vector relationship that affects parasite transmission (Hoare, 1972). Transmission of trypanosomes is dependent on the effectiveness of the parasites in evading the host immune system and their ability to develop within the mammalian host (Hoare, 1972). Trypanosoma cruzi undergoes stercorarian transmission to mammalian hosts by the triatomine bug (Triatoma infestans) through faecal contamination. This type of transmission is a less effective mode of transmission and depends on infected droppings being deposited on vulnerable sections of the skin, such as the mucous membranes or abraded skin. The inoculative method, as seen in the saliavarian transmission of African trypanosomes, is a more effective mode ensuring that the metatrypanosomes are inoculated directly into the

12 bloodstream. However it has been reported that the infection rate in Glossina spp. is generally low requiring multiple bites for infection to occur compared to the higher infection rate of stercorarian trypanosomes (Hoare, 1972).

The majority of known vectors of mammalian trypanosomes are insects which are particularly associated with haematophagous species of the order Hemiptera (eg. triatome bugs), Diptera (eg. flies) and Siphonaptera (eg. fleas) (Hoare, 1972).

Trypanosome species transmitted by biting flies include such species as T. b. rhodesiense, T. b. gambiense, and T. grayi transmitted by the tsetse , while T. evansi, and T. vivax are transmitted by tabanids outside of Africa (Hoare, 1972; Jakes et al.,

2001b). Trypanosoma lewisi and T. microti, known to infect many species of rodents and voles respectively, are transmitted by the flea (Hoare, 1972; Smith et al., 2005).

Ticks, bat mites and leeches have also been identified as vectors of trypanosomes

(Mackerras, 1959; Lukes et al., 1997; Stevens et al., 1999a; Hamilton et al., 2005a;

Thekisoe et al., 2007). Various leech species have been shown to play a role in the transmission of several trypanosome species such as T. rotatorium, T. boissoni and T. triglae found in the blood of aquatic vertebrates (Lukes et al., 1997). Interestingly, leeches have been reported to transmit many aquatic trypanosomes in Australia. For example, in 1968 Richardson (Richardson, 1968) reported trypanosomes in a single haemadipsid leech caught in New South Wales (NSW), Australia. In 2005, Hamilton et al. described three genera of leeches (Chtonobdella, Micobdella and Philaemon) infected with trypanosomes isolated from NSW, Victoria and Queensland, Australia, using DNA analysis (Hamilton et al., 2005a). In contrast, the vectors for many non- aquatic Australian native trypanosome species have not been identified, as little is known of these Australian trypanosomes and their life-cycles. It has, however, been hypothesized that potential vectors such as the wombat flea (Lycopsylla nova) and kangaroo ticks may be playing a role in the transmission of novel Trypanosoma species

13 isolated from wombats (Vombatus ursinus) and kangaroos (Macropus giganteus)

(Noyes et al., 1999). A study by Mackerras (1959) also reported the tick as a potential vector for native Australian trypanosomes with motile trypanosomes detected within tick nymphs (Ixodes holocyclus) isolated from bandicoots infected with Trypanosoma thylacis. A novel trypanosome isolate, KG1 from Japan, was also identified in naturally infected ticks (Haemaphysalis hystricis) (Thekisoe et al., 2007). The tick has also been suggested as a potential vector for the transmission of T. binneyi, which infects the platypus (Ornithorhynchus anatinus), together with sandflies and leeches (Mackerras,

1959; Noyes et al., 1999; Jakes et al., 2001b).

Although trypanosome transmission is generally associated with vectors, other modes do occur as seen with T. cruzi and include vertical transmission from mother to child via breast feeding, blood contamination, blood transfusions and mechanical transmission (between host animals with infected anal glands, e.g. opossums) and contaminated mouthparts of biting flies and bats (Hamilton et al., 2012). Another unique mode of transmission is through copulation as seen with T. equiperdum, where trypanosomes are transmitted in the seminal fluid from stallions and in the vaginal mucus of mares.

1.6 Life-cycle of trypanosomes

Members of the genus Trypanosoma are digenetic parasites, with a life-cycle that involves two hosts (Hoare, 1972). As a rule, the geographical distribution of the parasites coincides with that of their vectors and is determined by both favorable ecological and climatic conditions (Hoare, 1972). Trypanosomes in the bloodstream of mammals are ingested by the insect vector during a blood meal. Once within the vector, trypanosomes undergo a cycle of development in the mid- and hindgut which results in the development of infective metacyclic trypomastigotes (Vickerman, 1985). Two types

14 of life-cycles exist and are dependent on whether the trypanosome species are salivarian or stercorarian.

1.6.1 Salivarian life-cycle

The salivarian life-cycle within the mammalian host (Figure 1.5) is initiated when metacyclic trypomastigotes are inoculated through the skin by an infective feeding fly (Glossina spp.) (Hoare, 1972). Once the trypanosomes become established, they then develop into long slender trypomastigotes capable of maintaining a bloodstream infection (Langousis and Hill, 2014). These slender parasites may sometimes penetrate blood vessel endothelium and multiply in the connective tissues and lymphatics by longitudinal binary fission (Vickerman, 1985). These trypomastigotes are subsequently carried to the heart and other organs and may at a later stage invade the central nervous system (Taylor and Authie, 2004). Once infection has become established within the mammalian host, trypanosomes circulating within the peripheral circulation are taken up by a feeding tsetse fly. Trypanosomes that reach the intestines of the fly reproduce in both the mid- and hindgut, generating large numbers approximately ten days after infection (Vickerman, 1985). When large numbers are present, procyclic trypanosomes migrate to the salivary glands via the pharynx and salivary ducts. Within the salivary glands, the trypanosomes metamorphose into epimastigote forms and within two to five days further develop into metacyclic trypanosomes (Vickerman, 1985). Once infective, the fly introduces the metacyclic trypanosomes into the next host through saliva injected into the puncture wound during feeding, thus continuing the cycle of infection.

15

Figure 1.5. Diagrammatical representation of a typical salivarian life-cycle.

1.6.2 Stercorarian life-cycle

1.6.2.1 Development within the mammalian host

Trypanosoma cruzi has a typical stercorarian life-cycle, shown in Figure 1.6.

Infection of a mammalian host is initiated when an infected triatomine bug deposits metacyclic trypomastigotes within its faeces onto either the skin or mucous membranes around the eye or mouth. The trypanosomes either penetrate the mucous membranes or are scratched into the bite wound, entering the circulatory system (Vickerman, 1985).

The trypomastigotes of T. cruzi in the peripheral blood represent pleomorphic life-cycle stages consisting of slender, stout and broad forms (Brener, 1973). The stout forms are able to circulate in the bloodstream for days without penetrating the host cells and have been shown to be more resilient to the host immune system than the more sensitive slender forms, which disappear from the circulation in one to two hours. Once within the circulatory system, trypomastigotes enter nucleated mammalian cells (especially macrophages, fibroblasts, Schwann cells, smooth and striated myocytes) via ‘parasite- directed endocytosis’, resulting in the formation of a parasite-containing endocytic vacuole, which fuses with lysosomes within the host cell (Tyler and Engman, 2001).

16

Here they replicate as amastigotes leading to the formation of pseudocysts or a

“parasitic nest” which is the grouping of trypanosomes within the cell’s cytoplasm

(Tyler and Engman, 2001; Navarro et al., 2003). Once they attain high numbers, amastigotes transform into trypomastigote forms that rupture the host cell and migrate into the blood and lymph re-invading new cells (Tanowitz et al., 1992). Trypomastigote forms that fail to invade new cells undergo morphological changes to broad forms and then to amastigote forms (Tyler and Engman, 2001). This pathway of differentiation allows amastigotes to propagate infection via the phagocytic cells, which ingest the amastigotes (Tyler and Engman, 2001). The life-cycle is completed when a new vector ingests blood containing trypomastigotes.

Figure 1.6. Diagrammatical representation of a typical stercorarian life-cycle.

1.6.2.2 Development within the arthropod vector

Infection of the vector occurs when the appropriate insect ingests a pleomorphic population of trypomastigotes in a blood meal from an infected host. Once inside the stomach of the insect, the bloodstream forms develop into epimastigote and amastigote forms (De Souza, 2002). The amastigotes are three to five µm in diameter and are able to replicate and to transform into epimastigotes. Initially the amastigotes swell and 17 extend their flagellum to become sphaeromastigotes (Tyler and Engman, 2001); they then continue to elongate giving rise to the classical epimastigote form. Epimastigotes hydrophobically attach to the hindgut wall prior to differentiation into the metacyclic forms via longitudinal binary fission (Tyler and Engman, 2001). Once formed, they detach from the gut wall and migrate to the rectum. Here the metacyclic trypanosomes await excretion, where they initiate infection upon penetration of a vertebrate host.

1.7 Clinical signs and pathogenesis of trypanosomiasis

1.7.1 Pathogenesis of salivarian trypanosomes

Infection with pathogenic salivarian trypanosomes causes three diseases, nagana

(caused by T. congolense, T. vivax, T. simiae, T. brucei and T. suis), surra (caused by T. evansi) and dourine (caused by T. equiperdum) (Hoare, 1972). In animal trypanosomiasis, two main stages occur: one involving an acute response and the other a chronic phase of the disease. During the early phase of infection, trypanosomes in the dermis undergo extensive multiplication with the development of oedema after inoculation. After a period of time trypanosomes are observed in both the lymphatic and circulatory systems. Initially their presence results in high fevers during the first peaks of parasitaemia and then fluctuating fever thereafter in accordance with ‘waves’ of parasitaemia (Uilenberg, 1998). Anaemia then follows and this is a prominent feature of animal trypanosomiasis associated with weakness, lethargy and loss of condition

(Taylor and Authie, 2004). The most common type of anaemia during the acute phase is haemolytic anaemia and is associated with the detection of microspherocyte, echinocyte, schistocyte and dacrocyte cells within the blood. Reticulocytes are also present and diagnostically these immature red blood cells indicate blood loss or decreased red blood cell survival, and are prematurely released from the bone marrow to compensate for the loss of blood (Rodak et al., 2012). An auto-immune response is also

18 typical in animal trypanosomiasis. High numbers of white blood cells including macrophages, neutrophils and eosinophils capable of destroying trypanosomes have been shown to increase phagocytosis of leucocytes, platelets and erythrocytes resulting in leucopenia and thrombocytopenia (Taylor and Authie, 2004; Mallah et al., 2010).

The exact mechanism of haemolytic anaemia in trypanosomiasis is unknown. It is believed that in the acute stage, erythrocytes become coated with antigens from lysed trypanosomes leading to an autoimmune response (Taylor and Authie, 2004). Uilenberg

(1998) proposed that anaemia caused by phagocytosis increases due to trypanosome proteins, leading to the haemolysis of erythrocytes. Initially the haemopoietic system attempts to compensate for the loss of erythrocytes by increasing erythropoietic activity, but in the chronic phase, other factors from the parasites have a depressing effect on the haemopoietic system (Uilenberg, 1998).

In association with anaemia, pathological changes involving the spleen, liver and central nervous system have also been reported (Taylor and Authie, 2004). A typical feature of acute phase trypanosomiasis is splenomegaly, a condition consistent with erythrocyte and lymphocyte sequestration (Uche and Jones, 1992; Taylor and

Authie, 2004). Uche and Jones (1992) observed engorgement and hyperplasia of the spleen in rabbits infected with T. evansi, as well as marked changes within the white and red pulp regions due to an increase in macrophages, lymphocytes and plasma cells within the sinuses. During the chronic stage of infection, neurological signs became apparent. Impairment of motor functions, and neurological disorders leading to paralysis are common, and animals suffering from dourine generally display polyneuritis and neuronal degenerative lesions of the central nervous system (Taylor and Authie, 2004).

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1.7.2 Pathogenesis of stercorarian trypanosomes

The most significant stercorarian of medical importance is T. cruzi, the aetiological agent of Chagas’ disease. Two clinically distinct phases of the disease occur, one involving an initial or acute phase, which generally lasts for four to eight weeks, and a chronic phase that persists for the host’s life span (Fernandes and

Andrews, 2012). An intermediate stage is also known to occur and this stage displays no signs of clinical disease but tests positive by haemoculture and/or xenodiagnosis

(Teixeira et al., 2011).

The first clinical sign of disease is associated with the inflammatory response of the skin, often resulting in either a chagoma or Romana’s sign due to the insect’s bite, either on the skin or the mucous membrane of the eye, respectively. After an incubation period of approximately 72 hours, trypanosomes begin to multiply with parasite numbers in the blood stream at their highest. Fever, headache, joint and muscle pain, anorexia, vomiting, diarrhoea, drowsiness, apathy, lymphadenopathy, hepatosplenomegaly, oedema and convulsions are often associated with this initial stage

(Teixeira et al., 2011). The heart is also typically infected with an increase in size, congestion, and dilation present. It is at this stage that patients may die due to cardiac failure or encephalitis due to extreme inflammatory pathologies, linked to parasitic nests in the heart and parasite-induced cytolysis (Bonney and Engman, 2008).

In the chronic phase, parasites are not usually detected directly in the blood, as an equilibrium is established between parasite and host. Cardio-myopathy, megaoesophagus and megacolon are the three main clinical manifestations of chronic

Chagas’ disease. Hypertrophy of the heart is present in 94.5% of infected individuals, with tachycardia and brachycardia leading to abnormal cardiac dilation, arrhythmias, conduction abnormalities, valvular disturbances and congestive heart failure (Teixeira et al., 2011).

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The occurrence of digestive abnormalities is less common, with enlargement of the oesophagus and colon generally seen in 5.5% of chronic sufferers (Teixeira et al.,

2011). In the digestive system, the muscle layers and intramural nerve plexus are predominantly affected, with parasites frequently observed in the connective tissues.

Such disturbances lead to hypertrophy and dilation of the intestinal tract disrupting normal peristalsis and often resulting in intestinal obstruction (Hoare, 1972).

1.7.3 Human pathogenic trypanosomes

Three trypanosome species, T. b. rhodesiense, T. b. gambiense and T. cruzi, are pathogenic to humans because they have evolved to resist the lytic effects of human serum (Schenkman et al., 1986; Lai et al., 2009). In 1978, Rifkin demonstrated that a high density lipoprotein (HDL) found in human serum was trypanolytic (Rifkin, 1978).

Milner et al. (1999) then showed that HDL contained a minor subclass of lipoproteins termed trypanosome lytic factors (TLFs) that confer innate protection against infection with T. b. brucei, and are found in the blood of humans and some primates (Milner and

Hajduk, 1999; Molina-Portela Mdel et al., 2005). Two distinct TLF complexes have been identified in human blood; TLF1, which is a 500 kDa high density lipoprotein complex composed of both lipids and seven different proteins, and TLF2, a 1000 kDa lipid-poor protein immunocomplex (Molina-Portela Mdel et al., 2005). Both TLF1 and

TLF2 consist of apolipoprotein L-1 (apoA -1) and a human haptoglobin-related protein

(Hpr) with the latter being linked to the trypanolytic activity. This is in contrast to apoA

-1 which appears to have no direct role in trypanosome lysis, given its undetectable levels in patients with Tangier disease, who are still able to maintain trypanosome lytic activity (Tomlinson et al., 1995; Tomlinson and Raper, 1998). TLF1 has been shown to damage the membrane of trypanosomes by forming cation-selective pores (Tomlinson and Raper, 1998). Normally trypanosomes actively maintain a low intracellular Na+ level and a high intracellular K+ level. However, with the presence of the cation-

21 selective pores the osmotic balance is disrupted and an increase in Na+ influx into the cell occurs. This increase in Na+ leads to passive water diffusion resulting in cytoplasmic vacuolization, cell swelling and eventual trypanosome lysis (Molina-

Portela Mdel et al., 2005).

Trypanosome resistance to TLF has been linked to the presence of a serum resistance associated gene (SRA), which is responsible for a cell surface protein present only in serum resistant isolates of T. b. rhodesiense (De Greef and Hamers, 1994; Xong et al., 1998; Milner and Hajduk, 1999). This gene is not found in T. b. gambiense, however a SRA-like gene that is specific to T. b. gambiense (TgsGP) has been shown to be essential for human serum resistance (Capewell et al., 2013).

The resistance of T. cruzi to human serum occurs via another pathway and is based on resistance to the alternative complement pathway (ACP) (Cestari and Ramirez,

2010). Trypomastigotes of T. cruzi resist lysis because various specific membrane glycoproteins participate in preventing efficient complement activation of the C3 convertase (the central enzyme of the complement cascade) on the parasite membrane

(Tomlinson and Raper, 1998). This process can occur either by inhibiting the assembly of the C3 convertase or accelerating its decay (Joiner et al., 1988; Cestari et al., 2008).

In contrast, the amastigote stage activates the alternative complement pathway, and resists lysis by preventing the insertion of the lytic membrane attack complex (MAC or

C5b-9) into its surface membrane (Iida et al., 1989). Interestingly, the amastigote stage also has the potential to resist lysis by antibodies in immune sera, potentially maintaining chronic infection (Tomlinson and Raper, 1998). The mechanisms of resistance are, however, not conferred to all of the life-cycle stages of T. cruzi

(Schenkman et al., 1986). The epimastigote life-cycle stage derived from the vector’s gut is efficiently lysed in human serum, as its survival in the mammalian host is not required (Tomlinson and Raper, 1998).

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A trypanosome infectivity test developed by Rickman and Robson (1970) was originally designed to distinguish the human-infective T. b. rhodesiense from the non- human-infective T. b. brucei without recourse to human experimentation (Rickman and

Robson, 1970). This test, named the blood incubation infectivity test (BIIT) is based on the observation that non-human-infective trypanosomes are lysed by human serum while human-infective trypanosomes resist lysis. In brief, this test is performed by inoculating rodents with the trypanosome isolate to be tested. Once positive, cardiac blood is withdrawn from the rat and incubated in human blood for five hours and then re-inoculated into another rat. A positive result for human infective isolates occurs when the second rodent presents persistent parasitaemia. Initially this test proved to be a useful tool in the search for the animal reservoir host of T. b. rhodesiense and was widely used with BIIT-positive trypanosomes isolated from a waterbuck, lion, hyena, and cattle in Tanzania (Geigy et al., 1971). However, four years later Geigy et al. (1975) reported problems associated with the test and showed that trypanosomes from the first peak of infection were more likely to give a negative BIIT result compared to later populations. Geigy et al. (1975) also demonstrated that plasma resistant trypanosomes existed within the blood and less than 20% of these resistant parasites could not be detected by the BIIT. This was also demonstrated by Hawking (1976), who showed that trypanosome stocks can be composed of both serum resistant and serum sensitive individuals, undermining the validity of the BIIT. However, several successful infections of volunteers with BIIT-positive stocks have been reported (Robson et al.,

1972; Geigy et al., 1975) and most Trypanozoon (West African Trypanosoma) stocks isolated from man are BIIT positive (Hawking, 1976). It should be noted however that not all positive results indicate human pathogenic trypanosomes, as many stocks of T. lewisi, T. vivax and T. congolense are highly resistant to human blood, but infection to man is generally rare (Hawking, 1976). In addition, certain stocks of T. brucei ssp. have

23 also been shown to be originally human serum sensitive but through constant passage and rapid selection have the ability to develop human serum resistance (Turner et al.,

2004).

1.7.4 Drug therapies and vaccines

Current treatment for T. b. gambiense and/or T. b. rhodesiense includes melarsoprol, eflornithine, pentamidine and suramin, all of which are costly, highly toxic and have lost efficacy in several regions (Barrett et al., 2007; Jacobs et al., 2011).

Suramin, developed in 1920, and pentamidine, developed in 1940, are used prior to central nervous system involvement for diseases of T. b. rhodesiense and T. b. gambiense, respectively (Barrett et al., 2004). During the late stage of disease, melarsoprol and eflornithine are used, but are highly toxic, with melarsoprol derived from arsenic (Goodman-Gilman et al., 1996). One recent study reported that drug combinations of diminazene aceturate (DA) with either levamisole and/or vitamin C were more effective in the treatment of rats infected with T. b. brucei than DA alone

(Chekwube et al., 2014). Another study demonstrated that curcuminoid analogs had potent activity against T. b. brucei (Alkhaldi et al., 2015) and cysteine-rich secretory proteins (CRISPs) found in snake venom also show promise for the development of novel agents against trypanosomiasis (Adade et al., 2014).

Treatment of Chagas’ disease (T. cruzi) is based on the use of nifurtimox and benznidazole, two very toxic nitroheterocyclic compounds (Urbina, 1999). Neither is effective against chronic forms of the disease and have modest efficacy against the acute form, with both drugs significantly toxic to the host (Barrett et al., 2004). No vaccines for trypanosome diseases currently exist due to the extensive antigenic variation of the trypanosome surface coat (La Greca and Magez, 2011). The variable surface glycoprotein (VSG) in the surface coat of the trypanosome is responsible for antigenic variation (Barry and Carrington, 2004). There are over 1,000 possible VSGs

24 and therefore it is thought that the development of a vaccine is highly unlikely (La

Greca and Magez, 2011). In the absence of effective treatments and vaccines for trypanosomes, a better understanding of the epidemiology and transmission dynamics of these parasites becomes even more important.

1.8 History of Australian trypanosomes

In contrast to the wealth of information on economically important species such as T. cruzi and T. brucei, little is known of the diversity, morphology and the life-cycles of Australian trypanosomes (Hamilton et al., 2004). The first record of Australian

Trypanosoma in mammals was made by T.L. Bancroft in 1888 (Bancroft, 1888).

Bancroft, while looking for microfilaria, discovered T. lewisi (Figure 1.7) in the blood of rats captured in the suburbs of Brisbane. Further reports of observations of T. lewisi were also noted by Pound in 1905 in Brisbane (Pound, 1905), Cleland in Perth (Cleland,

1906, 1908) and Johnston in 1909 in Sydney (Mackerras, 1959). Studies of murids by

Mackerras (1959) detected T. lewisi in six bush rats (Rattus fuscipes) and in one water rat (Hydromys chrysogaster) from Queensland. In vivo studies by Mackerras (1959) on experimentally infected young rats reported an acute phase of infection, with T. lewisi initially rapidly multiplying and then gradually diminishing in numbers. In the chronic phase they disappeared from the circulatory system altogether (Mackerras, 1959).

Figure 1.7. Micrograph of T. lewisi in a blood film (Verma et al., 2011).

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The health implications of trypanosomes introduced to immunologically naïve

Australian native hosts was demonstrated by T. lewisi which has been implicated as the sole cause of extinction of the Maclear’s rat (Rattus macleari) on Christmas Island

(located in the Indian Ocean 2600 km North West of Perth, Western Australia) (Wyatt et al., 2008). Trypanosoma lewisi was believed to have been introduced to the island by fleas hosted on stowaway black rats (Rattus rattus) (Wyatt et al., 2008). With the advent of molecular DNA technologies, samples from museum specimens during the extinction window (AD 1888–1908) from both rodent species were examined. Positive results for the presence of T. lewisi from both native and introduced rats were reported by Wyatt et al. (2008) confirming the pathological cause of the extinction of the Maclear’s rat in

Australia.

The next trypanosome species detected in Australia was a bat trypanosome. In

1913, Breinl discovered Trypanosoma pteropi infecting a flying fox (Pteropus sp.) from

Townsville, in northern Queensland (Breinl, 1913). Morphologically this trypanosome was described as having a slender body, an under-developed undulating membrane and a long free flagellum with a total length ranging between 20-22 µm (Breinl, 1913).

Similar trypanosomes were also identified in flying foxes (Pteropus gouldii) by

Mackerras (1959) except they were slightly smaller in length, ranging from 18-20 µm, had a sharply pointed anterior end and a rounded kinetoplast. Morphologically they were shown to be closely related to Trypanosoma vespertilionis from European bats and reported to possess T. cruzi-like characteristics (Mackerras, 1959). A broad form was also identified in the flying fox, however measurements were not recorded.

Trypanosoma hipposideri, another bat trypanosome species was identified in the dusky horseshoe-bat (Hipposideros ater) caught in northern Queensland (Mackerras, 1959).

This species was characterised as very small and slender with a large kinetoplast located

26 near the posterior end and a delicate short free flagellum at the anterior end and measured 10.5-13 µm in total length (Mackerras, 1959).

Trypanosomes in domestic animals were discovered by Wenyon in 1926 and later by Turner and Murnane in 1930 (Mackerras, 1959) in the blood of cattle (Bos taurus) from Melbourne abattoirs (Turner and Murnane, 1930). This trypanosome was named T. theileri and has been described as having both a large and small form. The large form measures between 60-70 µm in length and the small form between 25-30 µm and both have been shown to be non-pathogenic (Mackerras, 1959).

In 1933, Owen was the first person to detect trypanosomes in the Australian monotreme, the platypus (Owen, undated). He described the parasite as a long slender trypanosome containing a narrow undulating membrane. A trypanosome isolated from a platypus caught in Interlaken, Tasmania was later described by Mackerras (1959) as being large and broad with a well-defined undulating membrane with numerous waves resembling forms similar to aquatic reptile trypanosomes (Trypanosoma primeti from the water snake (Tropidonotus piscator) and T. grayi from the crocodile) (Mackerras,

1959). The platypus trypanosome ranged in length from 47-67 µm with a long narrow anterior end and a short free flagellum and was described and named T. binneyi (Figure

1.8) (Mackerras, 1959). The morphological differences between T. binneyi and the trypanosome isolated by Owen were noted by Mackerras (1959) and Hoare (1972).

Mackerras (1959) suggested that the trypanosome isolated by Owen, with its slender long form and narrow undulating membrane, may represent a distinct species. The exact systematics of the trypanosome observed by Owen, was however, undetermined (Hoare,

1972).

27

Figure 1.8. Micrograph of T. binneyi in a blood film isolated from the platypus (Hamilton et al., 2005).

Mackerras was the first to detect and morphologically characterise a

Trypanosoma species isolated from an Australian marsupial, the northern brown bandicoot (Isoodon macrourus, originally named Thylacis obesulus) (Mackerras et al.,

1953). Trypanosomes were observed in the blood of 12 northern brown bandicoots captured in Brisbane Australia. The parasites were described as being moderately broad, with a well-developed flagellum and undulating membrane. A rod shaped kinetoplast was described as being at a considerable distance from a long finely tapered posterior end. The trypanosomes measured in length between 22-23 µm and the species was given the name Trypanosoma thylacis after the northern brown bandicoot (Mackerras,

1959). A later studie by Bettiol et al. (1998) also detected trypanosomes in blood smears from both the eastern barred bandicoot (Perameles gunnii) and southern brown bandicoot (Isoodon obesulus) in Tasmania (Figure 1.9). The trypanosomes from both host species were shown to be morphologically distinct with 30 trypanosome isolates from the southern brown bandicoot having a mean length of 37.97 ± 4.34 µm, while 20 trypanosome isolates from the eastern barred bandicoot had a mean length of 37.97 ±

3.13 µm. The overall length and nucleus-kinetoplast distance were similar to T. thylacis,

28 however, measurements for breadth and posterior–kinetoplast had a high degree of variability (Bettiol et al., 1998).

Figure 1.9. Micrograph of a Trypanosoma sp. in a blood film isolated from the southern brown bandicoot (Bettiol et al., 1998).

Not long after the detection of trypanosomes from Tasmanian bandicoots, Noyes et al. (1999) detected for the first time, by in vitro culture, trypanosomes from the

Macropodidate and Vombatidae families. Blood was taken from an eastern grey kangaroo (Macropus giganteus) and a common wombat (Vombatus ursinus) from

Victoria, with trypanosome forms detected 13 and 20 days respectively after inoculation into modified sloppy evan’s medium (MSEM). Morphologically the kangaroo trypanosomes resembled promastigotes with a very short flagellum. Small very motile trypanosome forms measuring 5.5 µm x 1.6 µm, along with epimastigotes (12 µm x 1.8

µm) and long thin nectomonads (26 µm x 1 µm) were also detected. The wombat trypanosomes morphologically were very different with sphaeromastigotes and epimastigotes (27 µm x 1.6 µm, n = 20) the main forms detected in culture. Attempts to infect immunosuppressed mice with the culture isolates were reported as unsuccessful

(Noyes et al., 1999). To help characterise and determine the evolutionary history of these novel trypanosomes, a nested polymerase chain reaction (PCR), sequence and phylogenetic analysis of the 18S rRNA gene were conducted. Interestingly, the kangaroo and wombat isolates were shown to be genetically distinct, with the kangaroo

29 isolate most closely related to a T. cruzi isolate (Noyes et al., 1999), demonstrating that

T. cruzi-like isolates are present in Australia (Hamilton et al., 2005a).

Trypanosome infections in Australian reptiles were reported by (Jakes et al.,

2001a) who detected T. chelodina in the short-necked tortoise (Emydura signata), the saw-shelled tortoise (Elseya latisternum) and the eastern snake-necked tortoise

(Chelodina longicollis), all captured in Southeast Queensland. Trypanosoma binneyi was also detected in this study, in the peripheral blood stream of a platypus sampled in

Tasmania (Jakes et al., 2001b). Phylogenetic relationships based on a partial 18S rRNA gene sequence from T. chelodina, T. binneyi and other Trypanosoma spp. indicated that

T. chelodina, and T. binneyi did not share a close phylogenetic relationship with other trypanosomes from Australian mammals, reptiles or amphibians, but were instead found to be more closely related to fish trypanosomes and each other (Jakes et al., 2001b).

Hamilton et al. (2005a) isolated trypanosomes using the same in vitro culture methods as Noyes et al. (1999) from the blood of the common wombat (Vombatus ursinus) (Figure 1.10A), a swamp wallaby (Wallabia bicolor) (Figure 1.10B), and a currawong (Strepera sp.) all from Victoria, Australia (Figure 1.10C). In culture, the wombat trypanosomes resembled trypomastigotes with a well-developed undulating membrane and a large kinetoplast, while in contrast the wallaby trypanosomes lacked a distinguishable nuclear and kinetoplast region. The currawong isolate was described as having a prominent kinetoplast adjacent to the nucleus with some individuals observed as having a long free flagellum (up to 42 µm). Within the same study eight platypuses from the Yan Yean Reservoir (near Melbourne, Victoria) were also examined for the presence of trypanosomes. Blood film analysis detected five out of the eight animals as positive for T. binneyi, but attempts to grow the platypus isolates in culture failed.

30

Figure 1.10. Micrographs of trypanosome species isolated from Australian animals. (A) Wombat AAP isolate. (B) Wallaby ABF isolate. (C) Currawong isolate (Hamilton et al., 2005a).

Interestingly, evolutionary relationships using the 18S rRNA gene showed that the Australian trypanosome isolates from the wombat, swamp wallaby and currawong grouped into separate clades (Hamilton et al., 2005a). The wallaby trypanosome grouped with T. cyclops and the T. theileri clade, together with a trypanosome isolated from a monkey (Macaca sp.) in Malaysia. The wombat trypanosome grouped with T. pestanai isolated from a badger (Meles meles) from Portugal, while the currawong trypanosome grouped with trypanosomes isolated from blood sucking insects. In 2005,

Hamilton et al. (2005b) also detected trypanosomes in Australian rabbits captured from

Victoria and NSW. Morphological and molecular phylogenetic analyses identified the rabbit isolates as Trypanosoma nabiasi, a trypanosome previously reported in European rabbits. The mean total length was 25.1 µm, and the free flagellum 10.6 µm. These two morphological parameters were the only significant differences noted between the

Australian and European isolates with the European Trypanosoma nabiasi measuring slightly larger with a mean total length of 26.4 µm and a shorted free flagellum of 8.5

µm (Hamilton et al., 2005b).

Previously trypanosomes inffecting Australian native marsupials were only detected from Eastern Australia and Tasmania (Mackerras, 1959; Bettiol et al., 1998;

Noyes et al., 1999; Hamilton et al., 2005a). During a routine health assessment,

McConnell (2001 unpublished observations) detected a novel trypanosome species for the first time in the blood of the critically endangered Gilbert’s potoroo from Western

31

Australia. Five years later, Clark and Spencer (2006) reported a novel Trypanosoma sp. in quokkas captured from the mainland of Western Australia, however, no morphological measurements were made (Clark and Spencer, 2006).

The detection of the novel trypanosome isolate from the Gilbert’s potoroo led to a survey on Western Australian wildlife by Averis et al. (2009) who detected trypanosomes by PCR in the boodie (Bettongia lesueur), brushtailed possum

(Trichosurus vulpecular), golden bandicoot (Isoodon auratus), Shark Bay mouse

(Pseudomys fieldi), bushrat (Rattus fuscipes), ash-grey mouse (Pseudomys albocinereus), dibbler (Parantechimus apicalis), bettong or woylie (Bettongia penicillata) and common planigale (Planigale maculata). Genetic analysis at the 18S rRNA locus showed the bush rats to be infected with T. lewisi, while the dibbler and ash-grey mouse were shown to be infected with T. lewisi-like trypanosomes. Novel trypanosome genotypes were shown to infect the golden bandicoot, Shark Bay mouse, boodie, woylie and common planigale (Averis et al., 2009).

Characterisation of another novel native Australian trypanosome isolated from the koala (Phascolarctos cinereus) was undertaken by McInnes et al. (2009) and the species named Trypanosoma irwini (Figure 1.11). Morphologically this parasite measured 32.1-38.7 µm in length, and was described as having a prominent kinetoplast, a developed undulating membrane, long free flagellum and a pointed posterior end.

Genetically, at the 18S rRNA and glyceraldehyde-3-phosphate dehydrogenase

(GAPDH) loci, T. irwini was found to be most similar to trypanosomes from birds and closely related to T. bennetti from an American kestrel (Falco sparverius) (McInnes et al., 2009).

32

Figure 1.11. Micrograph of T. irwini in a blood film isolated from the koala (McInnes et al., 2009).

In 2010, McInnes et al. characterised another Australian koala trypanosome isolate named Trypanosoma gilletti (McInnes et al., 2010). Morphologically however, only one slender unique trypomastigote stage was identified in a blood smear from koala Barbie, that also tested positive for T. irwini. This unique parasite measured 41.7

µm in length and 1.2 µm in breadth. Evolutionary relationships at both the 18S and

GAPDH loci showed T. gilletti to be closely related to novel trypanosome species isolated from woylies and a chuditch from Western Australia, previously genotyped by

Averis et al. (2009). Two of the harbored mixed trypanosome infections, with two different trypanosome species detected. These co-infections of trypanosomes in koalas are the first reports of dual trypanosome species within Australian marsupials

(McInnes et al., 2010).

In Western Australia, common brushtail possums (Trichosurus vulpecula) captured within the south-west region were identified as harboring trypanosomes

(Paparini et al., 2011). A total of five trypomastigotes were measured in a blood film from one infected possum. Morphologically these trypanosomes were described as elongated in shape with an undulating membrane extending over 85% of the entire length of the parasite (Figure 1.12). The possum trypanosome was aslo described as having tapered ends, with its kinetoplast and nucleus situated nearer to the posterior end

33 and a free flagellum at the anterior. The average total length of trypanosomes in the possum isolate was 35.6 µm with a breadth of 5.5 µm. Genetically these possum isolates were characterised for the first time and were shown to be closely related to a kangaroo trypanosome species. H25, and a bush rat trypanosome, species BRA2, with a genetic distance of 0.2% at the 18S rRNA locus (Paparini et al., 2011). During the same study, woylie samples were also examined for the presence of trypanosomes by molecular analysis. Like the koalas, three woylie isolates were also identified as having mixed Trypanosoma infections. The woylie isolates grouped together with isolates from a chuditch and four other woylie isolates previously genotyped by Averis et al. (2009) and were shown to be most closely related to T. gilletti from koalas (McInnes et al.,

2010).

Figure 1.12. Micrograph of the novel trypanosome isolate in a blood film isolated from the common brushtail possum (Trichosurus vulpecula) (Paparini et al., 2011). In a follow up study to Averis et al. (2009), Thompson et al. (2013) identified a very small trypanosome from woylies and named it Trypanosoma vegrandis (Figure

1.13). Trypanosoma vegrandis was described as having a curved body with a drawn out pointed posterior end with the nucleus located near the anterior end and the kinetoplast situated close to the nucleus. These small trypanosomes had a total length ranging from

6.92 µm – 10.50 µm and breadth ranging from 1.00 µm – 1.63 µm. They had a

34 relatively long flagellum recorded as being 20% of the total length of the parasite

(Thompson et al., 2013).

Figure 1.13. Micrograph of T. vegrandis in blood films isolated from the woylie (Thompson et al., 2013).

1.9 Trypanosomiasis in Australian marsupials

Few studies on the effects of trypanosome infection on the health of Australian wildlife have been undertaken. Experimental infection of agile wallabies with T. evansi

(Macropus agilis) and dusky pademelons (Thylogale brunii) was undertaken by Reid et al. (2001), and resulted in a high morbidity and mortality in both marsupial species. The observed clinical signs at six days post-infection included anorexia, weakness and ataxia, with a variety of histological lesions including pericarditis, splenomegaly, ulcerative gastritis and enteritis (Reid et al., 2001). Mortality was high and generally occurred within 61 days of infection. From these results, Reid et al. (2001) concluded that if this species ever became established in Australia, it could have devastating effects on Australian native marsupials and production animals.

In 2008, the extinction of the Maclears rat (Rattus macleari) on Christmas Island was linked to the aetiological agent, T. lewisi. Sick and dying rats were observed and postmortem examinations reported a markedly enlarged spleen and enlarged superficial lymphatic glands (Durham, 1908).

35

Preliminary analysis of the clinical impact of trypanosomes on koalas indicated that infection with trypanosomes was significantly associated with indicators of koala ill-health and non-survival (McInnes et al., 2011). Koalas infected with trypanosomes were 1.4 times less likely to survive than non-infected koalas (p = 0.034) and trypanosome infections were also associated with reduced packed cell volumes (PCVs)

(p = 0.041) indicative of anemia and lower (p = 0.014) body condition scores than uninfected koalas (McInnes et al., 2011). There is also evidence that trypanosomes contribute to immunosuppression of koalas, and significantly different patterns of parasitaemia and/or pathogenicity have been reported in mixed trypanosome infections

1.10 Characterisation of trypanosomes

Traditional approaches using comparative morphology have proven to be of value for the identification of Trypanosoma spp., and have been the cornerstone for characterising trypanosomes (Hoare, 1972). The most important features for characterising trypanosomes are the morphological structures of the adult bloodstream trypomastigote form, determined by light microscopy. Various species may differ in the shape and size of their body, have differences in the position of the nucleus and kinetoplast, as well as differences in the development of their undulating membrane and free flagellum. The posterior end of trypanosomes is also of diagnostic value, either being sharply pointed as seen with Herpetosoma trypanosomes, or rounded as characterised by T. vivax, with its club-shaped posterior. These biological characteristics, along with a variety of measurable parameters, are the classification requirements used to help distinguish between trypanosome species (Hoare, 1972).

The use of blood films to detect trypanosomes within a host’s circulatory system has been widely applied for the detection of this parasite particularly within endemic regions due to its ease of application and cost-efficiency (Singh et al., 2004). However, the sensitivity of this technique is confounded by the fluctuating numbers of parasites in

36 the peripheral circulation and may lead to false negatives, particularly in chronic trypanosomiasis, where the parasite can no longer be detected within the bloodstream

(Singh et al., 2004). Other biological methods such as in vitro culture and serological tests (enzyme linked immuno-sorbent assay, agglutination, and indirect fluorescent antibody tests) can be utilised to increase the sensitivity and specificity for identification of Trypanosoma species (Desquesnes et al., 2002; Hide and Tait, 2004; Singh et al.,

2004). In vitro culture provides an effective methodology for the detection of trypanosomes and for instance, is commonly used to detect T. theodori in goats, which are rarely observed in blood films but are successfully detected using these methods. A study by Noyes et al. (1999) also used this method to detect Australian native trypanosomes isolated from both a common wombat and eastern grey kangaroo, using modified Sloppy Evans medium (MSEM). Trypanosomes were detected up to 20 days after the initiation of blood cultures from both host species. This methodology too has its limitations as no standard culturing method suits all trypanosome species (Noyes et al., 1999).

1.11 Molecular characterisation of trypanosomes

With the advent of DNA technologies, molecular analysis is now being commonly used in prevalence studies, and has been shown to be particularly useful when addressing problems relating to the identification of trypanosome species (Hide and Tait, 2004). Molecular analysis has provided the ability to accurately identify trypanosomes and facilitate the study of evolutionary or ecological processes within these parasites. It is also being utilised for diagnosis and detection of disease pathologies for trypanosomes, as recently demonstrated in the population decline of woylies due to trypanosomes (Botero et al., 2013; Thompson et al., 2014b). For a genetic marker to be successful, ideally it should be able to detect levels of variation within as well as between species of closely related organisms to help facilitate

37 differentiation (Constantine, 2003). The Polymerase Chain Reaction (PCR) has enabled the development of a wide range of molecular tools for genetic typing, characterisation and phylogenetic analysis (Monis et al., 2002). PCR was first described in 1988 (Saiki et al., 1988) and today nearly all molecular research involves some facet of this process.

The main advantage of PCR lies within its sensitivity, enabling results to be obtained from very small samples of DNA (Glick and Pasternak, 2003). It also enables the possibility of differentiating trypanosomes, which are morphologically similar but have very different economic impacts (Duvallet et al., 1999). Molecular tools are therefore imperative to our understanding of Trypanosoma, providing information on epidemiology, disease transmission, parasite ecology, and ultimately the prevention and treatment of parasitic disease.

1.12 Phylogenetic analysis

Phylogenetic analysis uses the sequences of specific gene loci to measure the extent of genetic divergence between organisms allowing the genetic history of a population to be inferred (Hide and Tait, 2004). The genetic structure of taxonomic units can be influenced significantly by the effects of genetic exchange and recombination between organisms (Hide and Tait, 2004). The integrity of any taxonomic group is maintained by the absence of gene flow between populations and it is therefore crucial to understand such mechanisms to fully interpret phylogenetic data.

The selection of a locus to resolve relationships between taxa of interest is therefore crucial and the locus chosen must be evolving at an appropriate rate to obtain feasible data (Monis et al., 2002). The evolutionary relationships of the order Kinetoplastida were deduced from the analysis of the nuclear 18S ribosomal RNAs (18S rRNAs)

(Fernandes et al., 1993) as depicted in Figure 1.14. The 18S rRNA gene locus has been the phylogenetic marker of choice because it is conserved throughout the eukaryotes and consists of both rapidly and slowly evolving regions (Stevens et al., 1998). The

38 rapidly evolving regions are useful for determining the difference between closely related species, while the slowly evolving regions help to determine more distant relationships (Sogin et al., 1986). The range of conserved and variable regions within this gene has allowed for the study of diverse rates of genetic evolution, thus facilitating the study of closely related species. The high copy number of the 18S rRNA gene makes it suitable for PCR amplification and the size range (1,500 - 2,000 bp) makes it ideal for sequence analysis (Stevens et al., 1998). However to safely characterise a species based on genetic information alone, the use of more than one locus is recommended. The use of the 18S rRNA data alone to describe a new species has been considered inappropriate due to intraspecies variation at this locus. It is therefore suggested that significant portions of this gene inclusive of the variable regions V7-V8 be used alongside another genetic marker. The glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene (a key enzyme in glycolysis), is known to be conserved within the Kinetoplastida and has been shown to be monophyletic even when considering only two to five Trypanosoma species (Alvarez et al., 1996). This gene has been utilised in many recent studies to further examine the phylogeny of trypanosomatids (Hamilton et al., 2004; Hamilton et al., 2005a; Hamilton et al., 2007) and shown to be a more reliable phylogenetic marker over the time scale in which trypanosomes appear to have diverged. The GAPDH gene is under a different set of evolutionary constraints compared to the 18S rRNA gene and is evolving at a slower rate, making this gene a more suitable marker for studying evolution over a large time scale. It has been suggested that greater than 65% of this gene be used when comparing

Trypanosoma evolutionary relationships. Together with the use of the GAPDH gene and the 18S rRNA gene, Hamilton et al. (2004) were able to determine that the much debated evolutionary origin of the genus Trypanosoma was monophyletic (arose from a single origin), which is in contrast to the paraphyletic origin reported by earlier studies

39 of the 18s rRNA gene alone (Gomez et al., 1991; Fernandes et al., 1993; Maslov and

Simpson, 1995).

Figure 1.14. Eukaryotic rDNA array showing the 18S rRNA gene in relation to the other RNA subunits. Two internal transcribed spacers (i.e. ITS-1 and ITS-2) separate the 18S, 5.8S and 28S genes.

1.13 Western Australian marsupials the Gilbert’s potoroo and quokka

The critically endangered Gilbert’s potoroo (Figure 1.15A) was the first native marsupial from Western Australia reported to be harboring trypanosomes (McConnell,

2001 unpublished observations). The Gilbert’s potoroo is a small rat kangaroo with dense grey-brown fur, large eyes and weighs around 1 kilogram. Discovered near

Albany in 1840 by John Gilbert, fossil records show that the Gilbert’s potoroo lived within a narrow range along the south coast of Western Australia (Sinclair et al., 1996).

In the early 1900s, this species was so rare that it was thought to be extinct, but was rediscovered in 1994 at Mount Gardner headland at Two People Bay, Nature Reserve, near Albany (Sinclair et al., 1996). With only around 40 individuals still in existence, translocation programs have been implemented. In 2005 a small number of animals were translocated to Bald Island, located off the coast of Albany, Western Australia, to help ensure the survival of the species in case of fire, disease and predation. Quokkas

(Figure 1.15B) also habitat Bald Island as well as the south west corner of Western

Australia and Rottnest Island. Like the potoroo, they are small marsupials similar in appearance to a wallaby and are endemic to Western Australia.

40

Figure 1.15a. Photograph of the Gilbert’s potoroo (Potorous gilbertii) captured by Dick Walker.

Figure 1.15b. Photograph of the quokka (Setonix brachyurus).

41

To date, little is known about the potential impact of pathogenic parasites on the health of the Gilbert’s potoroo and quokkas. A study by Barker et al. (1988) identified nine new species of the pathogenic coccidian Eimeria from the family Potoroidae.

These parasites are the etiological agents for enteric coccidiosis, generally a disease associated with stressed or young animals and responsible for causing chronic diarrhoea

(Bennett et al., 2006). Within Australia, Eimeria has been described from various marsupial hosts and these include kangaroos, wallabies, quokkas (Macropodidae), wombats (Vombatidae), possums (Phalangeridae) and bandicoots (Peramelidae),

(Mackerras, 1959; Barker et al., 1988, 1989; Heckscher et al., 1999; Power et al., 2009;

Hill et al., 2012; Yang et al., 2012). Characterisation of marsupial Eimeria has largely been based on morphological description of both the sporulated oocysts and sporocysts

(Mackerras, 1959; Barker et al., 1989; Duszynski and Wilber, 1997). However, some

Eimeria species are morphologically similar and can occur in multiple hosts, compromising species identification (Power et al., 2009). To aid in morphological identification, molecular analysis is providing a more robust taxonomy system, allowing evolutionary relationships between Eimeria species to be determined.

42

The specific aims of this thesis were to,

1. Characterise species of trypanosomes isolated from Australian native

marsupials;

2. Determine the vector of trypanosome species identified;

3. Identify the different life-cycle forms of Australian native trypanosomes;

4. Investigate the prevalence of Australian trypanosomes in quokkas from three

different geographical locations (Two Peoples Bay, Bald Island and Rottnest

Island) using molecular analysis;

5. Test the relative susceptibility of trypanosome species identified to human

serum; and

6. Characterise and determine the prevalence of Eimeria species isolated from

quokka populations from Two Peoples Bay, Bald Island and Rottnest Island

from Western Australia.

43

CHAPTER 2

GENERAL MATERIALS AND METHODS

44

2.1 Sample collection and geographical locations

Blood and faecal samples from quokkas and the Gilbert’s potoroo were collected from Two Peoples Bay (34º 58’S, 118º 11’E), Bald Island (34º 55’ S, 118º 27’E) and

Rottnest Island (32º 00’S, 115º 31’E) in Western Australia (Figure 2.1). The animals used in this study were either live-trapped using cage traps (Mascot Wire Works collapsible rat/bandicoot traps) baited with peanut paste-rolled oat mix, or netted by hand with long-handled nets. Animals were trapped under Murdoch University animal ethics permit W2204/09 and DEC permit number SC000767. The animals were anaesthetised with isoflurane and approximately 200 microliters (µL) of blood was collected by venepuncture of the lateral caudal vein. The blood was added to commercial blood storage tubes containing ethylene diamine tetra-acetic acid (EDTA)

(Sestet, Australia) and stored at 4°C for later use. Faecal matter was collected at sampling where possible (i.e. if the animal defaecated during the sampling period) and stored at 4ºC until required.

Figure 2.1. Map of Western Australia showing the geographical location of the three marsupial sampling sites

45

2.2 Collection of ectoparasites

Ticks were collected from quokkas and Gilbert’s potoroos and placed into 1.5 mL microcentrifuge tubes containing 70% ethanol. Samples were identified to the species level by Mr Russel Hobbs at Murdoch University with the aid of the Australian ticks key (Roberts, 1970).

2.3 Detection of trypanosomes

2.3.1 Preparation of blood film

Thin blood smears were made from whole blood, prepared by smearing a drop of blood onto a microscope slide, which was then air dried and stained with Modified

Wright stain, using an automated slide stainer (Ames Hema-Tek®, Bayer, Germany). A cover-slip was placed over the stained blood smear and the preparation was examined microscopically at 200× and 400× magnification to check for the presence of trypanosomes. Smears containing trypanosomes were then observed at 1000× magnification and images of the parasites recorded using an Olympus DP71 Advance digital camera.

2.3.2 In vitro cultivation of trypanosomes

In vitro cultures were established by pipetting 20 µL of fresh blood from each animal into 1.8 mL cryopreservation vials (Nunc Cryo tubes vials, Apogent, Denmark) containing horse blood and modified sloppy Evans medium (MSEM) as previously described by Noyes et al. (1999). Cultures were incubated in the dark at room temperature for 10-14 days. Microscopic examination at 200× and 400× magnification of wet-smear preparations of the medium from each culture was performed weekly after the initial 10-14 day incubation to detect motile trypanosomes. An aliquot of 200 L of medium was removed every 10 days and placed into a new culture vial containing fresh

MSEM. Once trypanosomes were detected, Giemsa-stained thin smears were prepared

46 for further microscopic examination. Parasites that were detected in MSEM were subcultured into Cunningham’s liquid medium (CM). A 100 L aliquot from each

MSEM culture containing trypanosomes was placed into each well of a 6 well culture plate (Nunc), containing 5 mL of Cunningham’s liquid medium (CM) (Cunningham,

1977), supplemented with 10% foetal calf serum and 10 mg/mL gentamycin. The cultures were maintained at room temperature and monitored daily using an inverted microscope. Cultures were passaged every week by transferring 1 mL aliquots of culture media to a new culture plate containing fresh CM, and incubated in the dark at room temperature (25-27°C) as previously described (Hamilton et al., 2005a).

2.4 Immunofluorescence

To confirm the existence of novel trypanosome forms, immunofluorescent labelling using an antibody specific to the paraflagellar rod (PFR), which is an organelle only found in free-living dinoflagellates, euglenoids and kinetoplastids (which includes

Crithidia, Trypanosoma and Leishmania), was used (Kohl and Gull, 1998). Three mL of CM containing trypanosomes were centrifuged at 1,000 × g for 5 min. The majority of the supernatant was removed leaving approximately 200 µL of medium to allow the pellet to be re-suspended. The re-suspended pellet containing trypanosomes was placed on a poly-L-lysine coated slide and air-dried at room temperature for 30 min. The cells were fixed by incubating at -20˚C in methanol for 2 hours. After fixation cells were re- hydrated in 100 mL of PBS for 5 min. The PBS was removed and the cells re-hydrated with fresh PBS for another 5 min. Two hundred µL of a primary monoclonal antibody

(L8C4, neat), which binds to PFR (supplied by Dr Derrick Robinson University of

Bordeaux, France and Professor Keith Gull, University of Oxford, England) was added to the air dried cells and incubated in a moist chamber for 60 min at room temperature.

The primary antibody was removed and the cells washed twice with 50 µL of PBS. A

20 µL aliquot of FITC rabbit anti-mouse IgG (Sigma) was then added to the cells and 47 incubated in a dark moist chamber for 60 min at room temperature. The cells were washed twice with 50 µL of PBS. A 50 µL aliquot of 4',6-diamidino-2-phenylindole

(DAPI; 1 g/mL) (a fluorescent DNA intercalating dye) was added to the same cells and incubated for 4 min. The DAPI was then removed and the cells washed at 30 sec intervals twice with 50 µL of 100 mM HEPES. A drop of equilibrium buffer (Slowfade,

Molecular probes, Invitrogen, USA) was added to the cells and incubated at room temperature for 4 min. The buffer was removed and a drop of Component A (Slowfade,

Molecular probes, Invitrogen, USA) was added for fluorescence stability. Finally a cover slip was applied, the air expelled and the edges sealed with nail varnish.

2.5 Scanning electron microscopy (SEM)

A 100 µL aliquot of cultured trypanosomes was placed into 500 µL of 5% glutaraldehyde and incubated overnight at 4 ºC. The fixed trypanosome cells were washed in 1× PBS and placed onto a round coverslip (G401-10, Pro SciTech,

Townville, QLD) coated with 10% poly-L-lysine and air-dried. A 1% solution of

Dalton’s Chrome OsO4 (Pro Sci Tech Townville, QLD) was placed over the cells and incubated in the refrigerator for 1.5 hours. The cells were then dehydrated by a series of ethanol washes with 5 min intervals. The cells were first washed several times with 30% alcohol followed by 2 washes with 50%, 2 washes with 70%, 2 washes with 80%, 2 washes with 90%, 2 washes with 95% and then 3 final washes with 100% ethanol for 10 min each. After dehydration the cells were washed in 50:50 ethanol:amyl-acetate for 30 min, then washed twice in 100% amyl-acetate over a 60 min time period. The coverslip was then dried in a critical point dryer and then dry mounted onto a specimen stub with a carbon disc and sputler-coated with gold. The stub was stored in a desiccator until required.

48

2.6 Morphological analysis of trypanosomes

Digital micrographs of trypanosomes in blood films and in vitro cultures were taken at 1000× magnification using OPTIMUS 5.2 image analysis software. A digital image was captured and reproduced on a computer and the measurements recorded in

µm at 1000× magnification. Measurements of six key morphological features were performed using Image Pro Express version 5.1 (Media Cybernetics Inc., USA): total length (length of body measured along the mid-line including free flagellum), breadth

(maximum breadth measured at the level of the nucleus (including undulating membrane), PK (distance between the posterior end and the kinetoplast), KN (Distance between the kinetoplast and posterior edge of the nucleus), NA (distance between the anterior edge of the nucleus and the anterior end of the body) and FF (length of the free flagellum) (Figure 2.2). The morphological features used were based on parameters described by Mackerras (1959) and Hoare (1972). The mean measurement of each feature was calculated and the statistical significance of any difference between trypanosomes from different host species was determined using a one-way analysis of variance (ANOVA) and Tukey’s Honestly Significant Difference test at a 95% confidence limit.

Figure 2.2. Measurable morphological diagnostic parameters of trypanosomes (Hoare, 1972).

49

2.7 DNA analysis of trypanosomes

2.7.1 DNA extraction

Whole genomic DNA was extracted from both fresh blood samples and cultured trypanosomes using either a MasterPureTM DNA Purification Kit (Epicentre®

Biotechnologies, Madison, Wisconsin, U.S.A.) following the manufacturer’s instructions or a QIAamp®DNA Blood Mini Kit (Qiagen, Germany) following the manufacturer’s instructions. The DNA was stored at -20 ºC until required.

2.7.2 DNA amplification of the 18S rRNA gene

Fragments of trypanosome 18S rRNA gene were amplified using nested polymerase chain reactions (PCR). The fragments partially overlapped and covered an approximately 1.5 kb of the 18S rRNA gene. Both nested PCRs used external reverse primer S-762 and external forward primer S-LF as previously described by Maslov et al. (1996) and McInnes et al. (2009) respectively. One nested PCR used internal forward primer S-823 and internal reverse primer S-66R (producing ~900bp) and the other PCR used internal forward primer S-825 and internal reverse primer S-LIR

(producing ~959bp) as previously described by Maslov et al. (1996) and McInnes et al.

(2009) (Table 2.1). PCR amplification was performed in a 25 μL volume with the final mix containing 10-50 ng of trypanosome DNA, 0.8 µM of each primer, 0.02 U/µl Kapa

Taq (Kapa Biosystems, Massachusetts, USA), a final concentration of 0.4 mM of each dNTP and 2.5 µL of 10× buffer (Kapa Biosystems, Massachusetts, USA) with 1.5 mM

Mg (Kapa Biosystems, Massachusetts, USA) and Baxter Ultra-Pure H2O to a final volume of 25 µl. An initial activation PCR step was run at 94°C for 5 min, 50°C for 2 min and 72°C for 4 min (Jefferies et al., 2006), followed by 35 cycles of 94°C for 30 s,

55°C for 30 s and 72°C for 60 s. The PCR was completed with a final extension of 7

50 min at 72°C. One µL aliquot of the PCR mixture from the first PCR reaction was used as the template for the second PCR reaction.

2.7.3 DNA amplification of the GAPDH gene

An ~ 841bp fragment of the GAPDH gene was amplified by semi-nested PCR using primers previously described (Hamilton et al., 2004; McInnes et al., 2009). PCR amplification was performed in a 25 μL volume with the final mix containing 10-50 ng of trypanosome DNA, 0.8 µM of each primer, 0.02 U/µl Kapa Taq (Kapa Biosystems,

Massachusetts, USA), a final concentration of 0.4 mM of each dNTP and 2.5 µL of 10× buffer (Kapa Biosystems, Massachusetts, USA) with 1.5 mM Mg (Kapa Biosystems,

Massachusetts, USA) and Baxter Ultra-Pure H2O to a final volume of 25 µl. Cycling conditions consisted of a denaturation step of 94°C for 5 min, 50°C for 2 min and 72°C for 4 min, followed by 35 cycles of 94°C for 30 s, 50°C for 30 s (primary PCR) or 52°C for 30s (secondary PCR) with an extension step of 72°C for 2.20 min, followed by a final extension step of 7 min at 72°C. A 1µL aliquot of the PCR mixture from the first

PCR reaction was used as the template for the second PCR reaction.

2.7.4 PCR controls and agarose gel electrophoresis

All PCR amplifications performed included a negative control consisting of sterile molecular-grade H2O. The positive control for PCR reactions consisted of genomic DNA preparations of trypanosomes identified (and sequenced) during previous analyses. PCR amplification products were electrophoresed on a 1% agarose gel

(Promega, Australia) with a 100 bp DNA ladder (Fisher Biotec, Australia) and products detected using SYBR Safe DNA stain (Invitrogen, USA) on a UV transilluminator

(Fisher Biotec, Australia).

51

2.7.5 Purification of PCR products

Under UV illumination, amplified PCR products were visualised to determine whether single or multiple PCR products were present. PCR reactions containing one product were purified using a QIAquick® PCR Purification kit (Qiagen, Germany).

When more than one band was present, bands of the appropriate size were excised from the gel using a sterile scalpel blade for each band to prevent cross contamination and purified in an in-house filter tip method and used for sequencing without any further purification as previously described (Yang et al., 2013). DNA was stored at –20ºC for further use. DNA was either sequenced directly or cloned into a plasmid vector

(pGEM®-T, Promega).

2.8 Cloning

Two 10 µL ligation reactions were performed with two different concentrations of DNA. The first reaction was with a 1:1 insert:vector volume ratio, and the other reaction with at a 3:1 insert:vector volume ratio. 5 µL of 2X Rapid Ligation Buffer, 1

µL of T4 DNA ligase, and 1 µL of pGEM®-T Easy vector (Promega, USA) was in each reaction and deionised H2O added to a final volume of 10 µL. The ligation reaction was incubated overnight at 4ºC.

A vial of JM109 competent cells (Promega, USA) was thawed on ice for 5 min.

50 µL aliquots of competent cells were placed into 1.5 mL microcentrifuge tubes and placed on ice for 20 min. The cells were then incubated for 45-50 sec in a 42ºC water bath and then chilled on ice for 2 min. 800 µL of pre-warmed SOC medium was then added to the cells and incubated at 37 ºC for 1.5 hrs with shaking (~150 rpm). 100 µL of the transformed culture was then plated onto 2× YT agar containing ampicillin

(0.1mg/mL), IPTG (100mM) and X-GAL (50mg/mL) and incubated overnight at 37 ºC.

Ten colonies from each culture plate were chosen for PCR validation. White colonies were picked with a sterile P100 µL pipette tip and inoculated into 50 µL of 52 sterile H20, boiled for 5 min, and centrifuged (2000 × g, 5 min). The resulting supernatant was used for PCR amplification.

PCR was performed to detect the presence of the 18S rRNA or GAPDH DNA inserts in the plasmids using the appropriate PCR conditions. All PCR reactions were carried out using 0.2 mL tubes in a Perkin Elmer GeneAmp 2400 Thermocycler. The primer nucleotide sequences are listed below in Table 2.1. Ten µL of amplified product was electrophoresed on a 1.5% agarose gel to detect the presence of inserts of the correct size using a 100 bp DNA ladder (Fisher Biotec, Australia). The rest of each colony of the positive transformant was picked with a sterile P200 pipette tip and inoculated into 20 mL of LB-AMP broth. Cultures were incubated overnight at 37 ºC with shaking. Following overnight incubation, 400 µL of each culture was added to 600

µL of 40% glycerol stock solution, mixed thoroughly and stored at -80 ºC until required. Plasmid clones containing the correct size insert were purified using a

QIAquick® PCR Purification kit (Qiagen, Germany). Purified plasmid DNA was sequenced using SP6 forward promoter primer (TATTTAGGTGACACTATAG) and

T7 reverse promoter primer (TAATACGACTCACTATAGGG) as described in section

2.9.

53

Table 2.1. Nucleotide sequences of primers used in PCR assays

Product Primer Nucleotide sequence (5’- 3’) length Reference (bp)

18S rRNA Forward 20 McInnes et gene S-LF 5'GCTTGTTTCAAGGACTTAGC3' al., 2009 fragment 1 18S rRNA Reverse 20 Maslov et gene S-762R 5'GACTTTTGCTTCCTCTAATG3' al., 1996 fragment 1

18S rRNA Forward 20 Maslov et gene S823F 5'CGAACAACTGCCCTATCAGC3' al., 1996 fragment 2a Reverse 20 Maslov et 5'GACTACAATGGTCTCTAATC3' S662R al., 1996 18S rRNA Forward 20 Maslov et gene S825F 5'ACCGTTTCGGCTTTTGTTGG3' al., 1996 fragment 2b Reverse 19 McInnes et 5'ACATTGTAGTGCGCGTGTC3' SLIR al., 2009 GAPDH Forward 5’CTYMTCGGNAMKGAGATYG 22 McInnes et gene GAPDH AYG3' al., 2009 fragment 1 Reverse 20 McInnes et GAPDH 5’GRTKSGARTADCCCCACTCG3' al., 2009

GAPDH Reverse 20 Hamilton gene G4a 5’GTTYTGCAGSGTCGCCTTGG3' et al., 2004 fragment 2

2.9 Sequencing

Approximately 10 ng of purified PCR fragments (cloned and uncloned) were sequenced using the ABI Prism BigDye® terminator cycle sequencing kit (Applied

Biosystems, CA, USA). Each sequencing reaction contained 4 µL of Big dye (Applied

Biosystems, CA, USA), 2 µL of forward or reverse primer (3.25 pmol/µL) and 4 uL of template DNA. Cycling conditions consisted of 96°C for 2 min, followed by 25-35 cycles of 96°C for 10 s, 50°C for 5 s and 60°C for 4 min. Sequencing reactions were purified using ethanol precipitation by adding 25 µL of 100% ethanol, 1 µL of 125 mM

54

EDTA and 1 µL of 3M sodium acetate (pH 5.6). The solution was mixed and incubated at room temperature for 20 min then centrifuged at 2000 × g for 30 min. The supernatant was removed and 125 µL of 80% ethanol was added to the tube and centrifuged at 2000 × g for 5 min. Ethanol was carefully removed from the tube to avoid disturbing the DNA pellet. The tube was placed in a Nalgene vacuum desiccator until the pellet was completely dried. Purified sequencing reactions were sequenced on an

Applied Biosystems 3730 DNA Analyser instrument.

2.10 Phylogenetic analysis

Phylogenetic trees were constructed for Trypanosoma sp. for the 18S rDNA and

GAPDH genes with additional sequences retrieved from GenBank. Sequence data was analysed using the FinchTV 1.4 software

(http://www.geospiza.com/Products/finchtv.shtml) and imported into Bioedit Sequence

Alignment Editor (Hall, 1999), for manipulations and alignment by CLUSTAL W

(Larkin et al., 2007). MEGA v 4 (http://www.megasoftware.net/) (Tamura et al., 2007) was used for maximum-likelihood (ML) and parsimony analyses. Distance estimation was conducted using TREECONv1.3b (Van de Peer and De Wachter, 1994) based on evolutionary distances calculated with either the Kimura or Tajima and Nei models and trees constructed using neighbour-joining. Statistical support was provided for the ML, distance and parsimony trees by bootstrapping 1,000 replicates. Bootstrap values greater than 70% were considered statistically significant. Selected species were used as an outgroup species according to the gene locus used.

55

CHAPTER 3:

MORPHOLOGICAL AND MOLECULAR

CHARACTERISATION OF TRYPANOSOMA

COPEMANI (TRYPANOSOMATIDAE) ISOLATED

FROM GILBERT’S POTOROO (POTOROUS

GILBERTII) AND QUOKKA (SETONIX

BRACHYURUS)

Austen, J.M., Jefferies, R., Friend, J.A., Ryan, U., Adams, P., Reid, S.A., 2009.

Morphological and molecular characterisation of Trypanosoma copemani n. sp.

(Trypanosomatidae) isolated from Gilbert's potoroo (Potorous gilbertii) and quokka

(Setonix brachyurus). Parasitology 136, 783-792.

56

3.1 Introduction

To date only a few Trypanosoma species have been isolated from Australian native fauna. The first record of Australian trypanosomes in mammals was made by

T.L. Bancroft in 1888 with the discovery of T. lewisi in rats (Bancroft, 1888). Since then, a number of other species have been identified including: T. pteropi from the flying fox (Pteropus sp.), T. hipposideri from the dusky horseshoe-bat (Hipposideros ater) (Mackerras, 1959), T. binneyi from the platypus (Ornithorhynchus anatinus)

(McMillian and Bancroft, 1974), T. thylacis from the northern brown bandicoot

(Isoodon macrourus) (Mackerras, 1959; Mackerras and Mackerras, 1960), T. chelodina from the short-necked tortoise (Emydura signata), the saw-shelled tortoise (Elseya latisternum) and the eastern snake-necked tortoise (Chelodina longicollis) (Jakes et al.,

2001b) and more recently novel Trypanosoma spp. have been identified from the southern brown bandicoot (Isoodon obesulus) (Bettiol et al., 1998), the eastern grey kangaroo, common wombat (Noyes et al., 1999) and swamp wallaby (Wallabia bicolor)

(Hamilton et al., 2004).

In 2001 an unidentified Trypanosoma sp. was detected by light microscopy within blood films of Australia’s most endangered marsupial, the Gilbert’s potoroo during haematological assessment of the health status of the animals (McConnell, 2001 unpublished observations). The Gilbert’s potoroo was abundant in the 19th century in the vicinity of King George’s Sound near Albany, Western Australia. By the early

1900s the species had declined dramatically and was thought to be extinct until its rediscovery in 1994 on the Mount Gardner headland at Two Peoples Bay Nature

Reserve, near Albany (Sinclair et al., 1996). The discovery of trypanosomes in the

Gilbert’s potoroo prompted an investigation into the prevalence and molecular characterisation of this novel trypanosome in the Gilbert’s potoroo and the quokka and the species name Trypanosoma copemani is proposed.

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3.2 Materials and Methods

3.2.1 Study site and sample collection

A total of seven Gilbert’s potoroos and three quokkas, were trapped at Two

Peoples Bay (34° 58’S, 118°11’E) near Albany, Western Australia during the month of

November (Table 3.1) Ectoparasites including 19 fleas and 13 ticks were collected from the Gilbert’s potoroo and placed into 1.5 ml microcentrifuge tubes containing 70% ethanol. Flea and tick samples were identified to the species level by Mr Russle Hobbs at Murdoch University with the aid of the Dunnet and Mardon (1974) key and the

Australian ticks identified key (Roberts, 1970), respectively.

Table 3.1. A list of Gilbert’s potoroo and quokka isolates used in this study.

Animal identification Geographical location Host code (ear tags) animal captured

Gilbert’s potoroo P49 Two Peoples Bay Gilbert’s potoroo P50 Two Peoples Bay Gilbert’s potoroo P61 Two Peoples Bay Gilbert’s potoroo P63 Two Peoples Bay Gilbert’s potoroo P72 Two Peoples Bay Gilbert’s potoroo P83 Two Peoples Bay Gilbert’s potoroo P94 Two Peoples Bay Quokka Q3 (1416-1407) Two Peoples Bay Quokka Q5 (1837-1464) Two Peoples Bay Quokka Q10 (1354-1368) Two Peoples Bay

3.2.2 Detection of trypanosomes in whole blood

3.2.2.1 Preparation of blood films

Wet-smear preparations of the buffy coat from each potoroo blood sample were made by cutting blood-filled heparinised capillary tubes (after centrifugation at 12,000

× g for 12 min), approximately 1-2 mm below the red cell/buffy coat interface and expelling the buffy coat onto a glass microscope slide. A coverslip was placed over the

58 buffy coat and the preparation examined microscopically under 200× magnification for the presence of motile trypanosomes.

Light microscopy and staining was conducted as described in chapter 2 (section

2.3.1). Morphometric analysis and in vitro culture was conducted as described in chapter 2 (sections 2.6 and 2.3.2 respectively).

3.2.3 Genetic characterisation and phylogenetic analysis

DNA extraction and PCR were conducted as described in chapter 2 (sections

2.7.1 and 2.7.2).

The partial 18S rRNA gene sequences amplified from the Gilbert’s potoroo and quokka were aligned with sequences from 32 other trypanosome species/genotypes obtained from the GenBankTM database that represented each of the major clades described previously by Hamilton et al. (2007) (refer to Figures 3.4 and 3.5 for

GenBank accession numbers). The final alignment consisted of 1,165 characters of which 527 were variable and 395 were parsimony informative and was used for subsequent phylogenetic analysis and further aligned manually by eye.

Phylogenetic relationships were determined using distance, maximum likelihood and parsimony based methods using MEGA 4 (Tamura et al., 2007) and statistical support was provided by using 1,000 bootstrap replicates. Maximum likelihood analysis was conducted using PhyML (Guindon and Gascuel, 2003) based on the HKY85 nucleotide substitution model and using 500 bootstrap replicates. All trees included

Phytomonas serpens (AF016323), Leptomonas sp. (EF546786) and Leishmania tarentolae (X53916) as outgroup species.

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3.3 Results

3.3.1 Microscopy

Using light microscopy, trypanosomes were detected in the blood of 7 Gilbert’s potoroos (7/7), and three quokkas (3/3). Trypomastigotes in various developmental stages were observed in blood smears from the Gilbert’s potoroo (Figure 3.1 A-D), and only one trypomastigote-like form was observed in blood smears from the quokka

(Figure 3.2 A). The morphology of the trypanosomes detected in blood smears and buffy coat smears from the Gilbert’s potoroo were consistent with trypomastigote life- cycle stages (Hoare 1972). Three main trypomastigote-like forms were observed, representing a slender form, a medium form and a broad form (Figure 3.1 A-C). The broad form possessed myonemes, which gave this form a striated appearance (Figure

3.1 C). In all three forms, the kinetoplast was observed at some distance apart from the nucleus and generally situated closer to the posterior end. The trypomastigote-like stages generally had a pointed posterior end, a well-developed undulating membrane and a free flagellum. The dimensions of representative trypomastigotes are presented in

Table 3.2. Piroplasms and microfilariae (Figure 3. 2 B) were also observed in the potoroo blood smears (7/7 and 2/7 respectively). The fleas collected from the Gilbert’s potoroo were identified as Stephanocircus dasyuii of the family Stephanocircidae and the ticks were identified as Ixodes australiensis of the family Ixodidae.

60

Figure 3.1. Light micrographs of Trypanosoma copemani isolated from the blood of a Gilbert’s Potoroo. (A) Slender trypomastigote form in a Modified Wright stained blood film. (B) Medium trypomastigote form in a Modified Wright stained blood film. (C) Broad trypomastigote form in a Modified Wright stained blood film. (D) Dividing trypanosome form in a Modified Wright stained blood film. Scale bars represent 10 µm.

Figure 3.2. Light micrographs of Trypanosoma copemani isolated from the blood of a quokka and haemoparasites isolated from the Gilbert’s potoroo and quokka. (A) Trypomastigote form in a Modified Wright stained blood film from a quokka and a piroplasm (indicated by the arrow) in an erythrocyte from the quokka (B) Microfilaria in a Modified Wright stained blood film from a Gilbert’s potoroo. Scale bars represent 10 µm.

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3.3.2 In vitro culture

Trypanosomes from the Gilbert’s potoroos were successfully cultured in vitro in both MSEM and Cunningham’s media. Examples of the different morphological forms observed in in vitro culture are shown in Figure 3.3. Trypanosomes cultured in MSEM were highly polymorphic with sphaeromastigote (Figure 3 B), promastigote (Figure 3.3

D), epimastigote (Figure 3.3 A and C) and trypomastigote-like forms observed (Figure

3.3 A). The trypomastigotes were extremely slender in form with an absent or rudimentary undulating membrane. A nucleus and a prominent dark rounded kinetoplast were identifiable in each cell. Active cell division was observed in the epimastigote

(Figure 3.3 C) and sphaeromastigote-like stages. The position and size of the nucleus and kinetoplast in relation to each other varied considerably in the dividing stages. A free flagellum that varied in length was detected in the cultivated trypanosomes. The flagellum arose from near the kinetoplast, ran along the length of the body and emerged from the anterior end.

Trypanosomes cultured in Cunningham’s medium were observed as sphaeromastigote and epimastigote-like forms with multiplication observed in the last two trypanosome stages. The trypanosomes showed similar levels of polymorphism and evidence of multiplication as trypanosomes cultured in MSEM. The epimastigote form was the most abundant life-cycle stage detected in the CM cultures.

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Figure 3.3. Light micrographs of Trypanosoma copemani from in vitro cultures of blood from a Gilbert’s potoroo. (A) Trypomastigote and epimastigote forms in a Modified Wright stained film. (B) A dividing sphaeromastigote form in a Modified Wright stained film. (C) Dividing epimastigote form with two nuclei and two kinetoplasts and sphaeromastigote form in a Modified Wright stained film. (D) Promastigote form in a Modified Wright stained blood film. Scale bars represent 10 µm.

3.3.3 Statistical analysis of morphological measurements

The mean of the measurements for each morphological parameter (length, breadth, PK, KN, NA and FF) for 36 trypanosomes isolated from the Gilbert’s potoroo, and 12 trypanosomes isolated from the quokka are presented in Table 3.2. The mean breadth of the trypanosomes isolated from the Gilbert’s potoroo were significantly different from the trypanosomes isolated from the quokka (p<0.05).

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Table 3.2. The mean dimensions and standard error (SE) of morphological features of Trypanosoma copemani isolated from blood stream forms from the Gilbert’s potoroo and quokka.

Host Feature No. of Observed Mean  SE (µm) organisms Range measured (µm) Total length 23 25.0 - 46.9 36.9  1.0 Breadth 36 2.8 - 15.4 6.6  0.4 Gilbert’s PK 36 4.1 - 17.5 8.1  0.5 potoroo KN 36 4.3 - 10.9 5.8  0.1 NA 36 7.8 - 24.0 15.5  0.5 FF 23 8.7 - 16.3 10.8  0.4 Total length 12 31.2 - 44.3 37.8  1.1 Breadth 12 1.6 - 6.0 4.2  0.3 Quokka PK 12 5.2 - 8.9 6.5  0.3 KN 12 4.0 - 7.8 5.9  0.2 NA 12 8.8 - 18.1 13.7  0.7 FF 12 8.6 - 14.9 12.1  0.5

3.3.4 Sequence analysis of Trypanosoma copemani isolated from the

Gilbert’s potoroo and quokka

A 1,022 bp region of the 18S rRNA gene was amplified for all sampled Gilbert’s potoroo (7/7) and quokkas (3/3). As mixed chromatograms were initially detected, PCR products for each sampled host were cloned as described in chapter 2 (section 2.8) and three clones for each transformation were sequenced. Two distinct genotypes were detected, which had 12 single nucleotide substitutions (SNP’s) along the 1,022 bp of sequence. One of the genotypes (referred to as T. copemani genotype A) was sequenced from 5/7 Gilbert’s potoroos and 2/3 quokkas, while the second genotype (T. copemani genotype B) was also sequenced from 5/7 Gilbert’s potoroos and 2/3 quokkas. Both genotypes were identified in 3 Gilbert’s potoroos and 1 quokka. Sequences were deposited to the GenBank database under the accession numbers EU571231-34. A distance similarity matrix (data not shown) revealed that T. copemani genotype A and B were most similar to that of an unnamed Trypanosoma species from the common wombat (Vombatus ursinus) (0.3 % and 0.9 % difference, respectively) and were 99.8% 64 similar to each other. The lowest level of genetic difference between already established

Trypanosoma species using the same partial region of 18S rRNA gene sequences ranged from 0% between T. evansi and T. brucei, 1% between T. lewisi and T. microti and 5% between T. rangeli and T. conorhini.

3.3.5 Phylogenetic relationships of Trypanosoma copemani from

Gilbert’s potoroo and quokka

Phylogenetic analysis of the partial 18S rRNA gene sequence placed T. copemani genotype A, genotype B, and the Trypanosoma sp. from the wombat

(genotype C) together in a clade with T. pestanai isolated from European badgers

(Meles meles) and an unnamed Trypanosoma species from the tick Haemaphysalis hystericus (AB281091) with strong bootstrap support using distance (99%, Figure 3.4), parsimony (84%, Figure 3.4) and maximum likelihood analyses (99%, data not shown).

Further evolutionary relationships between the members within this clade was provided by analysis of a reduced number of Trypanosoma species and with the addition of a woylie trypanosome isolate (EU518939) with a limited available 503bp product of 18S rRNA gene (Figure 3.5). Six other clades of Trypanosoma species previously described by Hamilton et al. (2007), were also observed with significant bootstrap support produced for the ‘T. brucei’, ‘Lizard’, ‘T. theileri’ and ‘T. lewisi’ clades (Figure 3.4).

The relationship of the ‘T. copemani’ clade to the other Trypanosoma species, however, could not be accurately determined due to limited bootstrap support being observed between all major clades.

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Figure 3.4. Distance-based phylogenetic tree of 32 Trypanosoma species inferred using partial 18S rRNA gene sequences (790 positions). Relationships were determined using Neighbor- joining and Maximum Composite Likelihood methods. Bootstrap values are shown as percentages of 1,000 replicates and branches corresponding to partitions reproduced in less than 50% bootstrap replicates are collapsed. Values shown in bold represent support for each clade using distance, maximum parsimony and maximum likelihood algorithms respectively. The scale bar represents the proportion of base substitutions per site. Species from Australian marsupials are shown with an asterisk.

66

Figure 3.5. (A) Linearized distance-based phylogenetic tree inferred using partial 18S rRNA gene sequences (503 positions) showing the relationship of T. copemani to other closely related species. (B) Subsection of a distance-based phylogenetic tree inferred using partial 18S rRNA gene sequences (959 positions) revealing the 3 distinct genotypes within the species T. copemani (A, B and C). Relationships were determined using Neighbor-joining and Maximum Composite Likelihood methods. Bootstrap values are shown as percentages of 1,000 replicates and units are in number of base substitutions per site. New sequences are shown in bold.

Proposed species descriptions

On the basis of genetic characterisation of a partial region of the 18S rRNA gene and phylogenetic analysis a new species of Trypanosoma, Trypanosoma copemani was proposed and described by Austen et al. (2009).

Description: Trypomastigote form: Trypanosoma copemani appeared slightly curved with a pointed posterior end, a large rounded nucleus, prominent black stained oval kinetoplast, a well developed undulating membrane and a long free flagellum. The nucleus was usually located towards the middle of the body within a granular cytoplasm. Three main forms were observed; a slender form, a medium form and a broader form. The broad form displayed myonemes giving it a striated appearance. In all three forms, the kinetoplast was observed at some distance apart from the nucleus and generally closer situated to the posterior end. The total length ranged from 25 - 46

µm (mean 36.9 µm), the breadth including the undulated membrane ranged from 2.8 -

15.4 µm (mean 6.6 µm) the posterior to kinetoplast ranged from 4.1 - 17.5 µm (mean 67

8.16 µm), the kinetoplast to nucleus ranged from 4.3 - 10.9 µm (mean 5.8 µm) while the nucleus to anterior ranged from 7.8 - 24.0 µm (mean 15.53 µm) and the free flagellum from 8.7 - 16.3 µm (mean 10.8 µm).

Type host. Gilbert’s potoroo (Potorous gilbertii)

Other hosts. Quokka (Setonix brachyuris), common wombat (Vombatus ursinus).

Type locality. Two Peoples Bay Nature Reserve (34° 58’S, 118°11’E) near

Albany, Western Australia

Location in host. Systemic circulation

Prepatent period. Unknown

Patent period. Unknown

Etymology. Named after the late Associate Professor Douglas Bruce Copeman for his contribution to Australian parasitology

3.4 Discussion

Only recently have the diverse range of Trypanosoma species of Australian marsupials begun to be morphologically and genetically characterised and placed in an evolutionary context among other Trypanosoma species of the world. The discovery of two genetically unique trypanosomes within the Gilbert’s potoroo and quokka raises questions regarding the speciation of members of the Trypanosomatidae and the need for more clearly defined parameters regarding morphological and genetic based classification. This chapter proposes the name Trypanosoma copemani (subcategorized into three genotypes) to be used to describe trypanosomes from Gilbert’s potoroo, quokka and the common wombat, however further genetic and biological data are necessary before more conclusive taxonomic categorisation can be made.

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3.4.1 Infection and pathogenesis

While only a small number of animals were captured and sampled during this study due to small population size and the nocturnal and timid nature of these native

Australian animals, all Gilbert’s potoroo and quokka that were screened were found to be infected with T. copemani. This is at odds with previous studies that showed that

Australian mammals had a low prevalence of infection with trypanosomes which was considered to be a function of low parasitemia (Noyes et al., 1999). The observed differences between the findings of this study and previous studies may be because previous studies sampled animals housed either permanently, or wild animals temporarily for treatment, in zoological parks and not in free-living populations (Noyes et al., 1999; Hamilton et al., 2004). It is possible that captive animals are not exposed to the same degree of challenge from infected vectors and reservoir hosts as animals that are in their natural ecological niche.

Interestingly, piroplasms were observed in the blood of the Gilbert’s potoroo, which have been described as Theileria gilberti n. sp. (Lee et al., 2009). A study by Cox

(1977) showed that piroplasms enhance and prolong trypanosome infections, particularly in stercorarian transmitted trypanosomes, and generally occur together in the wild (Cox, 1977). This may account for the relative high prevalence of trypanosome infection seen in the Gilbert’s potoroo. Microfilaria were also detected in two of the

Gilbert’s potoroo sampled and to the authors’ knowledge, this is the first time that microfilaria have been reported in this host.

Despite the presence of these various parasite species, no obvious clinical signs were exhibited by either Gilbert’s potoroo or quokka. Parasitic infections have most likely co-evolved with their hosts and may be of low health risk to the endangered

Gilbert’s potoroo and quokka. However, if any changes occur in the ecology of infection within these animals then overt disease may occur. Constant wildlife

69 monitoring and management of the native fauna within Two Peoples Bay, Albany,

Western Australia is advisable to prevent the introduction and potential spread of new diseases that may pose a risk to the natural ecology of these animals.

3.4.2 Morphology

Morphological characterisation showed that T. copemani isolated from Gilbert’s potoroo and quokka was highly polymorphic, with various life-cycle stages detected in both blood films and in vitro cultures. The morphological characteristics observed in blood smears and in smears from in vitro cultures were typical of Trypanosoma trypomastigote, epimastigote, promastigote and sphaeromastigote like forms. The epimastigote, and trypomastigote forms were the most principal life-cycle stages in cultures, with the former the most abundant. These findings are similar to observations by Noyes et al. (1999) who successfully cultured novel native Australian Trypanosoma spp. isolated from a common wombat and an eastern grey kangaroo and observed that epimastigotes and sphaeromastigotes were the most abundant life-cycle stages in the wombat cultures while promastigotes were the most abundant form observed in the cultures from the eastern grey kangaroo. The promastigote life-cycle stage from the

Gilbert’s potoroo compared to promastigotes studied by Noyes et al. (1999) differed in both their abundance and morphological characteristics. Promastigotes isolated from the cultures of kangaroo blood had a swollen anterior end that lacked an emergent flagellum whereas the promastigote like-stages from the Gilbert’s potoroos were less abundant and contained an emerging anterior free flagellum.

The three different morphological trypomastigote blood stream stages observed in the Gilbert’s potoroo may represent different multiplication stages. The large stout trypanosome appears to represent a pre-division stage as seen in T. lewisi

(Herpetosoma), which is known to be accompanied by growth of the body and forward migration of the kinetoplast (Hoare, 1972). The slender stage showed similar

70 morphological characteristics to tick nymph (Ixodes holocyclus) trypanosomes isolated from wild adult bandicoots (Isoodon obesulus) (Mackerras, 1959) and may represent the vector stage which gives rise to the medium trypomastigotes stage. Definitive morphological characterisation of the trypanosomes isolated from the Gilbert’s potoroo will, however, require examination of all life-cycle stages that occur in vitro and in vivo to determine the complexity of each stage.

The relative size and the morphological characteristics of the trypanosomes isolated from the Gilbert’s potoroo suggest that they may either belong to the

Megatrypanum or Herpetosoma subgenus as described by Hoare (1972). The presence of a long free flagellum, prominent rod shaped kinetoplast and the relative large distance between the kinetoplast and the nucleus in the trypomastigote blood stream form suggests that the trypanosomes from the Gilbert’s potoroo are more likely to represent trypanosomes from the Herpetosoma subgenus, as Megatrypanum trypanosomes are characterised as having kinetoplasts situated close to the nucleus, a short to absent free flagellum, are large in length ranging from 54-79 μm (such as in T. binneyi) and have a small kinetoplast (Hoare, 1972). The total length of T. copemani isolates ranged between 25.0 µm - 46.9 µm, which falls within the known range for classification into the Herpetosoma subgenus. The length of existing members of the

Herpetosoma ranges from 16.8 µm (T. sigmodoni) to 47 µm (T. myceta) in length

(Hoare, 1972).

However, there have been concerns raised about the validity of the use of this terminology (Stevens et al., 1999a). Therefore, we have adopted the current terminology

(Hoare, 1972), which may require review once a new classification system has been defined.

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3.4.3 Evolutionary and taxonomic relationships

The 18S rRNA gene was chosen as the basis of molecular characterisation in this study because it is conserved throughout the eukaryotes (Stevens et al., 1998) and it is regarded as the gene of choice for phylogenetic analysis of kinetoplasts (Haag et al.,

1998; Stevens et al., 1998; Hamilton et al., 2004). Nucleotide variation within the 18S rRNA gene has been suggested as a means of species differentiation, although no universal number of mutational changes has been postulated for species level classification. Within this study, the level of genetic variation within the partial 18S sequence between established species ranged from 0 to 5%, making it difficult to assess accurately whether novel genotypes represented new species.

Two distinct genotypes of Trypanosoma from the same geographical location were found to infect both the Gilbert’s potoroo and the quokka, representing a new species with three genotypes. Trypanosoma copemani genotype A and genotype B were genetically similar to Trypanosoma sp. (genotype C) isolated from the common wombat

(Hamilton et al., 2005a). These three genotypes form a distinct clade with high statistical support. The evolutionary significance and resolution of the relationship of the T. copemani clade to the other Trypanosoma clades is difficult to determine due to alignment problems involving the 18S rRNA gene with its regions of high nucleotide variation and large insertions and deletions. Further phylogenetic analyses using the entire open reading frame of the 18S rRNA gene, and other loci such as the glycosomal glyceraldehyde phosphate dehydrogenase (GAPDH) and multiple gene analysis are therefore needed (Hannaert et al., 1998; Hamilton et al., 2004; Hamilton et al., 2007).

Interestingly, genotypes A, B and C of T. copemani were most closely related to the KG1 Trypanosoma tick (Haemaphysalis hystricis) isolate (Thekisoe et al., 2007) and T. pestanai which was isolated from a badger in Portugal. The close relationship of the different novel genotypes of T. copemani (A, B and C) to the KG1 Trypanosoma

72 isolate may indicate that these new species are transmitted by ticks. The tick species

Ixodes australiensis was collected from the Gilbert’s potoroo and may play a role in both the transmission of trypanosomes and piroplasms in the south-west of Western

Australia. Indeed, the slender form of trypanosomes cultivated from the potoroo had similar morphological characteristics to the trypanosomes isolated from wild bandicoot tick nymphs (Ixodes holocyclus) (Mackerras, 1959). Consequently, further investigation into the role of ticks as vectors for all members of this newly described clade is therefore warranted.

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CHAPTER 4

VECTOR OF TRYPANOSOMA COPEMANI

PUTATIVELY IDENTIFIED AS IXODES SP.

Austen, J.M., Ryan, U.M., Friend, J.A., Ditcham, W.G.F., Reid, S.A., 2011. Vector of

Trypanosoma copemani identified as Ixodes sp. Parasitology, 138, 866-872.

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4.1 Introduction

The epidemiology of diseases caused by Trypanosoma species are determined in part by the ecology of the vector and the host-vector relationship that affect parasite transmission (Hoare, 1972). The majority of known vectors of mammalian trypanosomes belong to the class Insecta, and are associated with haematophagous insects of the order Hemiptera (eg. triatome bugs), Diptera (eg. flies) and Siphonaptera

(eg. flea) (Hoare, 1972). However, Arachnida (ticks), bat mites and leeches are also vectors of trypanosomes (Mackerras, 1959; Lukes et al., 1997; Stevens et al., 1999a;

Hamilton et al., 2005a; Thekisoe et al., 2007). The presence of trypanosome species in terrestrial blood sucking leeches from both Asia and Australia has been reported

(Hamilton et al., 2005a). In Australia, potential vectors such as the wombat flea

(Lycopsylla nova) and kangaroo ticks have been implicated in the transmission of unidentified Trypanosoma species isolated from Australian mammals, the wombat and kangaroo (Noyes et al., 1999). Sandflies, leeches and the platypus tick (Ixodes ornithorhynchi) have been suggested as potential vectors for T. binneyi, which infect the platypus (Mackerras, 1959; Noyes et al., 1999; Jakes et al., 2001b). Another study identified Trypanosoma sp. found within tick nymphs (Ixodes holocyclus) isolated from bandicoots infected with T. thylacis (Mackerras, 1959). Ticks (Haemaphysalis hystricis) from Japan have also been found to be naturally infected with a novel trypanosome species, isolate KGI (Thekisoe et al., 2007). Trypanosomes found in Australian ticks and the close phylogenetic relationship of T. copemani to the KGI trypanosome spread by ticks lead to the examination of ticks as potential vectors of T. copemani along with opportunistic examination of fleas.

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4.2 Materials and Methods

4.2.1 Study site and sample collection

A total of 15 quokkas were trapped at Bald Island (34º 55’ S, 118º 27’E) near

Albany, and a Gilbert’s potoroo from Two Peoples Bay during the month of June 2010.

A total of 41 ticks were collected off quokkas from Bald Island and two ticks collected off a Gilbert’s potoroo from Two Peoples Bay (Table 4.1) and placed into 5 mL tubes

(SARSTEDT) for later identification using the Australian ticks key (Roberts, 1970).

Nineteen fleas were also collected from the Gilbert’s potoroos and placed into 1.5 ml microcentrifuge tubes containing 70% ethanol. Flea samples were identified to the species level by Mr Russel Hobbs at Murdoch University with the aid of the Dunnet and

Mardon (1974) key. Fresh whole blood samples were collected from three of the quokkas from Bald Island.

Table 4.1. A list of animal isolates and isolated tick species captured from Bald Island and Two Peoples Bay used in this study.

Geographical Number of tick Host species Animal isolate location animal tick species species identified captured Quokka 3333-3316 Bald Island Quokka 3556-4432 Bald Island Quokka 3588-3589 Bald Island Quokka 3348-3558 Bald Island I. myrmecobii 1 Quokka 4450-4449 Bald Island Quokka 4489-4491 Bald Island Quokka 2322-2351 Bald Island I. australiensis 5 Quokka 4455-3334 Bald Island Quokka 4442-4418 Bald Island Quokka 2917-2918 Bald Island I. australiensis 6 Quokka 4539-4540 Bald Island I. myrmecobii 1 I. hirsti 2 Quokka 4403-4402 Bald Island I. hirsti 1 Quokka 4545-4544 Bald Island I. australiensis 2

Quokka 4444-4445 Bald Island I. australiensis 2

Quokka 2383-2359 Bald Island I. australiensis 1 Gilbert’s Two Peoples P170 I. australiensis 2 potoroo Bay

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4.2.2 Detection of trypanosomes in ectoparasites

Once identified, a selection of ten Ixodes australiensis live ticks from quokkas and two ticks from the Gilbert’s potoroo were individually mounted onto house-hold pest glue boards (TOMCAT Motomco U.S.A) allowing immobilisation of the tick and leg restraint. The glue board was then stuck to a dissecting microscope stage with double sided tape, and the tick dissected according to published methods (Edwards et al., 2009). Haemolymph was collected on the end of a scalpel blade during the removal of the tick scutum and placed onto a microscope slide for examination. The midgut and salivary glands were also removed and placed into a 1.5 mL microcentrifuge tube containing 100 µL of RPMI 1640 (pH 7.2) supplemented with 10% foetal calf serum

(FCS), 1,000 international units (IU) / mL of penicillin and 10,000 micrograms (µg) / mL of streptomycin, with final concentrations of sodium bicarbonate 2.0 g / L, L- glutamine, 0.3 g / L and HEPES; 3.6 g / L. A small region of the midgut was placed onto a microscope slide containing 10 µL of RPMI 1640 culture medium, manually pulled apart and mixed using a sterile 10 µL pipette tip. A cover-slip was placed over the dissected midgut and the preparation examined microscopically at 200 magnification to detect any motile trypanosomes. Fragments of the midgut and salivary glands were also placed into MSEM with a 500 µL overlay of RPMI 1640

(supplemented as above) and incubated in the dark at room temperature.

Once identified, the fleas were mounted in paraffin and 4-8 µm histological sections were made and stained with haematoxylin and eosin (H&E) stain to enable visualisation of the tissue structure and detection of trypanosomes.

4.2.3 Detection of trypanosomes in tick faeces

Faeces from a collection of ticks isolated from eight individual quokkas from

Bald Island and a Gilbert’s potoroo from Two Peoples Bay were removed from the inside of 5 mL polypropylene tubes with a sterile 10 µL tip placed into 10 µL of 77 phosphate buffered saline (pH 7.2) (PBS) on a microscope slide and manually dispersed. The slide was air dried and stained with Modified Wright stain.

4.2.4 Morphological measurements

Digital micrographs were taken at 1000× magnification of five tick trypanosomes from faecal smears and 14 tick trypanosomes from the dissected midgut and blood meal. Measurements of key morphological features were made. Significant differences between trypanosome morphology measurements from different host species and sources were determined using SPSS 17.0 (Chicago, Illinois, USA.) by one- way analysis of variance (ANOVA) and the Tukey’s Honestly Significant Difference test for P < 0.05.

4.2.5 DNA extraction

Total genomic DNA was extracted using a DNeasy® blood and tissue kit

(Qiagen), from both the tick faeces and dissected midgut and salivary glands of the ticks collected from quokka Q2322-2351 and Q4489-4491 and from the tick collected from the Gilbert’s potoroo. DNA was extracted from quokka Q4489-4491 blood.

4.2.6 DNA amplification and sequencing

Fragments of the 18S ribosomal RNA (rRNA) gene were amplified by PCR as described in chapter 2 (2.7.2) and purified using a QIAquick® PCR Purification kit

(QIAGEN) and sequenced directly as described in chapter 2 (2.9).

4.3. Results

4.3.1 Identification of ectoparasites

Only 25 of the 41 ticks collected from 15 quokkas from Bald Island and two ticks collected from one Gilbert’s potoroo P170 from Two Peoples Bay were sufficiently intact morphologically to allow identification down to the species level.

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Ixodes australiensis was identified from both Bald Island and Two Peoples Bay, and was the most abundant tick species collected at 72% of the total. Two other tick species with similar morphology to Ixodes hirsti and Ixodes myrmecobii were also detected on

Bald Island at 12% and 8% of the total, respectively. Ixodes australiensis was the only tick species dissected for examination because the specimens of I. hirsti and I. myrmecobii did not survive longer than 30 days in storage. The fleas isolated from the

Gilbert’s potoroo were all identified as Stephanocircus dasyurii of the family

Stephanocircidae, but no trypanosomes were identified in the fleas.

4.3.2 Trypanosomes present in tick faeces

Of the eight tick faecal smears examined from ticks collected from eight individual quokkas and one Gilbert’s potoroo, the detection of trypanosomes was observed in one wet smear of dried faeces isolated from ticks collected from quokka

Q2322-2351 and to the authors knowledge is the first time that trypanosomes have been identified in tick faeces. Morphologically the trypanosomes were slender in shape with a narrow undulating membrane, a large PK value and a free flagellum (Figure 4.1). The mean measurement for each morphological parameter for the trypanosomes are presented in Table 4.2.

Figure 4.1. Light micrographs of trypanosomes detected in the faeces of the tick (I. australiensis) using a Modified Wright stain.

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Table 4.2. Mean dimensions and standard error (S.E.) of morphological features of trypanosomes isolated from I. australiensis tick faeces from quokka Q2322-2351, I. australiensis tick midgut from quokka Q2917-2918 and T. copemani blood stream trypanosomes from quokka blood. No. of Observed Parameter Source Mean±S.E.(µm) organisms Range (µm) tick faeces 5 27.0 - 44.1 35.5 ± 2.7

tick midgut 14 21.5 - 39.8 28.2 ± 1.5 L quokka 12 31.2 - 44.3 37.8 ± 1.1 blood tick faeces 5 0.8 - 1.2 1.2 ± 0.2

tick midgut 14 0.9 - 1.7 1.4 ± 0.1 B quokka 12 1.6 - 6.0 4.2 ± 0.3 blood tick faeces 5 5.5 - 13.4 9.9 ± 1.3

tick midgut 14 3.6 - 14.4 9.0 ± 0.6 PK quokka 12 5.2 - 8.9 6.5 ± 0.3 blood tick faeces 4 0.8 - 4.1 2.7 ± 0.8

tick midgut 14 5.3 - 10.2 7.9 ± 0.3 KN quokka 12 4.0 - 7.8 5.9 ± 0.2 blood tick faeces 4 10.2 - 22.8 17.1 ± 4.0

tick midgut 14 6.3 - 18.3 11.3 ± 0.9 NA

quokka 12 8.8 - 18.1 13.7 ± 0.7 blood tick faeces 5 7.7 - 16.7 10.6 ± 1.7

FF quokka 12 8.6 - 14.9 12.1 ± 0.5 blood

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4.3.3 Trypanosome present in ectoparasites

Motile trypanosomes were detected in two of the ten ticks collected from animals from Bald Island that were dissected and examined. Trypanosomes were observed in pieces of midgut dissected from I. australiensis ticks collected from quokkas Q2322-2351 and Q2917-2918. Numerous motile trypanosomes (identified as

T. copemani by PCR) were detected in the haemolymph (Figure 4.2), as well as in the midgut containing the blood meal from one tick collected from a Gilbert’s potoroo

(P170). Morphologically, the trypanosome tick isolates resembled the slender pleomorphic trypomastigote forms of T. copemani that are present in in vitro cultures isolated from the blood of the Gilbert’s potoroo as previously described in chapter 3.

However, the free flagellum of the trypanosomes within the midgut of the tick isolated from quokka Q2917-2918 were very short or absent (Figure 4.3). The mean measurement for each morphological parameter from trypanosomes isolated from within the tick collected from quokka Q2917-2918 are presented in Table 4.2. Attempts to culture the trypanosome tick isolate in MSEM were unsuccessful. No trypanosome like forms, were observed in any of the histological sections of the examined fleas.

Figure 4.2. Light micrograph of T. copemani in Modified Wright stain detected in the haemolymph of the tick collected from the Gilbert’s potoroo P170. Trypomastigote form (t), epimastigote form (e). Scale bar represents 10 µm.

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Figure 4.3. Light micrographs of trypanosomes in Modified Wright stain from the midgut region of a tick collected from quokka Q2917-2918. (A) Trypomastigote form (t). (B) Dividing epimastigote form (e). (C) Dividing trypomastigote form (d). Scale bar represents 10 μm.

4.3.4 Comparative analysis of morphological measurements of trypanosomes from quokka blood and from ticks.

Comparison of the morphological measurements taken from the trypanosomes found in tick faeces and midgut, with measurements taken from the blood stream forms of T. copemani in quokkas (Table 4.2), showed that trypanosomes from ticks were narrower (P < 0.001) and their PK value were greater (P < 0.001) than trypanosomes from quokkas. The length of the trypanosomes both from the tick faeces and host blood stream were similar, but a significant difference was noted in the length of the midgut trypanosomes which were shorter (P < 0.001). The length of the KN parameter was highly variable, however, differences between trypanosomes from each of the three sources were significant (P < 0.001). NA was only different between trypanosomes from the midgut and tick faeces (P < 0.001). Comparison of the free flagellar length revealed no significant differences (P > 0.329) between T. copemani trypanosomes from the blood of the quokka and the faeces of the tick. The midgut trypanosomes did not possess a free flagellum.

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4.3.5 Characterization of T. copemani isolated from blood and ticks

Two of the three quokkas (Q4489-4491 and Q4539-4540) from Bald Island were positive for trypanosomes by microscopy (Figure 4.4) and amplification of the 18S rRNA gene. A tick from one of the same quokkas (Q4489-4491) as well as a tick from another quokka from Bald Island (Q2322-2351) and a tick isolate collected from the

Gilbert’s potoroo P170 was also positive by PCR. The 18S rRNA sequences from the quokka blood (Q4489-4491) and the Gilbert’s potoroo tick isolate had 100% homology to each other and 100% homology to T. copemani (genotype A) and 99.9% homology to the T. copemani wombat AII isolate. The 18S rRNA sequences from the two Bald

Island tick isolates were also confirmed as T. copemani. They had a 100% homology to each other and just one bp difference compared to the trypanosome isolated from the quokka blood (Q4489-4491) and the Gilbert’s potoroo tick isolate. No amplification was detected from DNA extracted from the tick salivary glands and the faeces.

Figure 4.4. Light micrographs of T. copemani in Modified Wright stain detected in the blood of quokka Q4489-4491 captured from Bald Island.

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4.4 Discussion

Ticks are obligate ectoparasites of vertebrates that are capable of transmitting pathogens (viruses, bacteria, rickettsiae, helminths and protozoa) of veterinary and medical importance (Roberts, 1970). In Australia, various tick species of the family

Ixodidae are responsible for transmitting zoonotic diseases, such as tick typhus

(Roberts, 1970), and spotted fever (McBride et al., 2007). This is the first time ectoparasites from quokkas on Bald Island have been examined. Ixodes myrmecobii and

I. australiensis have previously been found occurring in various regions of the South

West of Western Australia and also within regions of Tasmania. To the best of our knowledge, I. hirsti has previously only been located in the eastern states of Australia in

NSW, Victoria and Tasmania (Roberts, 1970). This is therefore the first report of I. hirsti in Western Australia.

The trypanosomes found in the midgut, haemolymph and faeces of the I. australiensis ticks had similar morphology (long and slender) to trypanosomes detected in I. holocyclus nymphs removed from bandicoots (Mackerras, 1959). The long slender trypanosomes found in the tick faeces isolated from quokka Q2322-2351 are likely to be the metacyclic form of T. copemani, as previously suggested in chapter 3. Cultured trypanosomes from the blood of the potoroos were maintained at around 25ºC, which is suitable for the growth of the vector stages of trypanosomes (Vickerman, 1985).

Morphological comparison of the trypanosomes within the vector and those from infected hosts shows that ticks harbour pleomorphic trypanosomes with some tick trypanosome life-cycle stages found to represent the multiplication forms. The difference in PK, and KN values could be due to kinetoplast migration which occurs when trypanosomes are in their multiplication and division stage (Hoare, 1972).

In the present study, motile trypanosomes were observed in the midgut of I. australiensis ticks dissected 49 and 117 days after removal of the ticks from the quokka

84 and Gilbert’s potoroo respectively. Motile trypanosomes were also detected in the haemolymph 117 days after removal. When xenodiagnosis is used for the detection of low levels of T. cruzi infection in humans or animals, first-instar Triatoma nymphs are allowed to feed on the suspect individual and after 30 days the bugs are examined for the presence of trypanosomes (Machado et al., 2001). It is unlikely that the trypanosomes observed in this study originated from the blood meal of the ticks examined because of the extended period between the last meal and detection of trypanosomes (49 days and 117 days). This is longer than the 30 days used in the xenodiagnostic test for T. cruzi. It is interesting that live T. cruzi cannot be detected in the small intestines of reduviid bugs at the fifth instar 60 days after the bug’s last blood meal (Kollien and Schaub, 1998). The detection of motile trypanosomes at day 117 within the tick midgut, suggests that T. copemani may multiply in the tick. Indeed, the observation of dividing trypanosomes in stained smears from the gut and gut contents of the tick and the detection of slender trypanosomes in the tick haemolymph suggest that the ticks were systemically infected with T. copemani. These finding suggest that I. australiensis may be naturally infected and putatively the vector of T. copemani however, transmission studies are needed for confirmation.

Trypanosomes have previously been detected in the haemolymph and gut of a variety of tick species (Mackerras, 1959; Shastri and Deshpande, 1981; El Kady, 1998;

Latif et al., 2004; Thekisoe et al., 2007). However there is insufficient information to fully determine the mode of natural trypanosome transmission via ticks. The detection of intact trypanosomes and trypanosome DNA in smears of dried tick faeces and the detection of DNA in the midgut but not from salivary glands suggest that transmission is likely to be contaminative via tick faeces. The oral route of transmission that occurs with T. lewisi when rats eat infected fleas and/or their droppings containing metatrypanosomes (Hoare, 1972) could be a similar process of transmission that might

85 be occurring between T. copemani and its marsupial host. Ticks located on quokkas are generally found around the ears and the tail while on the Gilbert’s potoroo they are more widely distributed ranging from ears, tail, rim of pouch in females and around the scrotum in males as well as amongst the fur on the body, all of which are body parts generally accessible to grooming. The stercorarian nature of T. copemani is consistent with a faecal route of trypanosome transmission. However, it is not possible to rule out the salivary route of transmission because only a limited number of salivary glands were analysed. Furthermore, there is evidence that a similar trypanosome such as T. rangeli are transmitted via both the contaminative and inoculative routes (Guhl and Vallejo,

2003).

Sequence analysis of the amplified 18S rRNA gene from the DNA extracted from the tick midgut showed that I. australiensis on Bald Island and Two Peoples Bay are carrying a strain of T. copemani (genotype A) that is similar to T. copemani isolates from wombats in Victoria, Australia (Noyes et al., 1999). The single base-pair difference may have been due to an error introduced by the polymerase. The wide geographical distribution of T. copemani across Australian may be accounted for by the presence of Ixodes tick species, which have been found in most states of Australia.

Ticks show varying degrees of host specificity (Roberts, 1970). Ixodes ticks are generally not host-specific. Ixodes australiensis has been recorded from numerous hosts including small marsupials, such as bettongs (Bettongia lesueuri and B. penicillata), potoroos (Potorous tridactylus and P. gilbertii), the quokka, as well as kangaroos, dogs, humans and cows (Roberts, 1970; Lee et al., 2009). Leeches have been identified as vectors of trypanosomes in Australia (Hamilton et al., 2005a) and several authors have suggested various ectoparasites (fleas, biting flies, and ticks) as potential vectors for native Australian trypanosomes (Noyes et al., 1999; Jakes et al., 2001b).

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Australian trypanosomes have been found in a wide range of marsupials from mainland Australia (Mackerras, 1959; Noyes et al., 1999; Jakes et al., 2001b; Hamilton et al., 2005a; McInnes et al., 2009) as well as offshore islands (Bettiol et al., 1998). The identification of the vector for T. copemani is important for elucidation of the complete life-cycle. Knowledge of the life-cycle will aid in management decision making when reintroduction of naïve animals occurs into areas where the parasite is known to be present in the same or other species of marsupials. This is particularly important when a population of animals from the same species are genetically and morphologically different, as occurs with quokkas on Rottnest in comparison to mainland and Bald

Island quokka populations (Sinclair, 1998, 2001). The genetic composition of one population if different from another population of the same species may make it more resilient to parasitic disease, particularly if co evolution has occurred between the parasite and host of one of the populations and not the other.

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CHAPTER 5

INVESTIGATION OF THE MORPHOLOGICAL

DIVERSITY OF NATIVE AUSTRALIAN

TRYPANOSOMES IN QUOKKAS AND GILBERT’S

POTOROOS

Austen, J.M., Reid, S. A., Robinson, D. R., Friend, J. A., Ditcham, W. G. F., Irwin P.

J., Ryan,U. 2015. Investigation of the morphological diversity of the potentially zoonotic Trypanosoma copemani in quokkas and Gilbert’s potoroos. Parasitology, 142,

11, 1443-52.

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5.1 Introduction

The present distribution of the quokka includes a number of sites on mainland

WA, ranging from the Darling Plateau near Perth to the Green Range on the south coast east of Albany and two offshore islands, Bald Island and Rottnest (Sinclair, 1998).

Rottnest Island is located 18 km off the coast of WA and has been an important local holiday destination for over 50 years. Quokkas are the only native marsupials to inhabit

Rottnest Island and a close relationship has developed between tourists and quokkas

(Hart et al., 1985; Sinclair, 1998). To date, there is little available information on the potential impact of pathogenic parasites on the health of native wildlife in Australia. In chapter 3, quokkas captured from Two Peoples Bay in Albany were identified as being positive for trypanosomes, while epimastigote, trypomastigote, sphaeromastigote and promastigote stages were identified from in vitro culture. This native Australian species of trypanosome recently identified as T. copemani in chapter 3, is known to be infective to a variety of Australian marsupials including the critically endangered Gilbert’s potoroo, common wombat, koalas, woylies, southern brown bandicoot (Isoodon obesulus), tiger quoll (Dasyurus maculatus) and common brushtail possum (Noyes et al., 1999; McInnes et al., 2009; Botero et al., 2013; Thompson et al., 2013).

Importantly, as these animals have been shown to be infected with T. copemani, studies by McInnes et al. (2011) and Botero et al. (2013) have shown that T. copemani is associated with pathological impacts on both koalas and woylies. Significantly low packed cell volume (PCV) and regenerative anaemia were reported from koalas infected with T. copemani and other trypanosome species, while histopathological changes to cardiac and smooth muscle were reported in woylies, which were speculated to be due to the amastigote life-cycle stage (McInnes et al., 2011; Botero et al., 2013). Mixed trypanosome infections have been reported in woylies and koalas, with woylies co- infected with T. vegrandis and T. copemani and koalas with T. irwini, T. gilletti, T.

89 copemani and T. vegrandis (McInnes et al., 2010; McInnes et al., 2011; Botero et al.,

2013; Barbosa et al., 2016). The tick, Ixodes australiensis, has been identified as a possiable putative vector for T. copemani as previously described in chapter 4, but the vector for other trypanosome species infecting marsupials is unknown. Currently no treatment programs exist for native Australian trypanosomes. Importantly, T. copemani has also been shown to be resistant to normal human serum (chapter 7) and therefore may be potentially zoonotic.

Morphologically T. copemani is highly polymorphic with three main trypomastigote forms (thin, medium and broad) as previously described in chapter 3.

Broad and slender trypomastigote forms of T. copemani have also been observed in blood films from woylies (Thompson et al., 2013). The main aim of the present study was to elucidate the different morphological forms of T. copemani and other native

Australian trypanosomes within the circulatory systems of the quokka and the Gilbert’s potoroo. Knowledge of the different blood stream and in vitro forms of this parasite is imperative for clinical diagnostics and information gained from this study will help in monitoring and management of infected animals and allow for better treatment programs to be implemented.

5.2 Materials and Methods

5.2.1 Study site and sample collection

A total of 35, 34 and 41 blood samples were collected from quokkas captured at

Two Peoples Bay, Bald Island and Rottnest Island respectively Opportunistic samples from two Gilbert’s potoroos (P83 and P94) were also examined (Table 5.1).

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Table 5.1. A list of quokka and Gilbert’ potoroo isolates captured from the three different geographical locations used in this study.

Two Peoples Bay Two Peoples Bay Bald Island Rottnest Island Gilbert’s Potroo quokka isolates n =35 quokka isolates n=34 quokka isolates n=41 isolates n=2

Q1364-1356 Q1997-1961 RS1 P83 Q1416-1407 Q1 RS2 P94 Q1339-1364 Q2382-2373 RS3 Q1837-1464 (Q5) Q2522-2523 RS4 Q1996-1999 Q 2581-2582 RS5 Q1362 Q2351-2346 RS6 Q2399-2324 Q1947 RS7 Q1342-1338 Q2367-2332 RS8 Q1354-1368 (Q10) Q2377-2381 RS9 Q1369-1355 Q2363-2376 RS10 Q2088-2050 Q2383-2359 RS11 Q1403-1404 Q2578--2600 WS1 Q1382-1326 Q Q2933-3201 WS2 Q1340 Q3307 BS1 Q2031 Q3343-3304 BS2 Q1051 Q3355-3284 BS3 Q1767 Q3336-3325 BS4 Q1366 Q2385-2359 BS5 Q2990 Q3206-3207 BS6 Q4137-1443 Q3313-4458 BS7 Q4177-4185 Q3346-4462 BS8 Q4148-4151 Q3542-3543 BS9 Q4124-4119 Q3547-3548 BS10 Q4140-4178 Q4435-3539 BS11 Q4299-4300 Q3293-4460 BS12 Q4141-4193 Q4478-3340 BS13 Q4120-4129 Q4461-4497 BS14 Q4166-2325 Q4491-4489 BS15 Q4176-4133 Q4540-4539 WE2 Q4131-4159 Q3269-3268 WE3 Q4112-4117 Q3364-3201 WE4 Q4115-4192 Q3229-3228 WE5 Q4174-4102 Q3362-3317 WE6 Q4908-4846 Q3258-3257 WE7 Q4633-4613 WE8 WE9 WE10 WE11 WE12 WE13 WETL1

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5.2.2 Microscopy

Light microscopy and staining was conducted as described in chapter 2 (section

2.3.1). Scanning electron microscopy (SEM) was conducted as described in chapter 2

(section 2.5). Immuno-fluorescent and DAPI staining was conducted as described in chapter 2 (section 2.4).

5.2.3 In vitro cultivation of trypanosomes

5.2.3.1 BIIT medium

Modified Sloppy Evans medium and Cunningham’s liquid medium were prepared as described in chapter 2. The BIIT medium was prepared in the same way as

MSEM (Noyes et al., 1999), however human blood from a healthy volunteer was used as a supplement in the medium instead of horse blood. A 50 µL aliquot of fresh quokka blood was placed into separate tubes containing 250 μL of fresh undiluted human serum. These tubes were then incubated in a water bath at 37°C for 5 hours. The entire contents of each incubated tube was then added to individual tubes containing 1 mL of human MSEM and incubated at room temperature in the dark for 24 hours, before examination. Microscopic examination of wet-smear preparations of the culture was performed every day at 200× and 400× magnification to detect the presence of motile trypanosomes. If trypanosomes were detected, Modified Wright-Giemsa stained thin blood smears were prepared for further microscopic examination. The use of human subjects for this study was approved by the Murdoch University human ethics committee (project number 2010/053).

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5.2.4 Fluorescence in situ hybridisation (FISH)

To confirm the presence of novel trypanosome-like life-cycle forms that lacked a free flagellum, fluorescence in situ hybridisation (FISH) was performed on both blood and culture life-cycle forms using a commercially synthesised probe specific to the trypanosome 18S rRNA gene (TRYall1F 5’ ACCGTTTCGGCTTTTGTTGG 3’) manufactured with a 5' 6-carboxyfluorescein (FAM) fluorescent tag at the 5’ end

(GeneWorks, Adelaide, South Australia) as previously described (Thompson et al.,

2013), on MSEM culture and whole blood smears. The hybridised slides were examined with a BX51 microscope (Olympus, Japan) using ultraviolet light (330-385nm) through an emission filter (420 nm) producing green fluorescence. The slides were scanned at

400× and 1,000× magnification and digital images captured using an Olympus DP71

Advance digital camera.

5.2.5 Molecular characterisation

To confirm the presence of trypanosomes, universal primers were used to amplify and sequence an ~ 1,439 bp fragment of the 18S ribosomal RNA (rRNA) gene as previously described in chapter 2 (section 2.7.2). In addition, the samples were also screened with

T. vegrandis specific primers, as this species has previously been detected in marsupial blood from woylies, a Western grey kangaroo (Macropus fuliginosus), a quenda

(Isoodon obesulus), koalas, and a tammar wallaby (Macropus eugenii) (Botero et al.,

2013; Thompson et al., 2013; Barbosa et al., 2016). Briefly, an ~ 900 bp fragment of the

18S rRNA gene was amplified using primers S823F

(5'CGAACAACTGCCCTATCAGC3’) and S662 (5’ GACTACAATGGTCTCTAATC

3’), sourced from Maslov et al. (1996), as external primers in a reaction performed according to McInnes et al. (2009). The PCR product was run on a 2% agarose gel stained with SYBR safe (Invitrogen, USA). The gel band was excised and purified using an in-house filter tip method as previously described (Yang et al., 2013) and 1 µl 93 of gel-purified PCR was used in a secondary reaction with internal T. vegrandis-specific primers TVIF 5’ ACCAAAAACGTGCACGTG 3’ and TVIR 5’

AAATCGTCTCCGCTTTAAC 3’, which amplify a 350 bp product as previously described (Botero et al., 2013). All samples were also screened for piroplasms

(Theileria and Babesia), using 18S nested primers as previously described (Jefferies et al., 2007).

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5.3 Results

5.3.1 Trypanosome morphology

Using light microscopy, the morphology of trypomastigotes detected in blood smears from quokkas from both Two Peoples Bay and Bald Island were consistent with the different morphological forms of T. copemani from quokkas and Gilbert’s potoroos representing slender (Figure 5.1A), medium (Figure 5.1B) and broad forms (Figure

5.1C) as previously described in chapter 3. Trypanosomes representing typical trypomastigote blood cycle stages were not detected in any of the Rottnest Island quokka blood smears.

Figure 5.1. Light micrographs of trypanosomes in a Modified Wright-Giemsa stained blood film. (A) The slender stage in a blood film from quokka Q1369-1355 from Two Peoples Bay. (B) The medium stage in a blood film from quokka Q3355-3284 from Two Peoples Bay. (C) The broad stage in a blood film from quokka Q3355-3284 from Two Peoples Bay. Scale bars represent 10 µm.

In addition to the three morphologically different trypomastigote forms, other various potential trypanosome life-cycle stages were detected directly from blood samples from quokkas for the first time using light microscopy, in vitro cultivation,

SEM, immunofluorescence and FISH.

Trypanosome forms representing promastigote (Figure 5.2 A) and sphaeromastigote (Figure 5.3 A) stages were identified. In addition, an amastigote stage

(Figure 5.4) and three novel trypanosome life-cycle forms representing an oval form

(Figure 5.5), an extremely thin form (Figure 5.6), and an adherent form (Figure 5.7)

95 were detected directly within blood films and in vitro, while two novel culture forms representing a tiny form (Figure 5.8) and a circular form (Figure 5.9) were only detected in vitro. The detection rate of trypanosomes in direct blood films was very low and often required several smears from the same individual to be examined in order to detect a single trypomastigote. In comparison, trypanosome-positive blood cultured in vitro, yielded more abundant trypanosome life-cycle forms that were detected as early as 5 days post inoculation. The most prevalent life-cycle stages in vitro were epimastigotes, followed by thin trypomastigotes, sphaeromastigotes, amastigotes and promastigotes respectively. With regards to the potential novel trypanosome forms, the most prevalent form in culture and in blood films was the adherent form which is in contrast to the extremely thin form that was only detected once in both in vivo and in vitro.

During screening of blood smears several erythrocyte abnormalities were observed in the blood of infected quokkas. These included the appearance of acanthocytes (star shaped erythrocytes), echinocytes (erythrocytes with many small, evenly spaced thorny projections on the cell membrane), schistocytes (irregular shaped erythrocytes), dacrocytes (tear drop erythrocytes), microspherocytes (small round dense erythrocytes less than 4 µm with no central pallor) and burst erythrocytes (data not shown), which are all cell types associated with haemolytic anaemia (Rodak et al.,

2012).

5.3.1.1 Promastigote stage

Only one promastigote stage parasite (represented by an elongated form and an ante-nuclear kinetoplast with a flagellum arising near it and emerging from the anterior end of the body) was detected in a single blood film (Figure 5.2A) from quokka Q1340

(which was co-infected with T. vegrandis and T. copemani) captured from Two Peoples

Bay and from in vitro culture. Morphologically this stage was granular in appearance and was observed as having a long, pointed posterior, no undulating membrane, a

96 rounded anterior containing the nucleus, an antenuclear kinetoplast and a free flagellum.

The measurable morphological dimensions of this novel trypanosome stage in blood were; 17.0 µm in total length, 8.9 µm from posterior to kinetoplast, and 1.3 µm from nucleus to anterior.

The promastigote stage was more abundant in vitro and was detected using SEM

(Figure 5.2 B) and immunofluorescence. Immunofluorescent staining using a monoclonal antibody (L8C4) (specific for the paraflagellar rod for trypanosomes) and

DAPI showed bright green fluorescence along the length of the free flagellum and bright blue nuclear staining of both the nucleus and kinetoplast confirming the promastigote stage (Figure 5.2 C-F).

Figure 5.2. (A) Light micrograph of the trypanosome promastigote stage in a Modified Wright- Giemsa stained blood film from quokka Q1340 from Two Peoples Bay. (B) Scanning electron micrograph of the promastigote trypanosome stage from in vitro culture originally isolated from Gilbert’s potoroo P83. (C) Unstained trypanosome using differential interference contrast (DIC) from Gilbert’s potoroo P83. (D) Immunofluorescent staining of the paraflagellar rod of a trypanosome isolated from Gilbert’s potoroo P83 using monoclonal antibody (L8C4). (E) DAPI staining of nuclear DNA from Gilbert’s potoroo P83. (F) Combined images of C, D and E. Scale bars represent 10 µm.

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5.3.1.2 Sphaeromastigote stage

Only one sphaeromastigote stage was observed in a blood smear from one quokka (Q1342-1338) captured from Two Peoples Bay (Figure 5.3A) and a few from a quokka captured from Rottnest Island (QBS5). This is in contrast to the in vitro sphaeromastigote stages observed as the third most abundant life-cycle stage with epimastigotes and thin trypomastigotes the most prevalent forms. The in vitro sphaeromastigote forms originating from the blood of a Gilbert’s potoroo (P94) (Figure

5.3B) and a quokka from Bald Island (Q1) were detected using light microscopy and confirmed as a trypanosome stage with the use of the immunofluorescence monoclonal antibody L8C4 and DAPI (Figure 5.3C). Morphologically the sphaeromastigote was small and rounded with a free flagellum. The morphological dimensions of the sphaeromastigote in blood were; 6.28 µm in total length, 2.3 µm in breadth, and 3.3 µm in free flagellum.

Figure 5.3. Micrographs of sphaeromastigote trypanosomes. (A) Light micrograph of a sphaeromastigote stage (indicated by arrow) in a Modified Wright-Giemsa stained blood film from quokka Q1342-1338 from Two Peoples Bay. (B) Sphaeromastigote stage from in vitro culture originally isolated from Gilbert’s potoroo P94. (C) Immunofluorescent staining using monoclonal antibody (L8C4) of a sphaeromastigote in vitro isolated from a quokka Q1 from Bald Island. Scale bars represent 10 µm.

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5.3.1.3 Amastigote stage

The amastigote stage was observed in blood smears from quokka QRS3 from

Rottnest Island (Figure 5.4 A-B) and quokka Q4908-4846 and in vitro from quokka

Q1342-1338 (Figure 5.4 C-D) and confirmed using FISH. Morphologically amastigotes are small and round to oval, with a nuclear region and a peripheral kinetoplast with either a small or absent free flagellum. They measured 1.79 µm in length and 1.02 µm in width. Amastigotes were observed in culture generally clumped together, however single independent motile trypanosomes with short beating flagellum were also detected, generally swimming in a circular motion.

Figure 5.4. (A) Light micrograph of the trypanosome amastigote form in vivo from quokka QRS3. (B) FISH analysis of amastigote in vivo from quokka Q4908-4846. (C) Light micrograph of amastigote in vitro stained with Giemsa from quokka Q1342-1338 N = nucleus, K = kinetoplast. (D) FISH analysis of the same amastigote in in vitro culture. Scale bar represents 10 µm.

5.3.1.4 Oval form

Two singular oval form trypanosomes were observed in a blood smear (Figure

5.5 A) from quokka Q1051 captured from Two Peoples Bay and also in vitro from a

Gilbert’s potoroo P94 (Figure 5.5 B) and quokka Q2088-2050 (Figure 5.5 C).

Morphologically the oval form was granular in appearance with structures that stained with Modified Wright- Giemsa stain, had a short pointed posterior, oval body and free flagellum. The presence of the stained structures made it difficult to accurately determine the nucleus and kinetoplast. The morphological dimensions of this previously

99 unidentified novel trypanosome form in vivo were 9.4 µm in total length and 4.4 µm in free flagellum.

Figure 5.5. (A) Light micrograph of the oval form trypanosomes in vivo from quokka Q1051 from Two Peoples Bay. (B) Oval shape form trypanosome from in vitro culture originally isolated from Gilbert’s potoroo P94. (C) Oval form trypanosome in vitro culture from quokka Q2088-2050 stained with Modified Wright-Giemsa stain. Scale bars represent 10 µm.

5.3.1.5 Extremely thin form

An extremely thin trypanosome-like form was observed once in a blood smear

(Figure 5.6 A) from quokka Q3343-3304 captured from Bald Island and only detected once within in vitro culture from quokka Q2088-2050 (Figure 5.6 B). Morphologically this thin novel form was observed as having a kinetoplast, a nucleus, and a long pointed posterior end.

Figure 5.6. (A) Extremely thin trypanosome form in a Modified Wright-Giemsa stained blood film from quokka Q3343-3304 from Bald Island. (B) Light micrograph of the extremely thin trypanosome form in BIIT in vitro culture, originally isolated from Q2088-2050. Scale bars represent 10 µm.

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5.3.1.6 Adherent and free moving forms

Adherent trypanosome forms both in vivo and in vitro were first represented by small rounded independently rapid moving free forms and forms observed adherent to erythrocytes (Figure 5.7 A, Q2088-2050). Initially the adherent free form was observed as a tiny rounded circle, clear to greenish brown in appearance. It then appeared to develop into a small dark circle once it adhered to the surface of an erythrocyte. The adherent region developed further and could be seen with a cell body radiating out on either side of the nucleus when adhered to the surface of an erythrocyte (Figure 5.7 B,

Q3336-3325). Immunofluorescent staining using the L8C4 monoclonal antibody showed binding of the antibody (observed as green fluorescence) to the adherent flagella, and co-staining with DAPI stained the nucleus (Figure 5.7 C Q1). FISH analysis in vivo using a specific Trypanosoma probe further confirmed these forms, with binding of the DNA probe (observed as green fluorescence) to both the initial (Figure

5.7 D, Q4633-4613) and the more advanced forms (Figure 5.7 E, Q4633-4613) adherent to the surface of erythrocytes.

Figure 5.7. Light micrographs of (A) Initial adherent trypanosome form on the surface of the erythrocyte (indicated by arrow) from quokka Q2088-2050. (B) A more developed trypanosome adherent form on an erythrocyte in MSEM in vitro culture originally isolated from quokka Q3336-3325 from Bald Island. (C) Immunofluorescent staining using L8C4 monoclonal antibody of the adherent trypanosome form showing green fluorescence of the flagellum and co- staining of nuclei (blue fluorescence) using DAPI from quokka Q1. (D) FISH analysis of two initial adherent trypanosome forms showing green fluorescence (white arrows) adhered to the surface of an erythrocyte in vivo (Q4633-4613). (E) FISH analysis of the more advanced adherent trypanosome form (white arrow) on the surface of an erythrocyte in vivo isolated from quokka Q4633-4613 from Two Peoples Bay. Scale bars represent 10 µm.

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5.3.1.7 Tiny form

A very tiny round trypanosome-like form was detected by light microscopy

(Figure 5.8 A) and SEM (Figure 5.8 B) in cultures originally from the blood of a

Gilbert’s potoroo, P83. This form had a prominent pointed posterior end, a rounded anterior end and free flagellum and was often observed independently motile and rapid in motion.

Figure 5.8 (A) The tiny trypanosome from in vitro culture, originally isolated from Gilbert’s potoroo P83. Scale bars represent 10 µm. (B) SEM of the tiny trypanosome form (indicated by arrow) from in vitro culture originally isolated from Gilbert’s potoroo P83. Scale bars represent 2.5 µm.

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5.3.1.8 Circular form

A circular trypanosome form was detected in vitro (Figure 5.9, quokka Q1) using immunofluorescent staining with the L8C4 monoclonal antibody. Green fluorescence of the flagellum could be seen around the circumference of the parasite

(Figure 5.9 A) and DAPI co-staining of the nuclear region in the centre (Figure 5.9 B).

Figure 5.9. (A) Immunofluorescent staining (L8C4) of the circular trypanosome form in vitro showing green fluorescence (white arrows) of the flagellum around the circumference of the parasite (from quokka Q1). (B) The same form showing the nuclear region within the centre stained with DAPI. Scale bars represent 10 µm.

5.3.4 Molecular confirmation of trypanosomes and piroplasms

To confirm the presence of trypanosomes, all blood samples collected from quokkas and Gilbert’s potoroos were screened by PCR and positives were confirmed as

T. copemani by sequence analysis of the 18S rRNA gene. To determine the potential of trypanosome co-infections, all marsupial samples were also screened for T. vegrandis using a species-specific PCR. All samples tested negative, with the exception of quokka

Q1340, from Two Peoples Bay, which was positive for both T. vegrandis and T. copemani. The Rottnest Island quokkas were all negative for piroplasms, whereas all the quokkas from both Two Peoples Bay and Bald Island were positive for piroplasms by PCR.

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5.4. Discussion

The present study is the first report of trypanosomes (confirmed as T. copemani by DNA analysis) in quokkas on Rottnest Island. The previous lack of detection of trypanosomes on Rottnest Island, compared to the detection of the parasites in quokkas from Two Peoples Bay and Bald Island is likely due to the absence of the typical diagnostic trypomastigote stages in previously examined blood films (Austen et al.,

2011) Morphologically the trypanosomes from the Rottnest quokkas represented sphaeromastigote and amastigote stages.

Trypanosoma copemani has a polymorphic life-cycle with three trypomastigote forms, a thin, medium and broad form, described from the Gilbert’s potoroo and a medium form described from the quokka as reported previously in chapter 3. The thin and broad forms of T. copemani have also been described from woylies (Thompson et al., 2013). Previous studies, however, have detected only trypomastigote life-cycle stages in the peripheral blood of Australian native marsupials. In the present study, we have detected the promastigote, sphaeromastigote and amastigote forms (Hoare, 1972) directly in blood films, and have preliminary confirmation that these three stages are trypanosomes with the use of trypanosome specific FISH and immunofluorescent probes. These findings show that trypomastigotes are not the only life-cycle stages circulating within the blood stream of Australian native marsupials infected with trypanosomes.

The promastigote, sphaeromastigote and amastigote stages appeared quite granular in appearance with structures stained by Modified Wright-Giemsa, making it difficult to clearly distinguish the nucleus and kinetoplast regions, thus limiting our morphometric analysis. The small size and the lack of distinguishing features may be the cause of these stages being previously unidentified, as well as the low level of parasitaemia observed in Australian marsupials (Noyes et al., 1999). The promastigote

104 stage, however, had a more defined nuclear region and was shown to be more prevalent in culture compared to in vivo with only one promastigote stage identified directly from within the host.

The sphaeromastigote stage has previously been identified in vitro, from the

Gilbert’s potoroos (chapter 3). This is the first time that it has been detected directly in quokka blood and confirmed in vivo using immunofluorescent staining. The presence of the sphaeromastigote stage in blood supports the free circulation of amastigotes in the blood stream, since sphaeromastigotes represent transitional stages between amastigotes and epimastigotes in trypanosome stercorarian life-cycles (Hoare, 1972).

The amastigote stage was identified directly within blood films from quokkas for the first time to our knowledge and confirmed using FISH. Amastigote stages are generally considered intra-cellular, with multiplication occurring exclusively within cells. Studies however have shown that they can be present within the circulatory system (Ley et al., 1988; Tyler and Engman, 2001). Pyogranulomatous myocarditis, endocarditis, muscle degeneration and necrosis with structures present suggestive of amastigotes have been reported in histopathological studies of heart sections from woylies naturally infected with T. copemani (Botero et al., 2013). Further research however is needed to confirm these forms.

The small size of the presumed amastigote and its presence in the circulating blood could result in it being incorrectly identified as a platelet, which has implications for prevalence studies and clinical diagnosis. It is possible that what are currently thought to be platelets are actually amastigote forms or the infective metacyclic forms circulating within the blood. Future studies including platelet-specific stains and additional trypanosome specific probes are needed for additional confirmation.

Four novel trypanosome forms were detected in the present study representing an oval form, an extremely thin form, an adherent form and a tiny form. These forms

105 were identified both in vitro and in quokka blood films. A circular form was also identified only in vitro and may represent a transitional epimastigote rounding up and forming into an amastigote. The novel oval form may represent a transitional form between a sphaeromastigote and an epimastigote, as no clear nucleus or kinetoplast was distinguishable due to granular staining. This oval form appears to be morphologically similar to a cultured trypanosome isolate ABF, previously isolated from a wallaby, which also lacked a distinctive nucleus and kinetoplast region (Hamilton et al., 2005a).

The extremely thin form may represent a transitory trypomastigote form, just released from a ruptured host cell. The tiny form appears to morphologically resemble a metacylic trypomastigote with the exception of a shorter free flagellum, as described previously by Tyler and Engman, (2001). To fully validate these novel forms as trypanosomes, further analysis is needed such as binding of trypanosome specific probes to these potential life forms and the use of TEM for ultra-structural identification.

The adherent form characterised as a small rounded nuclear dot and later a more developed trypanosome was detected on the surface of erythrocytes in the blood smears of quokkas and Gilbert’s potoroos naturally infected with trypanosomes, and in vitro from quokkas captured from Two Peoples Bay, Bald Island and Rottnest Island. It is possible that this is a piroplasm life-cycle form as it has similar morphological characteristics to a marginal form of two piroplasm species (Babesia tachyglossi and

Theileria tachyglossi) identified in the blood of an echidna (Clark et al., 2004).

However, Babesia and Theileria were absent in all the Rottnest Island quokkas by both microscopy and PCR screening at the 18S rRNA locus. The binding of the L8C4 monoclonal antibody and DNA probe also supports the hypothesis that these novel forms are potential trypanosome forms because both these probes are specific for

Trypanosoma.

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Generally the adherent form was observed in a more advanced stage in vitro, containing a kinetoplast with a cell body radiating out from either side and attached to the surface of the erythrocyte. Whether the adherent form is actually attached to the surface of the erythrocyte is unknown. However, trypanosomes attaching to erythrocytes have previously been demonstrated in T. gambiense, T. rhodesiense, T. b. brucei, T. congolense and T. lewisi (Silva et al., 1995). Light and electron microscopy studies have reported the adhesion of T. b. brucei to erythrocytes in deer mice

(Peromyscus maniculatus) (Anosa and Kaneko, 1983), while Silva (1995) reported T. evansi adhered to erythrocytes from dogs and horses. These forms were also detected in the BIIT medium as discussed in chapter 7. Before the adherent form can be confidently validated, TEM analysis and the use of nuclear staining such as DAPI, are vital to show that these forms are freely independent and living trypanosome forms.

The most common type of anaemia during acute phase trypanosomiasis is haemolytic anaemia. The exact mechanism of erythrocyte destruction is unclear but it is believed that erythrocytes become coated with antigens from lysed trypanosomes leading to erythrophagocytosis due to antigen:antibody complement complexes bound to the surface of erythrocytes (Amole et al., 1982; Taylor and Authie, 2004). This process has been shown to occur in both human and animal trypanosomiasis (Woodruff et al., 1973; Kobayashi et al., 1976).

Therefore the adherent forms may have the potential to cause erythrocyte destruction by inducing immune antigen:antibody complexes and consequently leading to both erythrophagocytosis and haemolytic anaemia. Consistent with our findings of erythrocyte abnormalities, vacuolation, torocytes, acanthocytes, microspherocytes, schistocytes and dacrocytes have previously been reported in trypanosome-infected blood (Anosa and Kaneko, 1983; Silva et al., 1995). The latter four abnormalities are recognised clinical forms of anaemia with microspherocytes and schistocytes typically

107 associated with haemolytic anaemias (Mallah et al., 2010; Rodak et al., 2012).

Examination of erythrocytes from a quokka blood sample negative for trypanosomes was consistente with eosinophilic discocyte erythrocytes that showed a moderate central pallor (Clark et al., 2004) with the exception of a few dacrocytes detected. However, more in depth research is needed to determine if Australian native trypanosomes can cause trypanosomiasis. In particular, haematological and biochemical analysis of larger data sets of trypanosome positive and negative marsupials, is highly recommended to fully understand the effects of Australian trypanosomes on the health of marsupial erythrocytes.

The present study demonstrates our limited understanding of the complex life- cycles of Australian native trypanosomes. Given that all the quokkas and both Gilbert’s potoroo isolates were positive for T. copemani by sequence analysis at the 18S rDNA locus and negative for T. vegrandis, with the exception of one quokka (Q1340), the life- cycle forms identified in this study are highly suggestive of T. copemani. The promastigote identified in quokka Q1340, which was co-infected with both T. copemani and T. vegrandis by molecular characterisation, is unlikely to represent T. vegrandis as

T. vegrandis is the smallest trypanosome identified to date (Thompson et al., 2013).

Measurements for T. vegrandis promastigotes are not available, but trypomastigote stages measure 8.30 ± 0.28 (6.92 - 10.50) µm in length (Thompson et al., 2013). In contrast, the promastigote stage described from quokka Q1340 in the present study was

17.0 µm in length, which is too large to be T. vegrandis. However, to conclusively identify the trypanosome species represented by each life-cycle form, species-specific probes need to be applied. The importance of identifying and understanding all the stages of native Australian trypanosome life-cycles is important when considering clinical diagnosis, false negatives in epidemiological studies, management of native

108 wild life and translocation studies and preventing disease outbreaks as no treatment to date exists.

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CHAPTER 6

MOLECULAR CHARACTERISATION OF NATIVE

AUSTRALIAN TRYPANOSOMES IN QUOKKA

(SETONIX BRACHYURUS) POPULATIONS FROM

WESTERN AUSTRALIA

Austen, J. M., Paparini, A., Reid, S. A., Friend, J. A., Ditcham,W. G. F., Una Ryan, U.

2015. Molecular characterisation of native Australian trypanosomes in quokka (Setonix brachyurus) populations from Western Australia. Parasitology International, 65, (3),

205-208.

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6.1 Introduction

In contrast to the wealth of information on economically important species such as T. cruzi and T. brucei, knowledge of the prevalence and genetic diversity of native

Australian trypanosomes is considerably limited. The first phylogenetic analysis of

Trypanosoma sp. infecting native Australian mammals was conducted by Stevens et al.

1999. More recent phylogenetic analysis has extended this analysis in a range of marsupial hosts (Noyes et al., 1999; Stevens et al., 2001; Hamilton et al., 2005a;

McInnes et al., 2010; Paparini et al., 2011; Botero et al., 2013). In a study by Botero et al. (2013), eight novel trypanosome genotypes designated G1 to G8 were reported from a variety of Australian marsupials. G1 and G2 group with T. copemani isolates previously described from wombats and koalas, while G3-G7 clustered within the T. vegrandis clade (Botero et al., 2013; Thompson et al., 2013). Genotype 8 exhibited a

99% similarity to an Australian kangaroo trypanosome isolate (H25), which has previously been reported to be genetically similar to T. cruzi (Stevens et al., 1999a). The

18S rRNA locus has been the most widely utilised gene to determine evolutionary relationships of the order Kinetoplastida. However recent phylogenetic studies have shown that the use of the 18S rRNA locus alone to describe a new species is considered inappropriate due to intra-species variation at this locus (Hamilton et al., 2004). In the present study, molecular characterisation of both the 18S rRNA and GAPDH (which has been shown to be a more reliable phylogenetic marker over the time scale in which trypanosomes appear to have diverged) genes were used to screen for and elucidate relationships among trypanosome species associated with quokka (Setonix brachyurus) populations (vulnerable, small macropodid marsupials) in Western Australia.

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6.2 Materials and Methods

6.2.1 Study site and sample collection

Blood samples examined in the present study were collected from quokkas from

Two Peoples Bay (34º 58’S, 118º 11’E) (TPB) near Albany, Bald Island (34º 55’ S,

118º 27’E) (BI) and Rottnest Island (32º 00’S, 115º 31’E) (RI): all sites are in Western

Australia. A total of 35, 34 and 41 blood samples were collected from quokkas captured at Two Peoples Bay, Bald Island and Rottnest Island, respectively (Table. 6.1.).

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Table 6.1. A list of quokka isolates captured from the three different geographical locations used in this study.

Two Peoples Bay Bald Island quokka Rottnest Island quokka isolates n=35 isolates n=34 quokka isolates n=41

Q1364-1356 Q1997-1961 RS1 Q1416-1407 Q1 RS2 Q1339-1364 Q2382-2373 RS3 Q1837-1464 (Q5) Q2522-2523 RS4 Q1996-1999 Q 2581-2582 RS5 Q1362 Q2351-2346 RS6 Q2399-2324 Q1947 RS7 Q1342-1338 Q2367-2332 RS8 Q1354-1368 (Q10) Q2377-2381 RS9 Q1369-1355 Q2363-2376 RS10 Q2088-2050 Q2383-2359 RS11 Q1403-1404 Q2578--2600 WS1 Q1382-1326 Q Q2933-3201 WS2 Q1340 Q3307 BS1 Q2031 Q3343-3304 BS2 Q1051 Q3355-3284 BS3 Q1767 Q3336-3325 BS4 Q1366 Q2385-2359 BS5 Q2990 Q3206-3207 BS6 Q4137-1443 Q3313-4458 BS7 Q4177-4185 Q3346-4462 BS8 Q4148-4151 Q3542-3543 BS9 Q4124-4119 Q3547-3548 BS10 Q4140-4178 Q4435-3539 BS11 Q4299-4300 Q3293-4460 BS12 Q4141-4193 Q4478-3340 BS13 Q4120-4129 Q4461-4497 BS14 Q4166-2325 Q4491-4489 BS15 Q4176-4133 Q4540-4539 WE2 Q4131-4159 Q3269-3268 WE3 Q4112-4117 Q3364-3201 WE4 Q4115-4192 Q3229-3228 WE5 Q4174-4102 Q3362-3317 WE6 Q4908-4846 Q3258-3257 WE7 Q4633-4613 WE8 WE9 WE10 WE11 WE12 WE13 WETL1

6.2.2 PCR amplification and sequencing of trypanosomes

For initial screening, a nested set of trypanosome primers was used to amplify a partial fragment (~1,500 bp) of a variable region of the trypanosome 18S rRNA gene as

113 previously described in chapter 2 (section 2.7.2). An additional set of internal nested primers TRYall 1 Forward 5’ ACCGTTTCGGCTTTTGTTGG 3’ and TVEG Reverse

5’ AAATCGTCTTCGCTTTAACTTT 3’ which amplify ~468 bp of the 18S rRNA trypanosome gene (this study) were used on the Rottnest Island quokka samples, as previous attempts to amplify these samples using the universal 18S rRNA primers were unsuccessful. If mixed sequencing chromatograms resulted, then PCR products were cloned and plasmid inserts re-sequenced as described in 2.8. A subset of isolates (n =

26) that were positive at the 18S rRNA locus were amplified using a hemi-nested PCR of the GAPDH gene (~880 bp) as previously described (McInnes et al., 2009).

6.2.3 Phylogenetic analysis

After alignment, an 841 bp region of the 18S rRNA gene and a 664 bp region of the GAPDH gene (based on available shorter trypanosome sequences from this study and in GenBank) were analysed using MEGA6 (http://www.megasoftware.net/)

(Tamura et al., 2007). After selection of the most appropriate evolutionary model

(Kimura-2-parameter) using Model-Test in MEGA 6 (Tamura et al., 2013), maximum parsimony (MP), maximum likelihood (ML) and distance analyses were conducted using MEGA 6. Bootstrap analyses were conducted using 1,000 replicates to assess the reliability of inferred tree topologies. Bayesian Inference (BI) analysis was carried out by MrBayes (http://mrbayes.sourceforge.net), using the default options. For 18S, the

Kimura 2-parameter + G (0.19) model was used and for GAPDH analysis, the GTR + G

(0.30) was chosen. Trees were visualized by FigTree v1.4.0 (http://tree.bio.ed.ac.uk/).

6.2.4 Statistical analysis

Prevalences were expressed as the percentage of samples positive by PCR, with

95% confidence intervals calculated assuming a binomial distribution using the software Quantitative Parasitology 3.0 (Rozsa et al., 2000).

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6.3 Results

6.3.1 Prevalence of trypanosomes in quokka populations

The overall prevalence of trypanosomes in quokkas was 57.3% (48-65.5% CI)

(63/110) and varied from 4.9% on RI to 91.4% at TPB (Table 6.2.).

Table 6.2. Prevalence of T. copemani genotypes in quokka isolates from Two Peoples

Bay (TPB), Bald Island (BI) and Rottnest Island (RI) as determined by PCR and sequencing of partial 18S rRNA fragments. The 95% confidence intervals are given in parentheses. Mixed infections were determined by cloning and sequencing of amplicons producing mixed sequencing chromatograms.

Two Peoples Bay Bald Island Rottnest Island (TPB) (BI) (RI)

Overall 91.4% - 32/35 85.3% - 29/34 4.9% -2/41 Prevalence of (82.2-100.7% CI) (73.4-97.2) (0-11.5) Genotypes: (PCR positive) T. copemani 23.8% - 5/21 41.7% -10/24 50% -1/2 genotype A (5.6-42) (21.9-61.4) (0-199.3)

T. copemani 52.4% -11/21 50% -12/24 50% -1/2 genotype B (31-73.7) (30-70) (0-199.3)

Mixed T. copemani 23.8% - 5/21 8.3% -2/24 - genotype A and B* (5.6-42) (0-19.4)

6.3.2. Molecular characterisation of trypanosomes in quokka isolates at the 18S rRNA locus

Of the 32, 29 and 2 PCR positives at the 18S rRNA locus obtained from quokkas from Two Peoples Bay, Bald Island and Rottnest Island, respectively, sequences were obtained for 21, 24 and 2 positives. Sequence and phylogenetic analysis of these positive partial fragments of the 18S rRNA gene showed that all the trypanosome isolates from the quokkas were T. copemani (grouping into either T. copemani genotype A or B clades) (Table 6.2). The prevalence of T. copemani genotype 115

B was higher for both Two Peoples Bay and Bald Island compared to T. copemani genotype A (Table 6.2). The phylogenetic relationship of T. copemani to other trypanosome species at the 18S rRNA gene was analysed using Maximum Parsimony,

Maximum Likelihood, Distance and Bayesian analyses (Figure 6.1,Bayesian tree shown). For ease of phylogenetic analysis, two T. copemani genotype A representative

18S rRNA sequences from quokka isolates Q2031 from Two People’s Bay (TPB) and

Q3336-3325 from Bald Island (BI) (100% identity to all genotype A sequences) and two representative genotype B isolates from quokka Q1051 from TPB and quokka

Q4471-3340 from BI (100% identity to all genotype B sequences) were included in the tree (Figure 6.1), although phylogenetic analysis was conducted on all isolates.

Phylogenetic analysis produced trees with similar topologies with two distinct genotypes identified within T. copemani (Figure 6.1, Bayesian tree shown). The quokka isolates clustered within T. copemani genotypes A and B. Genotype B (represented by quokka isolates Q1051 - TPB and Q4478-3340 - BI) formed a separate clade consisting of trypanosomes isolated from quokkas and a T. copemani sequence from a woylie

(KC753530) previously designated as G1 by Botero et al (2013) which exhibited 1.9% genetic distance from T. copemani genotype B. Genotype A (represented by quokka isolates Q2031 - TPB and Q3336-3325 - BI) showed 100% identity with a T. copemani isolate from a woylie (KC753531) previously referred to as G2 by Botero et al. (2013), and also grouped with sequences from wombats (AA1- AJ620559, H26-AJ009169 and

APP- AJ620558) (Noyes et al., 1999). Trypanosoma copmani from koalas (GU966585-

GU966586 and GU966588) (McInnes et al., 2010) formed a separate clade. The quokka

T. copemani genotypes A and B were 1.8% distant from each other. Trypanosoma copemani genotype C (represented by wombat isolates AAI, AAP, H26) as previously described in chapter 3 was not supported in this study at the 18S rRNA locus, when analysing larger datasets

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Trypanosoma vegrandis isolates from woylies (G3-G7) (Botero et al., 2013;

Thompson et al., 2013) grouped with woylie isolates from an earlier study (Paparini et al., 2011). Considerable genetic variation was evident amongst T. vegrandis isolates with genetic distances of 0.4-3.2% within T. vegrandis isolates, with the largest distance of 3.2% between G3 (KC753533) and G4 (KC753532)/G7 (KC753536).

Another trypanosome sequence from a woylie (KC753537), previously designated as G8 by Botero et al. (2013), grouped with a trypanosome isolate (H25A) from a kangaroo (J009168), consistent with previous analysis by Botero et al. (2013).

Both the woylie and kangaroo isolates were genetically distinct from all other marsupial-derived trypanosome isolates and exhibited 0.7% genetic distance from each other, 8.2-8.6% genetic distance from T. cruzi and up to 26.9% genetic distance (G4 -

KC753532) from other marsupial-derived trypanosome isolates.

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Figure 6.1. Phylogenetic relationships of T. copemani genotype A and B quokka isolates (underlined this study) from Two Peoples Bay (TPB), Bald Island (BI) and Rottnest Island (RI) using the Bayesian Inference method of partial (841 bp) 18S rRNA sequences. Posterior probabilities are indicated on the main branches. Quokka isolates representing T. copemani genotype A (#) and T. copemani genotype B (*) are listed. Isolates designated ** were mixed T. copemani genotype A and B infections. The scale bar represents the proportion of base substitutions per site.

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6.3.3 Phylogenetic analysis of trypanosome isolates from quokkas at the

GAPDH gene

Sequences were obtained for 26 quokka isolates at the GAPDH locus.

Phylogenetic analysis of trypanosome partial GAPDH sequences was conducted using using Maximum Parsimony, Maximum Likelihood, Distance and Bayesian analyses

(Figure 6.2 - Bayesian tree shown). Four genotype A representative T. copemani

GAPDH sequences (from quokka isolates Q2031, Q3336-3325, Q1837-1464 and

QBS5) and four genotype B quokka isolates (Q2367-2332, Q4478-3340, Q2088-2050 and Q1051) were included in the tree although phylogenetic analysis was conducted on all 26 isolates.

In this analysis, quokka isolates from T. copemani genotype B formed their own clade while T. copemani genotype A quokka isolates clustered with koala and wombat isolates and the Rottnest quokka (QBS5). In contrast to the 18S rRNA analysis, genotype A quokka isolates also grouped with a G1 sequence (woylie), which had grouped in a clade at the 18S rRNA locus with T. copemani genotype B. One T. copemani isolate from a woylie representing G2 (KC812983), which grouped with genotype A at the 18S r RNA locus, formed a separate clade at the GAPDH locus. The genetic distances between T. copemani genotypes A and B was 1.8% at the GAPDH locus. The genetic distance between the sequence represented by G2 (KC812983) and T. copemani genotypes A and B was 2.9% and 3.1% respectively.

One isolate, Q4112-4117 from Two Peoples Bay, which grouped with T. copemani genotype B clade at the 18S rRNA locus (Figure 6.1), grouped with a very different clade at the GAPDH locus. This clade included a trypanosome sequence from a woylie (KC812988), previously designated as G8 by Botero et al. (2013), the trypanosome isolate (H25) from a kangaroo (AJ620276) and sequences from brushtail possums (JN315395 and JN315396) (Noyes et al., 1999; Hamilton et al., 2004; Paparini 119 et al., 2011). Quokka isolate Q4112-4117 from Two Peoples Bay was genetically distinct and exhibited 3.9% genetic distance from kangaroo isolate H25 and 1.8% genetic distance from brushtail possums (JN315395 and JN315396). The genetic distances between Q4112-4117 and H25 (AJ620276) from T. cruzi at the GAPDH locus were 11.1% and 11%, respectively. The T. vegrandis clade again exhibited considerable within species genetic diversity with the largest genetic distance (19.2%) between T. vegrandis isolates G3 (KC812984) and G4 (KC812985).

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Figure 6.2. Phylogenetic relationships of T. copemani genotype A and B quokka isolates (underlined this study) from Two Peoples Bay (TPB), Bald Island (BI) and Rottnest Island (RI) using the Bayesian Inference method based on partial (641 bp) GAPDH sequences. Posterior probabilities are indicated on the main branches. Quokka isolates representing T. copemani genotype A (#) and T. copemani genotype B (*) are listed. Isolates designated ** were mixed T. copemani genotype A and B infections. The scale bar represents the proportion of base substitutions per site

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6.4 Discussion

In the present study, molecular analysis was used to identify the genotypes of trypanosomes infecting Western Australian quokkas from three different geographical regions: Two Peoples Bay, Bald Island and Rottnest Island in Western Australia.

Phylogenetic analysis at the 18S rRNA locus found that all the quokka isolates grouped within the T. copemani marsupial clade together with isolates from wombats, koalas, woylies and a common brushtail possum, demonstrating that T. copemani is infective to a wide variety of Australia marsupials. Two main T. copemani genotypes were identified at the 18S rRNA locus T. copemani genotype A (represented by quokkas

Q3336-3325, Q2031) and T. copemani genotype B (represented by quokkas Q4471-

3340, Q1051). Trypanosoma copemani genotype C (represented by wombat isolates

AAI, AAP, H26) as previously described in chapter 3 was not supported in this study at the 18S rRNA locus, when analysing larger data sets.

Trypanosoma copemani genotype B was the most prevalent genotype infecting quokka populations from TPB and BI and has previously only been isolated from quokkas and the Gilbert’s potoroo. At the 18S rRNA locus, T. copemani genotype A isolates from quokkas were 100% identical to the G2 isolate from a woylie (Botero et al., 2013) indicating that genotype A is prevalent in multiple marsupial species, with G2 previously detected in woylies, a quoll and a quenda or southern brown bandicoot

(Botero et al., 2013).

In contrast to the 18S rRNA analysis however, the G2 genotype (KC812983) at the GAPDH locus grouped separately from T. copemani genotypes A and B which suggests that the G2 genotype may have been a mixed infection consisting of two genotypes, with different genotypes amplified at the 18S rRNA and GAPDH locus.

Most species of trypanosomes reproduce predominantly by clonal evolution (Telleria et al., 2004), which over time gives rise to multilocus genotypes (individuals display

122 exactly the same genotype at all loci) (Koffi et al., 2015) due to very rare genetic exchange. The observation that G2 grouped into different clades at different loci, demonstrates the multilocus genotypes of trypanosomes given that the topologies of both phylogenetic trees were similar with the exception of the G2 isolate.

A mixed infection in quokka isolate Q4112-4117 from Two Peoples Bay was also identified as this isolate grouped with T. copemani genotype B clade at the 18S rRNA locus (Figure. 6.1), but grouped with a clade by itself but closest to a clade that included the G8 genotype (Botero et al., 2013), as well as isolates from a kangaroo

(H25 - AJ620276) and brushtail possums (JN315395 and JN315396) at the GAPDH locus (Figure. 6.2). It has been previously reported that the G8 genotype is closely related to T. cruzi at the 18S locus (Stevens et al., 1999b; Paparini et al., 2011; Hamilton et al., 2012; Botero et al., 2013) however in the present study the genetic distance between this clade and T. cruzi at the 18S locus was 8.2-8.6%. At the GAPDH locus, the genetic distance was larger (11-11.1%), in agreement with a previous study by

Botero et al. (2013). This phenomenon of mixed trypanosome infections in native

Australian marsupials has previously been reported in woylies and koalas (McInnes et al., 2010; Botero et al., 2013; Barbosa et al., 2016) and highlights the need for species- specific primers and/or deep sequencing to identify the true level of mixed trypanosome infections in marsupials. For example, a recent study demonstrated that T. vegrandis was only detected in koalas when species-specific primers were used (Barbosa et al.,

2016). Identifying mixed infections has important clinical implications as mixed T. copemani and T. vegrandis infections are found more frequently in declining woylie populations compared to stable groups of woylies (p = 0.001) (Botero et al., 2013). Both

T. copemani genotype G2 (which corresponds to T. copemani genotype A) and T. vegrandis have been identified within the internal organs of the woylie, where they are thought to adversely affect the fitness and coordination of the host, thus increasing their

123 susceptibility to predation (Thompson et al., 2014b). In chapter 5, erythrocyte abnormalities, including microspherocytes and schistocytes in T. copemani infected quokka blood (chapter 5), were identified which are changes typically associated with haemolytic anaemias (Mallah et al., 2010; Rodak et al., 2012). This suggests that T. copemani genotype A may be a cause of anaemia in marsupials and it is also possible that T. copemani may be the cause of seasonal anaemia and low red blood cell counts in quokkas from Rottnest Island (Barker et al., 1974). However further clinical investigations are needed to determine the clinical impact of T. copemani on quokka populations.

Large genetic distances were evident within T. vegrandis isolates at both the 18S rRNA locus (0.4-3.2%) and the GAPDH locus (up to 18.5%). This indicates that T. vegrandis is not a uniform species but is in fact a species complex. Trypanosomes have few morphological features detectable using light microscopy which can adequately delimit species (Gibson, 2009) and the very small size of T. vegrandis (8.3 µm in length and 1.3 µm in width), renders this even more difficult. Previous studies have reported that a genetic distance of 3.75% at the GAPDH gene may be sufficient to delimit a new trypanosome species (McInnes et al., 2011). By this criterion, there are multiple valid species within T. vegrandis, which need to be characterised more fully in the future.

In conclusion, T. copemani consists of multiple genotypes, of which genotype A has previously been associated with pathogenic effects in woylies. Mixed infections were common and further work is required to determine the clinical impact of T. copemani on marsupial populations.

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CHAPTER 7

THE INNATE RESISTANCE OF TRYPANOSOMA

COPEMANI TO HUMAN SERUM

Austen, J. M., Ryan, U. M., Ditcham, W. G. F., Friend, J. A. and Reid, S. A. 2015. The innate resistance of Trypanosoma copemani to human serum. Experimental

Parasitology. 153, 105-110.

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7.1 Introduction

Only a few trypanosome species are known to cause disease in mammals. These include the etiological agents of animal trypanosomiasis (T. congolense, T. vivax, T. brucei brucei, and T. evansi) and human African trypanosomiasis (T. b. rhodesiense and

T. b. gambiense), as well as T. cruzi, which causes Chagas’ disease in the Americas

(Hoare, 1972). Trypanosoma rangeli has also been detected in humans but it is not considered to be pathogenic (Stevens and Brisse, 2004). Trypanosoma rangeli and T. cruzi share a large number of vertebrate reservoirs including animals in five orders:

Edentata, Marsupialia, Carnivora, Rodentia and Primates (Guhl and Vallejo, 2003).

Trypanosoma brucei rhodesiense and T. b. gambiense are resistant to the cytotoxic action of normal human serum because they are resistant to the trypanosome lytic factors (TLFs) that are naturally present in the blood of all humans and primates

(Milner and Hajduk, 1999; Molina-Portela Mdel et al., 2005). Resistance to TLFs is conferred by the presence of the serum resistance associated gene (SRA) in T. b. rhodesiense (De Greef and Hamers, 1994; Xong et al., 1998). In T. b. gambiense, an

SRA-like protein that is specific to T. b. gambiense (TgsGP) has been shown to be essential for human serum resistance (Capewell et al., 2013). The resistance of T. cruzi to human serum is based on resistance to the alternative complement pathway (ACP)

(Cestari and Ramirez, 2010). However, not all of the life-cycle stages of T. cruzi are resistant to direct serum lysis. For example, the epimastigote life-cycle stage derived from the vector’s gut is efficiently lysed in human serum, but the trypomastigote and amastigote stages from the vertebrate host are not (Tomlinson and Raper, 1998).

Trypomastigotes of T. cruzi resist lysis because they are able to prevent the initiation of the complement cascade by expression of complement system inhibitors (Joiner et al.,

1988).

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The blood incubation infectivity test (BIIT) developed by Rickman and Robson

(1970) was originally designed to distinguish the human-infective T. b. rhodesiense from the non-human-infective T. b. brucei. This test is based on the observation that non-human-infective trypanosomes are lysed by human serum while human-infective trypanosomes resist lysis. Therefore, this test can provide a method to identify new species of trypanosomes that are potentially human-infective (Turner et al., 2004). The present study was performed to determine the relative susceptibility of T. copemani to human serum.

7.2 Materials and Methods

7.2.1 Study site and sample collection

A quokka (Q2088), previously captured and identified as positive for T. copemani by PCR, was trapped at Two Peoples Bay. Blood was collected as described in chapter 2 (section 2.3.1). Fifty microliters (μL) of blood were used for the BIIT.

Thin-blood smears were prepared from 10 µL of blood and the number of trypanosomes counted to determine the initial numbers of trypanosomes / mL of blood. A haemocytometer was used to calculate the number of trypanosomes present in culture.

7.2.2 DNA extraction

Whole genomic DNA was extracted from both fresh blood samples and cultured trypanosomes as previously described in chapter 2 (section 2.7.1).

7.2.3 Molecular characterisation of T. copemani

To confirm that the trypanosomes isolated from quokka Q2088 were T. copemani, a 1,439 bp fragment of the 18S ribosomal RNA (rRNA) gene and an 841 bp fragment of the GAPDH gene were amplified and sequenced as previously described in chapter 2 (sections 2.7.2 and 2.7.3).

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7.2.4 In vitro human serum resistance: blood incubation infectivity test

Serum prepared from fresh whole blood from healthy human volunteers was used during this experiment. The use of human subjects for this study was approved by

Murdoch University human ethics committee (project number 2010/053).

The BIIT was performed in triplicate using serum freshly prepared from the blood of five healthy human volunteers as the test samples (labelled HA, HB, HC, HD and HE) and from horse blood as the control samples (labelled H0A, H0B and H0C).

Five sets of MSEM (Noyes et al., 1999), used for the growth of the trypanosomes following the BIIT challenge were made up with each sample of the homologous human blood (HuMSEM) along with three sets of horse blood MSEM (HoMSEM). The BIIT was performed by adding a 50 µL aliquot of fresh quokka blood (containing ~five trypanosomes) into separate tubes, each containing 250 μL of one of the five samples of fresh undiluted human serum, or horse serum. These tubes were then incubated in a water bath at 37°C for 5 hours. The entire contents of each incubated tube were then added to individual tubes containing 1 mL of HuMSEM or HoMSEM, and incubated at room temperature in the dark for 24 hours before examination. In addition, a 50 μL control sample of fresh quokka blood was directly transferred into a HuMSEM and a

HoMSEM without the initial incubation in serum, and incubated at room temperature in the dark. Microscopic examination of wet-smear preparations from each culture was performed every day at 200× and 400× magnification to detect the presence of motile trypanosomes. If trypanosomes were detected, Giemsa-stained thin blood smears were prepared for further microscopic examination.

On day 14 when high numbers of motile trypanosomes were observed in

HuMSEM, 100 μL volumes of the culture (approx. 7.5 x 107 organisms) were transferred into 1 mL of HuMSEM with a 0.5 mL overlay of RPMI 1640 supplemented with 10% horse serum, 1,000 IU/mL of ICN penicillin and 10,000 MCG/mL of

128 streptomycin. Cultures were incubated at 37ºC with 5% CO2 to mimic mammalian conditions and determine survival of T. copemani in liquid culture.

7.2.5 Statistical analysis

The relative rate of replication of T. copemani in HuMSEM and HoMSEM after the BIIT was assessed by performing triplicate counts on three human cultures (HA,

HB, HC) and 1 horse culture (H0C) at several time points after inoculation using a haemocytometer counting chamber. Half of the minimum level of detection (300) was added to each of the triplicate counts of T. copemani in HuMSEM and HoMSEM after the BIIT and log10 transformed. The mean log-transformed count was plotted against time for each of the 4 cultures (HA, HB, HC and H0C). Non-linear regression was performed on the log-transformed counts from each culture using the ‘plateau followed by one phase association’ function in GraphPad Prism version 5.00 for Windows

(GraphPad Software, San Diego, USA). The counts recorded for the human serum culture HA on days 18 and 19 were omitted from the analysis to improve the goodness of fit. Outputs of the regression analysis include: day of initiation of exponential growth, maximum trypanosome density and replication rate and their 95% confidence intervals. Values were considered significantly different if the 95% CI’s for each output variable from the regression analysis did not overlap.

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7.3 Results

7.3.1 Microscopy

Blood from quokka Q2088 contained approximately 100 trypanosomes per mL.

The morphology of the trypanosomes detected in the blood smear was consistent with the trypomastigote life-cycle stage (Figure 7.1) (Hoare, 1972).

Figure 7.1. Light micrograph of T. copemani in a Modified Wright stained blood film from quokka Q2088. Scale bar represents 10µm.

7.3.2 Molecular characterisation of T. copemani

The identity of the trypanosome infecting quokka Q2088 was confirmed as T. copemani (genotype B) using sequence analysis of both the 18S rRNA and GAPDH genes before and after the BIIT test. The 18S rRNA (1,439 bp) and GAPDH (841 bp) gene sequences of the quokka trypanosome isolate were found to be 100% identical to reference T. copemani sequences both before and after the BIIT challenge, and therefore were deposited in the GenBank database under the accession numbers (18S rRNA)

HQ267094 and (GAPDH) HQ267095.

7.3.3 Blood incubation infectivity test (BIIT)

Trypanosoma copemani isolated from quokka Q2088 multiplied successfully in

MSEM containing either human or horse blood, after undergoing the BIIT. There was

130 no significant difference in the time to initiation of exponential growth in cultures containing human blood or horse blood and trypanosomes were detected in all cultures by day 5. Plots of the log-transformed counts of T. copemani in cultures containing serum from 3 humans and 1 horse are presented (Figure 7.2). Motile T. copemani was also observed in both the control HuMSEM and HoMSEM inoculated with blood taken directly from the quokka.

Figure 7.2. Mean log -transformed count (plus half minimum detection level) of T. copemani 10 grown in cultures containing serum from 3 humans (HA, HB, HC) and one horse (H0C). The trend line represents the mean log-transformed count and markers () represent individual counts.

Unfortunately complete growth curves for T. copemani directly inoculated into

HuMSEM and HoMSEM from infected blood were not possible due to the delayed detection of trypanosomes in the first 9 days. Numbers of trypanosomes/mL were, however, measured at several time points from day 10 with T. copemani shown to multiply at a slightly faster rate in HuMSEM at days 10, 12 and 14 compared to the growth rate of T. copemani in HoMSEM (Figure 7.3). On day 17, the numbers of

131 trypanosomes in HuMSEM dropped compared to that detected in HoMSEM (Figure

7.3).

Figure 7.3. Numbers of trypanosomes / mL measured at several time points after direct inoculation of infected quokka blood into HuMSEM and HoMSEM.

The first trypanosome life-cycle stages to be detected in both HuMSEM and

HoMSEM post-BIIT were slender, rapidly moving trypomastigotes and thin epimastigotes (Figure 7.4 A), with the former life-cycle stage being the most abundant.

Larger epimastigotes and sphaeromastigotes (Figure 7.4 B) were detected by day 10, but in fewer numbers than the trypomastigotes and epimastigotes. There were no obvious morphological differences between the epimastigote or trypomastigotes life- cycle stages seen following culture in either human or horse blood.

Motile T. copemani was observed in the subcultures grown in human MSEM overlayed with RPMI that were incubated at 37°C. Trypomastigotes were the most abundant life-cycle stage detected with similar morphology to the trypomastigotes observed in quokka blood (Figure 7.4 C).

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Figure 7.4. Light micrographs of T. copemani grown in vitro, in a culture of blood from the quokka, which had undergone the blood incubation infectivity test. (A) Epimastigote (e) and trypomastigote (t) forms at day 10, in a Modified Wright stained smear from HuMSEM. (B) Sphaeromastigotes (s), epimastigotes and trypomastigote forms at day 10, in a Modified Wright stained smear from HuMSEM. (C) Trypomastigote form of T. copemani at day 18, grown in HuMSEM overlaid with RPMI at 37ºC. Scale bars represents 10µm.

7.4 Discussion

This study is the first to assess the susceptibility of an Australian mammalian trypanosome to human serum. Results from comparing the growth curves of T. copemani in cultures containing human and horse serum showed that there was no consistent effect on the trypanosome attributable to either host species. Trypanosoma copemani was detected as early as day 5 after the BIIT challenge, compared to direct inoculation into either HuMSEM or HoMSEM, in which trypanosomes were first detected on day 10. This suggests a faster rate of multiplication with the addition of liquid serum allowing the trypanosomes to readily gain access to the required nutrients compared to the solidified blood agar medium. The use of solidified MSEM by Noyes et al. (1999) to detect novel trypanosome species isolated from the blood of a wombat and kangaroo shows similar findings with trypanosomes first detected on days 13 and

20, respectively. Overall the growth of T. copemani inoculated directly into HuMSEM and HoMSEM appeared to be similar with only a slight increase in multiplication observed initially in the HuMSEM. Molecular analyses of the 18S rRNA and GAPDH

133 genes were conducted to confirm that the trypanosomes isolated from the blood of quokka Q2088 were T. copemani.

The BIIT is designed to enable the identification of potentially human infective trypanosomes. In previous studies (Rickman and Robson, 1970; Hawking, 1978), the

BIIT protocol included inoculation of human serum-exposed trypanosomes into rats and mice to determine viability. Inoculation of rodents was not done in the present study because early attempts to infect rodents with T. copemani were unsuccessful. Failure of rodent infectivity with T. copemani may have resulted in the lack or low prevelence of metacyclic trypanosomes within the inoculum at the time of infection. For natural infections to occur, multiple bites are needed from Glossina spp. for example, to initiate trypanosome infection (Hoare, 1972). The immune response of the host may have also overcome the initial inoculum of T. copemani, however, future studies are required to confirm this and to also understand the infective stages of T. copemani and their modes of transmission.

The in vitro methodology used in this study was adapted from a previous method (Tomlinson et al., 1995) where an in vitro assay was used to analyse the human serum resistance of various T. brucei genotypes. To reduce the likelihood of false positive results, only fresh whole human blood was used in this study to eliminate the potential effect an anticoagulant may have had on either the complement system or the human serum high density lipoprotein (HDL) trypanolytic factor. This is important because it has been shown that lipoprotein lipase activity and the level of HDL in rats infected with T. b. gambiense were increased if heparin was administered parenterally, which caused a reduction in the number of trypanosomes in the rat (Nishimura et al.,

2005). Furthermore, in vitro culture of T. b. gambiense was inhibited by the addition of plasma from infected rats treated with heparin (Nishimura et al., 2005). In contrast, the commonly used anticoagulant EDTA was found to inhibit the trypanolytic action of

134 normal human serum on T. congolense TC35U (Ferrante and Allison, 1983). This uncoated trypanosome is normally lysed by human serum but is resistant to lysis by human serum containing EDTA. In 2006 T. evansi was identified within a human patient that lacked Apolipoprotein L-1 (APOL1), a human specific protein that binds to

HDL (Vanhollebeke et al., 2006). Together APOL1 and HDL cause the osmotic swelling of T. brucei and eventual death (Vanhollebeke et al., 2006). The potential role of APOL1 in the results of this study were not investigated because trypanolytic factors

(TLFs), of which APOL1 is a major component, have been shown to have no effect on the replication of T. cruzi in an in vivo model (Samanovic et al., 2009). This is important because T. copemani shares morphological similarities with T. cruzi (Botero et al., 2013). For example, studies by Botero et al. (2013), found T. copemani in various marsupial tissues, and has described T. copemani as having a life-cycle similar to T. cruzi with T. copemani trypomastigotes having the ability to infect cells in vitro and transform into amastigotes (Botero et al., 2015).

The low numbers of trypomastigotes (100 trypanosomes/mL of blood) detected in the quokka’s blood at the time of sampling resulted in exposure of only about five trypomastigotes to human serum in the BIIT and subsequent culturing in HuMSEM.

The low inoculum is both representative of a natural challenge, and also ensures that any potential defence mechanisms present in the blood samples being used in the test are not overwhelmed by a large inoculum, giving a spurious apparent survival of trypanosomes. The low number of trypanosomes used in the present study is in contrast to both the large inocula of 2 x106 T. congolense exposed to 50% NHS by Xong et al.

(2002), and of 1x107 T. b. brucei exposed to 25% NHS by Turner et al. (2004). These studies (Xong et al., 2002; Turner et al., 2004) also used diluted serum in contrast to the undiluted normal human serum used in the present study, which shows that T. copemani blood life-cycle stages have a high level of resistance to human serum. The low

135 inoculum used in the present study may have accounted for the initial lag phase in growth, which prevented the detection of viable parasites in the first four days of the

BIIT. Detection of T. copemani was first possible on day five, a time lag which may have allowed the parasites to efficiently multiply and adapt to the vector life-cycle stages, given that their maintenance in vitro is a methodology used to mimic the conditions of the vector (Hoare, 1972).

The observation that T. copemani is able to survive in the presence of human serum is interesting but must be interpreted cautiously. Survival of trypanosomes when subjected to the BIIT test is strongly correlated to pathogenicity in studies of T. b. brucei (Rickman and Robson, 1970). In addition, T. lewisi for instance, which is considered a rodent trypanosome (Hawking, 1978), has been reported to infect humans on eight occasions (Lun et al., 2009). Similarly, T. evansi which is responsible for a widely distributed disease called ‘‘surra’’ in domestic and wild animals found in Asia,

Africa, South America, and even Europe, has been identified in humans on four occasions (Lun et al., 2009). However, survival in the BIIT test does not necessarily correlate with an ability to infect humans and cause disease. Furthermore, the risk of human infection with T. copemani would require interaction between the marsupial hosts, their putative tick vector and susceptible humans. Trypanosoma copemani does, however, have a broad marsupial host range, and increasing human encroachment on marsupial habitats, where ticks coexist with their natural marsupial hosts, may increase the risk of humans becoming infected with T. copemani.

The mechanism of resistance to NHS by T. copemani is unclear. If the mechanism is similar to T. cruzi, then the trypomastigotes may be able to inhibit the assembly, or accelerate the decay of C3 convertase, the central enzyme of the complement cascade (Tomlinson and Raper, 1998). Chronic, non-pathogenic infection in the quokka may be maintained by the production of antibodies, which render the

136 trypomastigotes sensitive to lysis via the alternative complement cascade, as occurs in mammalian hosts infected with T. cruzi (Krautz et al., 2000). There are significant morphological similarities between the life-cycle stages observed in T. copemani cultures and blood smears compared to T. cruzi. Therefore, it should not be surprising that blood-stream trypomastigote stages of T. copemani are resistant to human serum.

This is because trypomastigotes and amastigotes of T. cruzi from the vertebrate host are resistant to direct serum lysis from the complement system and epimastigotes from the gut of the vector are not resistant (Tomlinson and Raper, 1998; Krautz et al., 2000). It has been shown that cultured trypomastigotes of T. cruzi can form into both extracellular and intracellular amastigotes, both of which are infective to human monocytes in vitro, and may help to maintain the T. cruzi mammalian life-cycle (Ley et al., 1988). Amastigote stages of T. copemani have been observed in culture and in quokka blood (chapter 5), and may contribute to the resistance of T. copemani to human serum lysis.

It is tempting to draw bold conclusions from the results of this study that T. copemani, and possibly other Australian trypanosomes, represent an extensive and latent pool that could give rise to new emerging infectious diseases (EIDs). Indeed, approximately 75% of EIDs that have affected human populations in the past 30 years have been zoonotic (Daszak et al., 2007). In addition to T. copemani, Australian trypanosomes from kangaroos (H25), possums (Pseudocheirus peregrinus), woylies, a banded hare wallaby (Lagostrophus fasciatus) and boodies (Bettongia lesueur) have closer phylogenetic relationships with T. cruzi than T. copemani (Noyes et al., 1999;

Stevens et al., 1999a; Paparini et al., 2011; Hamilton et al., 2012; Botero et al., 2013). In addition, there are a small but significant number of atypical human infections with

“animal” trypanosomes that raises the possibility that many human infections remain undiagnosed (Lun et al., 2009; Truc et al., 2013). Trypanosoma cruzi is principally a

137 parasite of sylvatic animals and it did not undergo the prolonged period of co-evolution with humans experienced by the T. brucei group. Humans became a host only when they became ‘available’ in the sylvatic life-cycle of T. cruzi ~9,000 years ago, which corresponds with the period when humans developed settled (rather than nomadic) populations. The presence of these sedentary populations and their dwelling places altered the life-cycle of the vector, creating a new non-sylvatic cycle and a new ‘human’ disease (Aufderheide et al., 2004). Therefore, the observation that T. copemani is able to resist NHS is insufficient to conclude that it has significant zoonotic potential per se.

Rather, a more complex chain of events would have to occur to significantly alter the life-cycle of the parasite and its mammalian hosts. It may also be possible that T. cruzi is unique in the trypanosome world in that the vector and sylvatic cycle contained the correct ingredients for establishment in the “human” domestic environment. Whilst the human population of Australia is unlikely to undergo a significant societal change equivalent to South America 9,000 years ago, Australians live in relatively close proximity to native marsupials and the level of exposure to marsupial-derived tick vectors is unknown. It is, therefore, possible that isolated cases of human trypanosomiasis may occur in the Australian population. If this does occur, then information on the susceptibility of native trypanosomes to currently available trypanocidal drugs would be of enormous value.

138

CHAPTER 8

FURTHER CHARACTERISATION OF TWO

EIMERIA SPECIES (EIMERIA QUOKKA AND

EIMERIA SETONICIS) IN QUOKKAS (SETONIX

BRACHYURUS)

Austen, J.M., Friend J.A., Yang, R., Ryan, U.M. 2014. Further characterisation of two

Eimeria species (Eimeria quokka and Eimeria setonicis) in quokkas (Setonix brachyurus). Experimental Parasitology 138, 48-54

139

8.1. Introduction

Within the genus Eimeria, there are hundreds of different species, which cause disease in birds, reptiles and mammals (Barker et al., 1988; Cox, 1998; Power et al.,

2009). Pathogenic species of Eimeria, which commonly cause morbidity and mortality, are a major concern for the agriculture community, resulting in economical losses within the livestock and poultry industries (Barta et al., 1997; Cox, 1998; Aarthi et al.,

2010).

In native mammals, over 50 Eimeria species have been described from a range of marsupial hosts in Australasia and the Americas including kangaroos, wallabies, wombats, quokkas, possums, bandicoots and opossums (Mackerras, 1959; Barker et al.,

1988, 1989; Heckscher et al., 1999; Bennett et al., 2006; Power et al., 2009; Hill et al.,

2012). Description of these native Eimeria species has largely been based on morphological characterisation by determining the size and shape of sporulated oocysts and sporocysts, as well as by various other distinguishing characteristics (Mackerras,

1959; Barker et al., 1989; Duszynski and Wilber, 1997). The geographical location as well as the host species has also been used to aid in the classification of novel Eimeria species. However it has been shown that many Eimeria species are not strictly host- specific and can naturally have very large geographical ranges (Duszynski and Wilber,

1997).

Molecular characterisation coupled with the more traditional morphological classification methodologies results in a more robust taxonomie system and provides more information on evolutionary relationships between Eimeria species. The gene of choice commonly used for aiding in the evolutionary diversity of Eimeriidae is the 18S rRNA gene due to its conserved nature throughout the eukaryotes (Stevens et al.,

1999a). More recently the cytochrome c oxidase subunit 1 (COI) gene has been used because it has higher resolving power for Eimeria spp., especially with respect to recent

140 speciation events (Ogedengbe et al., 2011) and is the target gene of choice for DNA bar coding of Eimeria spp. (Hebert et al., 2003).

There is limited information on the evolutionary relationships of Eimeria species isolated from Australian marsupials. To date only Eimeria trichosuri, a species found in brushtail possums of the genus Trichosurus, E. macropodis from tammar wallabies

(Macropus eugenii) and several unidentified macropod Eimeria species have been genetically characterised (Power et al., 2009; Hill et al., 2012; Yang et al., 2012). A previous study by Barker et al. (1989) morphologically described three Eimeria species;

E. volckertzooni, E. quokka and E. setonicis isolated from quokkas. However, no molecular analysis was undertaken. In the present study, we characterised two Eimeria species from quokkas, which conformed morphologically to E. quokka and E. setonicis, at two loci (18S rRNA and COI) and determined their evolutionary relationships to other marsupial-derived Eimeria species to expand our knowledge of the diversity and evolution of marsupial-derived Eimeria species.

8.2. Materials and Methods

8.2.1 Sample collection

Fresh voided faecal samples were collected from 8 quokkas captured at Two

Peoples Bay, 20 faecal samples were collected from quokkas from Bald Island and 23 faecal samples collected from quokkas on Rottnest (Table 8.1) and stored at 4ºC until required.

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Table 8.1. Quokka isolates in which faecal samples were collected from, for use in this study.

Two Peoples Bay quokka Bald Island quokka Rottnest Island quokka isolates n=8 isolates n=20 isolates n=23

4177-4185 3361-3360 RS3 4148-4151 4409-4408 RS4 4124-4119 2473-2414 RS6 4142-4193 3578-4517 RS7 4112-4117 4481-4250 RS9 4174-4102 4479-4480 WS2 4125-4169 4526-4563 BS1 1340-1956 3506-2965 BS3 2411-2410 BS4 2367-2332 BS5 4580-4584 BS10 2929-4443 BS11 4529-3548 BS12 2435-4533 BS13 3267-3266 BS14 4547-4546 WE2 4581-4503 WE3 4435-3539 WE4 4527-4578 WE5 3258-3257 WE6 WE8 WE9 WE10

8.2.2 Screening and morphological analysis of oocysts

To detect oocysts in quokka faeces, a zinc flotation method was used as previously described (Daugschies et al., 1999), combined with microscopic examination of wet mounts using a 400× magnification. Oocysts were confirmed as Eimeria by their oval appearance, presence of four sporocysts and thick cell wall. If oocysts were identified, the faecal samples were placed into 2% potassium dichromate and left at room temperature for sporulation to take place to aid morphological identification.

Sporulated oocysts were observed at 1,000× magnification using Nomarski or differential interference contrast (DIC) microscopy and measured with an ocular

142 micrometer. Images of oocysts were recorded using an Olympus DP71 Advance digital camera and Image J (Schneider et al., 2012).

8.2.3 DNA isolation and PCR amplification

Genomic DNA was extracted from 250 mg of each faecal sample using a Power

Soil DNA Kit (MolBio, Carlsbad, California) according to the manufacturer’s instructions. Samples were screened using a nested PCR which amplified ~ 502 bp region of the 18S rRNA using primers as previously described (Eberhard et al., 1999).

Positive samples were also amplified using a hemi nested PCR to amplify a longer fragment of the Eimeria 18S rRNA gene (~ 1,300 bp) as previously described (Yang et al., 2012). Amplification of a 465 bp region of the COI locus from samples that were positive at the 18S rRNA locus was conducted as described by Ogedengbe et al. (2011).

8.2.4 Sequencing and phylogenetic analysis

PCR products were processed and purified as described in chapter 2 (2.7.5) and sequenced directly as described in chapter 2 (2.9).

Phylogenetic trees were constructed for Eimeria spp. at the 18S rRNA and COI genes with additional isolates from GenBank. MEGA5 (http://www.megasoftware.net/) was used for maximum-likelihood (ML) and Parsimony analyses (Tamura et al., 2007).

Distance estimation was conducted using TREECON (Van de Peer and De Wachter,

1994), based on evolutionary distances calculated with the Tamura-Nei model and grouped using Neighbour-Joining. Bootstrap analyses were conducted using 1,000 replicates to assess the reliability of inferred tree topologies.

8.2.5 Statistical Analysis

Prevalences were expressed as percentage of positive samples with 95% confidence intervals calculated assuming a binomial distribution, using the software

Quantitative Parasitology 3.0 (Rozsa et al., 2000).

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8.3. Results

8.3.1 Eimeria Prevalence

The prevalence of Eimeria oocysts in faecal samples isolated from quokkas captured from each of the three studied geographical locations (Two Peoples Bay, Bald

Island and Rottnest Island) by microscopy was 62.5% (5/8) (29.0 - 96.0, 95% CI), 45%

(9/20) (23.2 - 66.8, 95% CI) and 43.5% (10/23) (23.2 - 63.7, 95% CI), respectively. The prevalence by PCR (18S rRNA locus) for Two Peoples Bay, Bald Island and Rottnest

Island was 62.5% (5/8) (29.0 - 96.0, 95% CI), 85.0% (17/20) (69.4 - 100.6, 95% CI) and

78.3% (18/23) (61.4 - 95.1, 95% CI), respectively (Table 8.2).

Table 8.2. The prevalence (and 95% confidence intervals) of Eimeria in quokkas detected by both microscopy and PCR (18S rRNA locus) from three geographical locations.

Geographical No of Microscopy Microscopy PCR PCR % locations faecal positives % Positives prevalence samples prevalence 18S rRNA (95% CI) examined (95% CI) locus Two Peoples 62.5 (29.0- 62.5 (29.0- Bay 8 5 5 96.0) 96.0)

Bald Island 45.0 (23.2- 85.0 (69.4- 20 9 17 66.8) 100.6) Rottnest 43.5 (23.2- 78.3 (61.4- Island 23 10 18 63.7) 95.1)

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8.3.2 Morphological characterisation of Eimeria oocysts

Morphological identification of Eimeria species from quokkas captured at Two

Peoples Bay and Bald Island were determined from sporulated oocysts incubated in potassium dichromate (Table 8.3). Unfortunately the Eimeria oocysts from the Rottnest quokkas failed to sporulate when incubated. Eimeria oocysts were measured from three

(Q4125-4169, Q4177-4185, Q4124-4119) and five (Q4546-4547, Q2367-2332, Q4527-

4578, Q4547-4546, Q2929-4443) individual quokkas from Two Peoples Bay and Bald

Island, respectively, and two morphologically different species of Eimeria were identified. Based on the dimensions of the sporulated oocysts and sporocysts, along with other morphological characteristics, oocysts resembling E. quokka (Figure 8.1 A and B) and E. setonicis (Figure 8.1 C) were identified. In the present study, sporulated oocysts from both Two Peoples Bay and Bald Island conformed to the previously reported characteristics for E. quokka (Barker et al., 1988) with the exception that the oocyst length, which ranged from 13.7 - 22.5 µm and 12.5 - 23.7 µm respectively, was slightly larger (Table 8.3) and sporocysts ranged from 5.0 - 10.0 µm in length and 3.7 -

7.5 µm in width.

Figure 8.1. Nomarski interference-contrast micrographs of Eimeria oocysts isolated from quokka faeces (quokka isolate Q2929-4443) from Two Peoples Bay, Western Australia, resembling E. quokka based on morphological characteristics (A and B) and (quokka isolate Q4124-4119) from Bald Island, Western Australia, resembling E. setonicis based on morphological characteristics (C).

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Sporulated oocysts from both Two Peoples Bay and Bald Island conformed to E. setonicis characteristics but ranged in length from 26.2 - 37.5 µm and 28.7 - 35 µm and in width from 17.5 - 22.5 µm and 17.5 - 25 µm respectively and are therefore slightly larger than the previously published morphometrics for E. setonicis (Barker et al.,

1988). Sporocysts measured 7.5 - 13.7 µm in length and 7.5 - 10 µm in width (Table

8.3).

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Table 8.3. Morphometric (µm) comparison of Eimeria oocysts from quokkas* from Two Peoples Bay, and from quokkas● from Bald Island with previously published morphometrics for E. quokka and E. setonicis from Rottnest Island quokkas (Barker et al., 1988).

E. quokka E. quokka E. quokka E. setonicis E. setonicis E. setonicis Two Peoples Bay Bald Island Rottnest Two Peoples Bald Rottnest Island Bay Island Island

This study This study Barker et al., This study This study Barker et al., 1988 1988 n 20 58 45 11 12 26 Oocyst length Mean 18.3 18.0 18.0 29.7 31.9 29.9 SD 2.3 3.2 1.7 3.2 1.8 1.71 Range 13.7 - 22.5 12.5 - 23.7 13.6 - 21.6 26.2 - 37.5 28.7 - 35.0 26.4 - 33.6

Oocyst width Mean 11.8 12.2 10.8 19.5 19.5 17.9 SD 1.24 1.4 1.1 2.6 2.2 0.6 Range 10.0 - 15.0 10.0 - 15.0 8.8 -15.2 17.5 - 22.5 17.5 - 25 16.8 - 19.2

Sporocyst length Mean 7.0 6.4 7.7 12.1 11.0 12.0 SD 1.3 1.2 0.6 1.3 1.9 0.79 Range 5.0 - 10.0 5.0 - 8.7 6.4 - 9.6 8.7 - 13.7 7.5 -13.7 10.4 - 13.6

Sporocyst width Mean 5.0 5.0 5.0 8.5 8.8 7.7 SD 0.7 0.8 0.5 1.0 0.8 0.4 Range 3.7 - 7.5 3.7 - 7.5 4.0 - 6.4 7.5 -10.0 7.5 - 10.0 7.2 - 8.8

n=number, SD =standard deviation.

*(Q4125 - 4169), (Q4177 - 4185), (Q4124 - 4119), ●(Q4546 - 4547), (Q2367-2332), (Q4527 - 4578), (Q4547 - 4546), (Q2929 - 4443).

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Eimeria quokka was the most prevalent Eimeria species detected in quokka faeces from both Two Peoples Bay and Bald Island (26% and 74% of samples respectively) compared to E. setonicis (48% and 52% of samples respectively). Mixed infections of both Eimeria species were generally observed in the faeces of individual animals. However, single infections of both E. quokka and E. setonicis were detected in individual quokkas and determined by the uniformity of the oocyst. Quokka isolate

Q2929-4443 was identified as having a single infection of E. quokka, while quokka isolate Q4124-4119 was infected with only E. setonicis. DNA extracted from these two quokkas was amplified, sequenced and used for phylogenetic analysis. DNA was also extracted from quokka isolates BS3, WS2 and WE1 from Rottnest Island and characterised genetically.

8.3.3 Phylogenetic analysis of E. quokka, E. setonicis and Eimeria spp. from the quokka at the 18S rRNA gene

Phylogenetic analysis of a partial nuclear sequence (~481 bp) of the 18S rRNA gene from quokka isolate Q2929-4443 (positive with E. quokka from Two Peoples Bay) and quokka isolate Q4124-4119 (positive with E. setonicis from Bald Island) using distance, parsimony and ML analysis produced trees with similar topographies (Figure.

8.2, distance tree shown). The overall tree topology was similar to previously produced phylogenetic trees for Eimeria (Power et al., 2009; Yang et al., 2012). Both E. quokka and E. setonicis grouped within the marsupial clade together with E. macropodis, E. spp. from western grey kangaroos and E. trichosuri. A distance similarity matrix generated using Tajima - Nei distance (data not shown) at the 18S rRNA locus showed that E. quokka exhibited 97.3% similarity with E. setonicis, 97.3% similarity with E. trichosuri from possums, 96.7% similarity with E. macropodis from the tammar wallaby and 97.3% similarity with E. sp from western grey kangaroos (Yang et al.,

2012). Eimeria setonicis exhibited 99.6% similarity to E. trichosuri, 98.1% similarity to 148

E. macropodis (clone 10) and 99.2% similarity to an Eimeria sp from western grey kangaroos. Phylogenetic analysis was also conducted on the larger partial nuclear sequence (~1,288 bp) of the 18S rRNA gene for quokka isolate Q2929-4443 and

Q4124-4119 using distance, parsimony and ML analysis and produced the same topographies as the smaller partial 18S rRNA gene sequence (data not shown).

In addition to E. quokka and E. setonicis sequences, two novel sequences were identified. A novel sequence (referred to hereafter as novel Eimeria sequence 1) was identified from quokka isolates BS3 and WS2 from Rottnest. Direct sequencing of the

18S rRNA gene fragment from these two quokka isolates produced clean chromatograms, indicating that only one sequence was present. Sequences from isolates

BS3 and WS2 were 100% identical to each other and grouped most closely with the marsupial clade (Figure 8.3). Novel Eimeria sequence 1 exhibited 98.8% similarity to

E. trichosuri (clone A), 98.8% similarity to E. macropodis (clone 10), 98.3% similarity to an Eimeria sp. (2534) from western grey kangaroos, 96.7% similarity to E. quokka

(16 single nucleotide polymorphisms, SNPs over 481 bp of sequence) and 98.3% similarity to E. setonicis (8 SNPs over 481 bp of sequence). A second novel sequence

(novel Eimeria sequence 2) was identified from quokka isolate WE1 from Rottnest

Island, which grouped within the rodent clade and exhibited 99.8% and 99.2% similarity with E. telekii and E. separate, respectively. The partial 18S rRNA gene sequences generated in the present study have been submitted to GenBank under

Accession KF225636 and KF225639-KF225642.

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Figure 8.2. Evolutionary relationships of Eimeria quokka, Eimeria setonicis and Eimeria sp. inferred by distance analysis of partial (481bp) 18S rRNA sequences. Percentage support (>50%) from 1,000 pseudoreplicates from neighbor-joining analyses is indicated at the left of the support node. Shaded isolates are from this study.

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8.3.4. Phylogenetic analysis of E. quokka and E. setonicis from the quokka at the COI gene

Phylogenetic analysis of the mitochondrial COI partial sequence (~426 bp) from the E. quokka isolate (Q2929-4443) and the E. setonicis isolate (Q4124-4119) produced trees of similar topography to the 18S rRNA tree and grouped both E. quokka and E. setonicis within the marsupial clade (Figure. 8.3, distance tree shown). A distance similarity matrix generated using Tajima - Nei distance (data not shown) at this locus showed that E. quokka exhibited 96.1% similarity to E. setonicis, 93.8% similarity to E. trichosuri and 88.9% similarity to E. macropodis, while E. setonicis exhibited 96.9% similarity to E. trichosuri and 91.7% similarity to E. macropodis. Unfortunately, sequences from quokka isolates BS3, WS2 and WE1 from Rottnest could not be amplified at this locus. The partial COI sequences generated in the present study have been submitted to to GenBank under Accession KF225637 and KF225638.

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Figure. 8.3. Evolutionary relationships of Eimeria quokka and Eimeria setonicis from the quokka inferred by distance analysis of the mitochrondial cytochrome oxidase gene (CO1) (426 bp). Percentage support (>50%) from 1,000 pseudoreplicates from neighbor-joining analyses is indicated at the left of the support node.Shaded isolates are from this study.

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8.4 Discussion

In this chapter E. quokka, E. setonicis and two novel Eimeria species from the quokka were characterised for the first time using molecular tools. Eimeria species have previously been isolated and described morphologically from quokkas habituating

Rottnest Island only (Barker et al., 1988). Our results also extend the geographical range of Eimeria species from quokkas to Bald Island and the mainland of Western

Australia.

In the study by Barker et al. (1988), three different Eimeria species from quokkas were morphologically identified and described: E. quokka, E. setonicis and E. volckertzooni. In the present study, Eimeria oocysts resembling E. quokka and E. setonicis only were identified in quokka faeces collected from Two Peoples Bay, Bald

Island and Rottnest Island. Eimeria volckertzooni was described as having irregular blunt ellipsoid oocyst, ranging in length from 20 - 24µm and width from 11.6 - 13.6µm, often flattened on one side and containing a smooth clear colourless double cell wall, with the presence of a polar granule and oocyst residuum. Sporocysts were long and narrow and ranged in length from 11.2 - 15.2 µm and in width from 4.8 - 6.4 µm with an inconspicuous Steida body and two elongated sporozoites (Barker et al., 1988).

Oocysts similar to this were not detected in the present study.

Microscopic analysis showed E. quokka was the most prevalent Eimeria species from Two Peoples Bay and Bald Island and was previously reported as being the most prevalent Eimeria species in quokkas on Rottnest Island (Barker et al., 1988). Oocyst and sporozoite length and width dimensions for the Eimeria species identified in the present study were slightly larger than previously published measurements for E. quokka and E. setonicis. Eimeria quokka previously has been described as having ellipsoidal oocysts (13.6 - 21.6 µm in length and 8.8 -15.2 µm in width), with a double- layered wall, outer wall smooth, clear colourless, inner wall clear; polar granule present;

153 four ellipsoidal sporocysts (6.4 - 9.6 µm in length and 4.0 - 6.4 µm in width), with a small Stieda body, two sporozoites with granular cytoplasm, spheroidal sporozoites, refractile body and sporocyst residuum as granular clustals (Barker et al., 1988).

Eimeria setonicis has previously been described as having ellipsoidal oocysts (26.4 -

33.6 µm in length and 16.8 - 19.2 µm in width) which were slightly pointed at one end with a double-layered wall, outer wall smooth, clear colourless, inner wall clear; polar granule present; four ellipsoidal sporocysts (10.4 - 13.6 µm in length and 7.2 - 8.8 µm in width) with a Stieda body, two sporozites, spheroidal prominent sporozoites, refractile body and sporocyst residuum as granular clustals (Barker et al., 1988). The overall oocysts characteristics for E. quokka and E. setonicis from this study were consistent with descriptions of E. quokka and E. setonicis described by (Barker et al., 1988).

Morphological variation in sporulated oocysts within individual Eimeria species is well documented with polymorphism reported both within and between host species from different geographical locations (Duszynsk, 1971; Gardner and Duszynski, 1990). For example, variation in both oocyst and sporocyst morphometrics within E. cochabambensis from three different marsupial host species has been previously described (Heckscher et al., 1999). However, as no molecular analysis was conducted in those studies, the possibility that multiple species were present cannot be ruled out. A more recent study, however, which did conduct molecular analysis, reported variation in both oocyst and sporocyst morphometrics in E. trichosuri oocysts from brushtail possums (Power et al., 2009). The time of infection also has been reported to play a role in morphometric differences from single Eimeria species from the same host

(Duszynsk, 1971). This is because some members of the Eimeriidae are known to display polymorphism during patency and sporulated oocyst size can increase during this period (Duszynsk, 1971).

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Unfortunately, the quokka faecal samples from Rottnest Island did not sporulate when incubated in potassium dichromate. This may have been largely due to time of collection and storage conditions, as samples were collected in early January during peak summer conditions with temperatures reaching 30 - 40ºC. These high temperatures may have inactivated the oocysts, as ambient temperatures of 20 - 23ºC are more suited for sporulation (Duszynski and Wilber, 1997).

Molecular characterisation techniques such as sequencing of the 18S rRNA and

COI loci are currently being used to characterise and detect Eimeria species (Hill et al.,

2012; Yang et al., 2013). In the present study, molecular analysis was used to generate for the first time sequences for E. quokka and E. setonicis at two loci (18S rRNA and

COI) as well as two additional novel Eimeria sequences at the 18S rRNA locus.

Molecular detection was also more sensitive than microscopy and detected 85 vs. 45% prevalence for Eimeria in quokka faeces from Bald Island and 78.3 vs. 43.5% prevalence for Rottnest Island. Microscopy, however, is still vital for morphological characterisation of Eimeria oocysts from marsupial faecal samples, as marsupials are known to have multiple Eimeria species within the one host, and also as the availability of DNA sequences for marsupial Eimeria are limited (Power et al., 2009).

Phylogenetic analysis from both loci grouped E. quokka and E. setonicis within the marsupial clade together with E. trichosuri from brushtail possums and E. macropodis from tammar wallabies. Eimeria spp. from western grey kangaroos also grouped within the marsupial clade at the 18S loci. Unfortunately sequences from the

Eimeria spp. from the western grey kangaroos were not available at the COI locus.

Eimeria quokka and E. setonicis exhibited only 97.3% and 96.1% similarity to each other at the 18S rRNA and COI loci respectively. Eimeria quokka was most closely related to E. setonicis at the COI gene, while at the 18S rRNA gene it had 97.3% similarity with E. trichosuri from brushtail possums, Eimeria sp. from western grey

155 kangaroos and E. setonicis from quokkas. Eimeria setonicis was most closely related to

E. trichosuri at both loci with 99.6% and 96.9% similarity at the 18S rRNA and COI gene respectively. The differences seen in the levels of genetic diversity exhibited by the 18S rRNA and COI genes for Eimeria support comments by Hill et al. (2012) and

Yang et al. (2013), who highlight the need to use more than one genetic marker to fully understand the evolutionary relationship of marsupial Eimeria species.

Novel Eimeria sequence 1 (isolates WS2, BS3) from Rottnest Island was genetically distinct and formed its own clade within the marsupial group. At the 18S rRNA locus, it exhibited 98.8% similarity to E. macropodis and E. trichosuri. This novel isolate may represent a new Eimeria species, however, morphological and genetic analysis at the COI gene need to be conducted for this species before it can be properly validated. Novel Eimeria sequence 2 (isolate WE1) grouped within the rodent clade and grouped most closely to E. telekii from a striped grass mouse (Lemniscomys striatus) from Kenya (Slapeta et al., 2001). To the authors’ knowledge this is the first time that a rodent-like Eimeria species has been isolated from a marsupial host. Novel Eimeria sequence 2 was most likely acquired via mechanical transmission, as Eimeria sp. from mice are known to be host specific at the genus and species level (Hnida and Duszynski,

1999). However it has been suggested that transfers may occur among animals occurring together in the same habitat at the same site (syntopic hosts) under natural conditions (Hnida and Duszynski, 1999). This theory may account for our findings and also the recent detection of an Eimeria rodent isolate (genetically similar to E. falciformis and E. vermiformis) from a faecal sample from a King’s skink (Egernia kingii) from Western Australia (Yang et al., 2013).

In conclusion, the present study characterised E. quokka and E. setonicis for the first time at both the 18S rRNA and COI loci and demonstrated that they are closely related to Eimeria from other marsupials (E. trichosuri and E. macropodis) suggesting

156 co-evolution of marsupial Eimeria species regardless of the host. These findings also support the hypothesis that Eimeria found in marsupials diverged prior to Eimeria species from placental mammals (Power et al., 2009). Phylogenetic analysis has enabled the evolutionary placement of these two Eimeria species and has contributed to our knowledge on the genetic diversity of Eimeria species isolated from native marsupials.

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CHAPTER 9

GENERAL DISCUSSION

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9.1 Blood-borne and enteric parasites in marsupials and management implications

This thesis has encompassed characterisation of trypanosome and Eimeria species in the critically endangered Gilbert’s potoroo and the vulnerable quokka and has allowed for a significant increase in the current knowledge regarding the life-cycle and molecular epidemiology of these parasites. A novel trypanosome T. copemani, was identified and characterised in the Gilbert’s potoroo and the quokka, and a novel T. cruzi-like isolate and T. vegrandis were identified in individual quokka samples. In addition, two potentially novel Eimeria species were identified in quokkas and the first molecular characterisation of two Eimeria species, E. quokka and E. setonicis (which had previously been characterised morphologically only), was conducted.

Parasites can have pronounced or subtle effects on hosts, affecting host behaviour, growth, fecundity, and mortality. In the present study, the prevalence of T. copemani was 100% in the Gilbert’s potoroo and averaged 57.3% in quokka populations (range 4.9-91.4%). This is also the first report of T. copemani from Rottnest

Island. To date, little is known about the clinical effects of trypanosomes on Gilbert’s potoroos and quokkas, however, some trypanosomes, normally considered as non- pathogenic, are capable of inducing detrimental effects in the host, particularly when the parasite encounters new or naïve host species following their introduction into a new habitat (Maraghi and Molyneux, 1989), or when an infected host is exposed to additional or increased levels of stress (Brown et al., 2000; Wyatt et al., 2008). For example, some studies have shown that T. theileri, a non-pathogenic trypanosome of bovids that infects most cattle worldwide, may be considered potentially pathogenic in the presence of stress, gestation, poor nutritional status, and concurrent infections

(Hussain et al., 1985; Doherty et al., 1993; Seifi, 1995). Therefore, the potential for diseases (such as those caused by trypanosomes and coccidians) to reduce the fitness of

159 wildlife hosts and influence their population decline, is of particular concern, particularly given recent evidence linking T. copemani and T. vegrandis to the decline of the critically endangered woylie (Botero et al., 2013; Thompson et al., 2014b). In those studies, inflammation and tissue degeneration in woylies were reported, and principally occurred in the heart, skeletal muscle, oesophagus and tongue, with occasional detection of inflammatory cells around blood vessels (Botero et al., 2013).

The pathology seen in the heart of 3 woylies infected with T. copemani showed a multifocal, severe and chronic pyogranulomatous myocarditis and endocarditis accompanied by muscle degeneration and necrosis. Multifocal, chronic, pyogranulomatous oesophagitis and glossitis were also noted and accompanied by prominent skeletal muscle degeneration. The histopathological findings from the woylies in the report by Botero et al. (2013) were similar to those seen in Didelphis marsupialis infected with T. cruzi. In a study by Thompson et al. (2014b), a possible temporal connection implicating T. copemani as the disease agent linked with the recent decline of the Kingston indigenous woylie population within the Upper Warren region of Western Australia was proposed (Thompson et al., 2014b). Three different trypanosomes (T. vegrandis, T. copemani and Trypanosoma sp. H25) and two different strains of T. copemani that vary in virulence were detected by PCR screening. The spatial pattern of trypanosome infection varied among the five study sites, with a greater proportion of woylies from the Perup indigenous population being infected with T. copemani than from the neighbouring Kingston indigenous population. For an established infection, T. copemani detection was temporally inconsistent during the longevity of infection. The more virulent strain of T. copemani appeared to regress at a faster rate than the less virulent strain, with the infection possibly transitioning from the acute to chronic phase. In addition, it was also reported that interspecific competition may also exist between T. copemani and T. vegrandis, where an existing T. vegrandis

160 infection may moderate the sequential establishment of the more virulent T. copemani

(Thompson et al., 2014b). There is also evidence that trypanosomes contribute to host immunosuppression as significantly different patterns of parasitaemia and/or pathogenicity have been reported in mixed trypanosome infections (Dargantes et al.,

2005).

Fossil records show that the Gilbert’s potoroo lived within a narrow range along the south coast of Western Australia (Maxwell et al., 1996; Sinclair et al., 1996). It was thought to be extinct but was re-discovered in 1994, confined to Mount Gardner headland at Two People Bay, Nature Reserve, near Albany (Sinclair et al., 1996). When this project was initiated in 2006, the estimated total population size of the Gilbert’s potoroo was 40 animals. In 2015, the overall population is currently estimated at 100. A small number of animals were translocated to Bald Island in 2005, located off the coast of Albany, Western Australia, where the current estimated population size is 35 and therefore populations are still critically endangered. Quokka populations are confined to

Bald Island, Rottnest Island and the south-west corner of Western Australia. Although not the main focus of this study, the identification of erythrocyte abnormalities in quokka blood, including microspherocytes and schistocytes, changes typically associated with haemolytic anaemias (Mallah et al., 2010; Rodak et al., 2012), suggests that T. copemani may have a clinical effect on its host. This is supported by the studies in woylies discussed above that reported that the localisation of T. copemani in the capillaries and/or cells of the internal organs may be chronically pathogenic, adversely affecting the long term fitness and coordination, and making the host more vulnerable to predation (Thompson et al., 2014b).

Trypanosoma copemani has a broad host range, which includes the Gilbert’s potoroo, quokka, koala, common wombat, common brushtailed possum, tiger quoll and southern brown bandicoot (Bettiol et al., 1998; Noyes et al., 1999; McInnes et al., 2010;

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Botero et al., 2013; Thompson et al., 2014a; Thompson et al., 2014b). Its geographic range is also the largest of the native Australian trypanosomes and includes WA,

Queensland, New South Wales, Victoria and possibly Tasmania (McInnes et al., 2010;

Botero et al., 2013; Thompson et al., 2014a). As more mammals are sampled from inland habitats and from other states and territories (such as South Australia and

Northern Territory), both the host and geographic ranges of T. copemani and other trypanosomes are likely to increase. It is therefore clear that further research on the pathogenicity of T. copemani in marsupial hosts is urgently required.

In additional to trypanosomes, four Eimeria species were detected in quokkas at a prevalence of 61.4-85%. Eimeria species are the etiological agents for enteric coccidiosis, which is generally a disease associated with stressed or young animals and responsible for causing chronic diarrhea (Bennett et al., 2006). Multiple infections

(simultaneous infections with multiple parasite species in an individual host), also called polyparasitism, are the rule, rather than the exception, in the wild (Bordes and

Morand, 2011). Despite being recognised for many years, investigations on the impacts of multiple infections on individual hosts are still scarce in disease ecology (Bordes and

Morand, 2011). While individual pathogen infections may be tolerated and only present modest clinical signs of disease, interaction effects among multiple pathogens may dramatically alter their pathogenicity. For example, some studies have reported that co- infections with multiple parasite species resulted in more severe clinical disease

(Ezeamama et al., 2008). The clinical impact of Eimeria on the quokka is unknown, but it is possible that polyparasitism may decrease the fitness of quokkas and is an area that needs to be explored further.

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9.2 Morphology versus molecular characterisation

An important aspect of this study was the morphological and molecular characterisation of T. copemani. Morphological analysis demonstrated that T. copmani is a highly polymorphic parasite with three different trypomastigote forms identified.

Previous studies have also reported on morphological polymorphism in T. copemani with thin and broad trypomastigote forms described from woylies (Thompson et al.,

2013). In addition to trypomastigotes, life-cycle stages representing a promastigote, sphaeromastigote and amastigote stage were identified directly in blood films, for the first time, in samples from Australian marsupials. Four previously unidentified trypanosome forms (extremly thin form, adherent form, tiny form and circular form) were also identified both in vivo and in vitro and confirmed as trypanosomes by the use of trypanosome-specific antibody probes and FISH. The knowledge of life-cycle stages has very important implications for wildlife conservation, as it is possible that trypanosome infections are not being detected by microscopy as stages other than trypomastigotes are present in blood, and what was once believed to platelets may actually be trypanosome life-cycle stages.

Molecular characterisation at both the 18S rRNA and GAPDH loci identified genetic variation within T. copemani with two different genotypes (A and B) identified within potoroo and quokka populations with genotype B being the most prevalent.

Trypanosoma copemani genotype B was only found infecting marsupials in Western

Australian being restricted to quokkas and Gilbert’s potoroos, while genotype A was found to be more widely distributed and affects a wide variety of marsupials from both

Western and Eastern Australia (Noyes et al., 1999; McInnes et al., 2011; Botero et al.,

2013; Thompson et al., 2013). The identification of mixed trypanosome infections in quokkas also became apparent with co-infections of both T. copemani genotypes and one quokka infected with both T. copemani and a novel T. cruzi-like isolate and another

163 quokka infected with both T. copemani and T. vegrandis. Interestingly, it has been suggested that T. copemani genotype A (genotype G2 - Botero et al., 2013) is more virulent than other T. copemani genotypes, as it has been implicated in pathological changes to heart, oesophagus, tongue and skeletal muscle with reported clinical characteristics being similar to those of Chagas’ disease in humans (Botero et al., 2013;

Thompson et al., 2014b). This has important implications for immunologically naïve and endangered species, with the potential of disease reducing the fitness of the host and making it more susceptible to predation and population decline.

Although not a key aim of this project, phylogenetic analysis identified large genetic distances within T. vegrandis isolates at both the 18S rRNA locus (1.5-9.8%) and the GAPDH locus (16.9-18.5%). This indicates that T. vegrandis is not a uniform species but is in fact a species complex. As T. vegrandis is the smallest trypanosome identified to date (8.3 µm in length and 1.3 µm in width), morphological characterisation using scanning or transmission electron microscopy will be required to better characterise species within this complex.

Morphological and molecular analysis was also conducted on Eimeria with PCR analysis detecting a prevalence of 62.5%, 85.0% and 78.3% for Eimeria in quokkas from Two Peoples Bay, Bald Island and Rottnest Island, respectively, compared to

62.5%, 45% and 43.5% for microscopy. Phylogenetic analysis at both the 18S rRNA and CO1 loci showed that E. quokka, E. setonicis and one novel Eimeria sequence grouped within the Eimeria marsupial clade but the fourth Eimeria sequence clustered within a rodent clade. Morphological characterisation is still required to fully characterise the two novel Rottnest Eimeria isolates (Eimeria sequence 1 and Eimeria sequence 2).

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9.3 Transmission of T. copemani

The transmission of native Australian trypanosomes is another issue that requires urgent attention, as very little is known about their life-cycle, vectors

(including cyclical and mechanical vectors) and transmission dynamics (stercorarian vs. salivarian). Identification of the vector(s) that transmit trypanosomes in various marsupial populations will have important implications for our understanding of both the epidemiology of these diseases and for conservation strategies (Hoare, 1972). For example, in humans and other mammals, the prevalence of trypanosomes is directly proportional to the density of the vector population (Sehgal et al., 2001). Thus knowledge of the vectors and transmission routes will aid in management decisions such as avoiding re-introduction of naïve animals into areas where both the vectors and pathogens are known to be present in the same or other species of marsupials.

In this study, phylogenetic analysis of T. copemani indicated that it fell within the stercorarian group of trypanosomes, which use faecal contamination of skin as a route of transmission. Ectoparasites (ticks and fleas) found dwelling and defaecating on marsupial hosts were collected and motile trypanosomes were observed in the haemolymph, midgut and faecal matter isolated from ticks and confirmed as T. copemani using molecular analysis. Transmission studies are however required to fully validate the tick as the vector of T. copemani but given the general vulnerability and endangered status of many Australian marsupials, ethical constraints may hinder such future studies. Confirmed identification of the vector for T. copemani as I. australiensis will allow for more targeted management and conservation strategies of marsupial hosts in the future.

9.4 The zoonotic potential of Australian trypanosomes

The zoonotic potential of T. copemani was investigated using a blood incubation infectivity test, to determine its potential for growth in human serum. To eliminate any 165 effects of anticoagulants on the complement system and on human high density lipoprotein (HDL), only fresh whole human blood was used. Trypanosoma copemani was observed by microscopy in all human blood cultures from day 5 to day 19 post inoculation (PI) and confirmed using molecular analysis. The mechanism for normal human serum (NHS) resistance in T. copemani is not known. The results of this study show that at least one native Australian trypanosome species may have the potential to be infective for humans. Approximately 75% of new emerging infectious diseases

(EIDs) that have affected human populations in the past 30 years have been zoonotic

(Daszak et al., 2007). However, the observation that T. copemani is able to resist NHS is insufficient to conclude that it has significant zoonotic potential per se. Further research is required to challenge the vectors metacyclic life-cycle stages to the BIIT and to determine the prevalence of trypanosomes in ticks removed from humans (and other ectoparasites capable of transmitting trypanosomes such as tabanids and ) as well as in human blood.

9.5 Future studies

The complexity of the morphology of the different life-cycle stages associated with Australian trypanosomes has been highlighted in this study and further research into this area is warranted. Species-specific molecular probes for FISH analysis would be beneficial to determine the different morphological phenotypes associated with each life-cycle stage of T. copemani and help to reduce false descriptions of other potential trypanosome species, given the detection of co-infection with different trypanosome species within some quokka isolates. Such tools would further aid in characterising novel life-cycle forms, as observed in this study and help to elucidate the complete life- cycle of T. copemani. In addition, a standard marker such as mitotracker probe, which is a cell-permeant mitochondrion-selective dye that accumulates in active mitochondria

(Vassella et al., 1997), would help to clarify the viability of novel trypanosome forms.

166

The use of transmission electron microscopy is also necessary to confirm the presence of both nuclear and kinetoplast regions.

In the present study, Sanger sequencing identified T. copemani in the majority of isolates screened. However, it is possible that Sanger sequencing is not detecting the true prevalence of mixed infections as less prevalent species may not be amplified or may produce mixed chromatograms. Next generation deep sequencing platforms such as MiSeq (Illumina) should therefore be applied for identification of levels of mixed infections.

Whole genome sequencing (WGS) should also be conducted on T. copemani and other marsupial-derived marsupial species to (1) determine the phylogenetic relationships between trypanosome parasites from marsupials, and (2) to look for genetic evidence of how long these trypanosomes have been associated with marsupial hosts. Knowing whether trypanosomes are recent or long-term pathogens in different marsupial host species is important because emerging diseases are different from established pathogens due to the fact that pathogen interactions with novel hosts are unpredictable and are more likely to lead to host population declines and extinctions

(Tobler et al., 2012). Almost all disease-threatening extinctions result from a host encountering a novel pathogen that it has had no exposure to in evolutionary time

(McCallum, 2012). This process was clearly demonstrated with the extinction of a native rodent species (Rattus macleari) on Christmas Island when exposed to

Trypanosoma lewisi believed to have been introduced to the island by fleas hosted on stowaway black rats (Rattus rattus) (Wyatt et al., 2008).

With regards to transmission, future studies are required to determine if the tick is capable of transmitting T. copemani to other host species through either the contaminative or inoculation route or both. Information gained from such a study would be imperative for reducing the distribution of this parasite through vector control

167 management and would be vital for future translocation programs, particularly associated with immunologically naive and endangered host species.

Due to the absence of the typical trypomastigotes stage in blood films and the low prevalence of trypanosomes in the Rottnest Island quokkas, additional prevalence studies are needed to ascertain the epidemiology of T. copemani on Rottnest Island, given the close association between tourist and quokka.

9.6 Conclusions

Overall this study has demonstrated the complexity and polymorphic nature of

Australian native trypanosomes and has allowed the characterisation of T. copemani.

Multiple genotypes have been identified within this species, with T. copemani genotype

A from the quokka and Gilbert’s potoroo having 100% homology at the 18S rRNA locus to a novel trypanosome isolate (G2 - Botero et al., 2013), known to cause clinical pathologies similar to T. cruzi. Morphological analysis in combination with molecular probes has enabled the description of novel trypanosome forms and has shown that the parasite has the potential to cause erythrocyte destruction and retrospectively induce haemolytic anaemia, which is a common feature of animal trypanosomiasis, with the exact cause to date unknown.

The prevalence of T. copemani is wide-spread within marsupial populations around Australia, with the tick identified as the putative vector. Given the often close association of humans and wildlife such as humans and quokkas on Rottnest Island, the need to understand how T. copemani is transmitted is imperative and demonstrates the need for future transmission studies on T. copemani. It is clear from this study that many questions have been answered and have contributed to our understanding of

Australian parasites. However, it is also apparent that our newly gained knowledge leads to many more questions that need to be addressed, with additional research crucial

168 to better understanding the epidemiology and transmission dynamics of parasites in marsupial hosts.

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REFERENCES

Aarthi, S., Raj, G.D., Raman, M., Gomathinayagam, S., Kumanan, K., (2010). Molecular prevalence and preponderance of Eimeria spp. among chickens in Tamil Nadu, India. Parasitology Research 107, 1013-1017. Adade, C.M., Carvalho, A.L., Tomaz, M.A., Costa, T.F., Godinho, J.L., Melo, P.A., Lima, A.P., Rodrigues, J.C., Zingali, R.B., Souto-Padron, T., (2014). Crovirin, a snake venom cysteine-rich secretory protein (CRISP) with promising activity against trypanosomes and Leishmania. PLoS Neglected Tropical Diseases 8, e3252. Alkhaldi, A.A., Creek, D.J., Ibrahim, H., Kim, D.H., Quashie, N.B., Burgess, K.E., Changtam, C., Barrett, M.P., Suksamrarn, A., de Koning, H.P., (2015). Potent trypanocidal curcumin analogs bearing a monoenone linker motif act on Trypanosoma brucei by forming an adduct with trypanothione. Molecular Pharmacology 87, 451-464. Alvarez, F., Cortinas, M.N., Musto, H., (1996). The analysis of protein coding genes suggests monophyly of Trypanosoma. Molecular Phylogenetics and Evolution 5, 333-343. Amole, B.O., Clarkson, A.B., Shear, H.L., (1982). Pathogenesis of anemia in Trypanosoma brucei infected mice. Infection and Immunity 36, 1060-1068. Anosa, V.O., Kaneko, J.J., (1983). Pathogenesis of Trypanosoma brucei infection in deer mice (Peromyscus-Maniculatus) .V. Macrophage ultrastructure and function. Veterinary Pathology 20, 617-631. Aufderheide, A.C., Salo, W., Madden, M., Streitz, J., Buikstra, J., Guhl, F., Arriaza, B., Renier, C., Wittmers, L.E., JR.,, Fornaciari, G., Allison, M., (2004). A 9,000- year record of Chagas' disease. Proceedings of the National Academy of Sciences of the United States of America 101, 2034-2039. Austen, J.M., Ryan, U.M., Friend, J.A., Ditcham, W.G., Reid, S.A., (2011). Vector of Trypanosoma copemani identified as Ixodes sp. Parasitology 138, 866-872. Averis, S., Thompson, R.C., Lymbery, A.J., Wayne, A.F., Morris, K.D., Smith, A., (2009). The diversity, distribution and host-parasite associations of trypanosomes in Western Australian wildlife. Parasitology 136, 1269-1279. Bancroft, T.L., (1888). Note on haematomonas in rat's blood. Proceedings of the Royal Society Queensland 5, 31-32. Barbosa, A., Austen, J., Gillett, A., Warren, K., Paparini, A., Irwin, P., Ryan, U., (2016). First report of Trypanosoma vegrandis in koalas (Phascolarctos cinereus). Parasitology International 65, 316-318. Barker, I.K., Ocallaghan, M.G., Beveridge, I., (1988). Eimeria spp. (Apicomplexa:Eimeriidae) parasitic in wallabies and kangaroos of the genera Setonix, Thylogale, Wallabia, Lagorchestes and Dendrolagus (Marsupialia:Macropodidae). International Journal for Parasitology 18, 955- 962. Barker, I.K., Ocallaghan, M.G., Beveridge, I., (1989). Host-parasite associations of Eimeria spp (Apicomplexa, Eimeriidae) in kangaroos and wallabies of the genus Macropus (Marsupialia, Macropodidae). International Journal for Parasitology 19, 241-263. Barker, S., Glover, R., Jacobsen, P., Kakulas, B.A., (1974). Seasonal anemia in Rottnest quokka, Setonix brachyurus (Quoy + Gaimard) (Marsupialia-Macropodidae). Comparative Biochemistry and Physiology 49, 147-157.

170

Barrett, M.P., Boykin, D.W., Brun, R., Tidwell, R.R., (2007). Human African trypanosomiasis: pharmacological re-engagement with a neglected disease. British Journal of Pharmacology 152, 1155-1171. Barrett, M.P., Coombs, G.H., Mottram, J.C., 2004. Future prospects in chemotherapy for trypanosomiasis, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The trypanosomiases. CABI Publishing, Wallingford, pp. 445-460. Barry, D., Carrington, M., 2004. Antigenic variation, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The trypanosomiases. CABI Publishing, Wallingford, pp. 25- 37. Barta, J.R., Martin, D.S., Liberator, P.A., Dashkevicz, M., Anderson, J.W., Feighner, S.D., Elbrecht, A., PerkinsBarrow, A., Jenkins, M.C., Danforth, H.D., Ruff, M.D., ProfousJuchelka, H., (1997). Phylogenetic relationships among eight Eimeria species infecting domestic fowl inferred using complete small subunit ribosomal DNA sequences. Journal of Parasitology 83, 262-271. Bastin, P., Ellis, K., Kohl, L., Gull, K., (2000). Flagellum ontogeny in trypanosomes studied via an inherited and regulated RNA interference system. Journal of Cell Science 113 ( Pt 18), 3321-3328. Bastin, P., Sherwin, T., Gull, K., (1998). Paraflagellar rod is vital for trypanosome motility. Nature 391, 548. Bennett, M.D., Woolford, L., O'Hara, A.J., Nicholls, P.K., Warren, K., Hobbs, R.P., (2006). A new Eimeria species parasitic in western barred bandicoots, perameles bougainville (Marsupialia : Peramelidae), in Western Australia. Journal of Parasitology 92, 1292-1294. Bettiol, S.S., Jakes, K., Le, D.D., Goldsmid, J.M., Hocking, G., (1998). First record of trypanosomes in Tasmanian bandicoots. Journal of Parasitology 84, 538-541. Bonney, K.M., Engman, D.M., (2008). Chagas’ heart disease pathogenesis: one mechanism or many? Current Molecular Medicine 8, 510-518. Bordes, F., Morand, S., (2011). The impact of multiple infections on wild animal hosts: a review. Infection Ecology and Epidemiology 1. Botero, A., Thompson, C.K., Peacock, C.S., Clode, P.L., Nicholls, P.K., Wayne, A.F., Lymbery, A.J., Thompson, R.C., (2013). Trypanosomes genetic diversity, polyparasitism and the population decline of the critically endangered Australian marsupial, the brush tailed bettong or woylie (Bettongia penicillata). International Journal for Parasitology. Parasites and Wildlife 2, 77-89. Brack, C., (1968). Elektronenmikroskopische untersuchungen zum lebenszyklus von Trypanosoma cruzi. Unter besonderer berucksichtigung der entwicklungsformen im uebertrager Rhodnius prolixus. Acta Tropica 25, 289. Breinl, A., (1913). Parasite protozoa encountered in the blood of Australian native animals. The Australian Institute of Tropical Medicine, 30-38. Brener, Z., (1973). Biology of Trypanosoma cruzi. Annual Review of Microbiology 27, 347-382. Brown, M.J.F., Loosli, R., Schmid-Hempel, P., (2000). Condition-dependent expression of virulence in a trypanosome infecting bumblebees. Oikos 91, 421-427. Bruce, D. 1895. Preliminary report on the tsetse fly disease or nagana in Zululand (Durban : Bennett and David). Brun, R., Hecker, H., Lun, Z.R., (1998). Trypanosoma evansi and T. equiperdum: distribution, biology, treatment and phylogenetic relationship (a review). Veterinary Parasitology 79, 95-107. Capewell, P., Clucas, C., DeJesus, E., Kieft, R., Hajduk, S., Veitch, N., Steketee, P.C., Cooper, A., Weir, W., MacLeod, A., (2013). The TgsGP gene is essential for resistance to human sserum in Trypanosoma brucei gambiense. PLoS Pathogens 9. 171

Cestari, I., Ramirez, M.I., (2010). Inefficient complement system clearance of Trypanosoma cruzi metacyclic trypomastigotes enables resistant strains to invade eukaryotic cells. PLoS One 5, e9721. Cestari, I.D., Evans-Osses, I., Freitas, J.C., Inal, J.M., Ramirez, M.I., (2008). Complement C2 receptor inhibitor trispanning confers an increased ability to resist complement-mediated lysis in Trypanosoma cruzi. Journal of Infectious Diseases 198, 1276-1283. Chagas, C., (1909). Neue trypanosomen Archiv für Schiffs- und Tropen-Hygiene 13, 120. Chekwube, A.I., Onyema, E.I., Ikenna, U.E., Ezeokonkwo, R.C., (2014). Effect of diminazene aceturate, levamisole and vitamin C combination therapy in rats experimentally infected with Trypanosoma brucei brucei. Asian Pacific Journal of Tropical Medicine 7, 438-445. Clark, P., Adlard, R.D., Spratt, D.M., 2004. Haemoparasites of Australian mammals. In: Haematology of Australian mammals. CSIRO, Collingwood, Australia. Clark, P., Spencer, P., (2006). Haematological characteristics of wild quokka (Setonix brachyurus). Comparative Clinical Pathology 15, 82-86. Cleland, J.B., (1906). The haemogregarine of mammals (H. balfouri), and some notes on rats. Journal of Tropical Medicine, 296-297. Cleland, J.B., (1908). Some remarks on the natural history and disease of the rats of Perth and Fremantle, Western Australia. Report of the Australian Association for the Advancement of Science for 1907, 516-520. Constantine, C.C., (2003). Importance and pitfalls of molecular analysis to parasite epidemiology. Trends in Parasitology 19, 346-348. Cox, F.E.G., (1977). Interactions between trypanosomes and piroplasms in mice. Protozoology 111, 129-134. Cox, F.E.G., (1998). Control of coccidiosis: lessons from other sporozoa. International Journal for Parasitology 28, 165-179. Cunningham, I., (1977). New culture medium for maintenance of tsetse tissues and growth of trypanosomatids. The Journal of Protozoology 24, 325-329. Dargantes, A.P., Campbell, R.S., Copeman, D.B., Reid, S.A., (2005). Experimental Trypanosoma evansi infection in the goat. II. Pathology. Journal of Comparative Pathology 133, 267-276. Daszak, P., Epstein, J.H., Kilpatrick, A.M., Aguirre, A.A., Karesh, W.B., Cunningham, A.A., (2007). Collaborative research approaches to the role of wildlife in zoonotic disease emergence. Current Topics in Microbiology and Immunology 315, 463-475. Daugschies, A., Imarom, S., Bollwahn, W., (1999). Differentiation of porcine Eimeria spp. by morphologic algorithms. Veterinary Parasitology 81, 201-210. De Greef, C., Hamers, R., (1994). The serum resistance-associated (SRA) gene of Trypanosoma brucei rhodesiense encodes a variant surface glycoprotein-like protein. Molecular and Biochemical Parasitology 68, 277-284. De Souza, W., (2002). Basic cell biology of Trypanosoma cruzi. Current Pharmaceutical Design 8, 269-285. Desquesnes, M., Ravel, S., Cuny, G., (2002). PCR identification of Trypanosoma lewisi, a common parasite of laboratory rats. Kinetoplastid Biology and Disease 1, 2. Doherty, M.L., Windle, H., Voorheis, H.P., Larkin, H., Casey, M., Clery, D., Murray, M., (1993). Clinical disease associated with Trypanosoma theileri infection in a calf in Ireland. Veterinary Record 132, 653-656. Durham, H.E., (1908). Notes on nagana and on some haematozoa observed during my travels. Parasitology 1, 227-235. 172

Duszynsk, D.W., (1971). Increase in size of Eimeria separata oocysts during patency. Journal of Parasitology 57, 948-&. Duszynski, D.W., Wilber, P.G., (1997). A guideline for the preparation of species descriptions in the Eimeriidae. Journal of Parasitology 83, 333-336. Dutton, J.E. 1902. Preliminary note upon a trypanosome occuring in the blood of man (Thomson Yates Laboratory Report), p. 445. Duvallet, G., de la Rocque, S., Reifenberg, J.M., Solano, P., Lefrancois, T., Michel, J.F., Bengaly, Z., Sidibe, I., Cuisance, D., Cuny, G., (1999). Review on the molecular tools for the understanding of the epidemiology of animal trypanosomosis in West Africa. Memorias Do Instituto Oswaldo Cruz 94, 245-248. Eberhard, M.L., da Silva, A.J., Lilley, B.G., Pieniazek, N.J., (1999). Morphologic and molecular characterization of new Cyclospora species from Ethiopian monkeys: C. ercopitheci sp.n., C. colobi sp.n., and C. papionis sp.n. Emerging Infectious Diseases 5, 651-658. Edwards, K.T., Goddard, J., Varela-Stokes, A.S., (2009). Examination of the internal morphology of the ixodid tick, Amblyomma maculatum Koch, (Acari: Ixodidae); a “how-to” pictorial dissection guide Midsouth Entomologist 2, 28-39. El Kady, G.A., (1998). Protozoal parasites in tick species infesting camels in Sinai Peninsula. Journal of the Egyptian Society of Parasitology 28, 765-776. Ezeamama, A.E., McGarvey, S.T., Acosta, L.P., Zierler, S., Manalo, D.L., Wu, H.W., Kurtis, J.D., Mor, V., Olveda, R.M., Friedman, J.F., (2008). The synergistic effect of concomitant schistosomiasis, hookworm, and trichuris infections on children's anemia burden. PLoS Neglected Tropical Diseases 2. Fernandes, A.P., Nelson, K., Beverley, S.M., (1993). Evolution of nuclear ribosomal RNAs in kinetoplastid protozoa: perspectives on the age and origins of . Proceedings of the National Academy of Sciences of the United States of America 90, 11608-11612. Fernandes, M.C., Andrews, N.W., (2012). Host cell invasion by Trypanosoma cruzi: a unique strategy that promotes persistence. FEMS Microbiology Reviews 36, 734- 747. Ferrante, A., Allison, A.C., (1983). Alternative pathway activation of complement by African trypanosomes lacking a glycoprotein coat. Parasite Immunology 5, 491- 498. Gardner, S.L., Duszynski, D.W., (1990). Polymorphism of eimerian oocysts can be a problem in naturally infected hosts - an example from subterranean rodents in bolivia. Journal of Parasitology 76, 805-811. Geigy, R., Jenni, L., Kauffmann, M., Onyango, R.J., Weiss, N., (1975). Identification of T. brucei-subgroup strains isolated from game. Acta Tropica 32, 190-205. Geigy, R., Kauffmann, M., Steiger, R., Brun, R., (1971). Influence of bloodmeals from different donors on the infection rates of Trypanosoma brucei in Glossina. Acta Tropica 28, 164-169. Gibson, W., (2009). Species-specific probes for the identification of the African tsetse- transmitted trypanosomes. Parasitology 136, 1501-1507. Glick, B.R., Pasternak, J.J., 2003. Molecular biotechnology: principles and applications of recombinant DNA, third Edition. ASM Press, Washington, D.C. Gomez, E., Valdes, A.M., Pinero, D., Hernandez, R., (1991). What is a genus in the Trypanosomatidae family - phylogenetic analysis of 2 small ribosomal-rna sequences. Molecular Biology and Evolution 8, 254-259. Goodman-Gilman, A., Hardman, J.G., Limbird, L.E., Molinoff, P.B., Ruddon, R.W., 1996. Goodman and Gilman's the pharmacological basis of therapeutics. McGraw-Hill, Sydney.

173

Guhl, F., Vallejo, G.A., (2003). Trypanosoma (Herpetosoma) rangeli Tejera, 1920 - An updated review. Memorias Do Instituto Oswaldo Cruz 98, 435-442. Guindon, S., Gascuel, O., (2003). A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Systematic Biology 52, 696-704. Haag, J., O'HUigin, C., Overath, P., (1998). The molecular phylogeny of trypanosomes: evidence for an early divergence of the Salivaria. Molecular and Biochemical Parasitology 91, 37-49. Hall, T.A., (1999). BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series 41, 95-98. Hamilton, P.B., Gibson, W.C., Stevens, J.R., (2007). Patterns of co-evolution between trypanosomes and their hosts deduced from ribosomal RNA and protein-coding gene phylogenies. Molecular Phylogenetics and Evolution 44, 15-25. Hamilton, P.B., Stevens, J.R., Gaunt, M.W., Gidley, J., Gibson, W.C., (2004). Trypanosomes are monophyletic: evidence from genes for glyceraldehyde phosphate dehydrogenase and small subunit ribosomal RNA. International Journal for Parasitology 34, 1393-1404. Hamilton, P.B., Stevens, J.R., Gidley, J., Holz, P., Gibson, W.C., (2005a). A new lineage of trypanosomes from Australian vertebrates and terrestrial bloodsucking leeches (Haemadipsidae). International Journal for Parasitology 35, 431-443. Hamilton, P.B., Stevens, J.R., Holz, P., Boag, B., Cooke, B., Gibson, W.C., (2005b). The inadvertent introduction into Australia of Trypanosoma nabiasi, the trypanosome of the European rabbit (Oryctolagus cuniculus), and its potential for biocontrol. Molecular Ecology 14, 3167-3175. Hamilton, P.B., Teixeira, M.M., Stevens, J.R., (2012). The evolution of Trypanosoma cruzi : the 'bat seeding' hypothesis. Trends in Parasitology 28, 136-141. Hannaert, V., Opperdoes, F.R., Michels, P.A.M., (1998). Comparison and evolutionary analysis of the glycosomal glyceraldehyde-3-phosphate dehydrogenase from different kinetoplastida. Journal of Molecular Evolution 47, 728-738. Hart, R.P., Bradshaw, S.D., Iveson, J.B., (1985). Salmonella infections in a marsupial, the quokka (Setonix brachyurus), in relation to seasonal changes in condition and environmental stress. Applied and Environmental Mcrobiology 49, 1276- 1281. Hawking, F., (1976). The resistance to human plasma of Trypanosoma brucei, T. rhodesiense and T. gambiense. II. Survey of strains from East Africa and Nigeria. Transactions of the Royal Society of Tropical Medicine and Hygiene 70, 513-520. Hawking, F., (1978). The resistance of Trypanosoma congolense, T. vivax and T. evansi to human plasma. Transactions of the Royal Society of Tropical Medicine and Hygiene 72, 405-407. Hebert, P.D.N., Ratnasingham, S., deWaard, J.R., (2003). Barcoding animal life: cytochrome c oxidase subunit 1 divergences among closely related species. Proceedings of the Royal Society B-Biological Sciences 270, S96-S99. Heckscher, S.K., Wickesberg, B.A., Duszynski, D.W., Gardner, S.L., (1999). Three new species of Eimeria from Bolivian marsupials. International Journal for Parasitology 29, 275-284. Herrera, C.P., Licon, M.H., Nation, C.S., Jameson, S.B., Wesson, D.M., (2015). Genotype diversity of Trypanosoma cruzi in small rodents and Triatoma sanguisuga from a rural area in New Orleans, Louisiana. Parasites & Vectors 8, 1-9.

174

Hide, G., Tait, A., 2004. Genetics and molecular epidemiology of trypanosomes, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The Trypanosomiases. CABI Publishing, Wallingford, pp. 77-93. Hill, N.J., Richter, C., Power, M.L., (2012). Pinning down a polymorphic parasite: New genetic and morphological descriptions of Eimeria macropodis from the tammar wallaby (Macropus eugenii). Parasitology International 61, 461-465. Hnida, J.A., Duszynski, D.W., (1999). Cross-transmission studies with Eimeria arizonensis, E. arizonensis-like oocysts and Eimeria langebarteli: Host specificity at the genus and species level within the muridae. Journal of Parasitology 85, 873-877. Hoare, C., 1972. The trypanosomes of mammals. A zoological monograph, In: Blackwell Scientific Publishing, Oxford, England. Hussain, K., Brodie, B., Ott, R.S., Montealegre, F., (1985). Prevalence of Trypanosoma theileri in cows and fetuses at slaughter. American Journal of Veterinary Research 46, 1256-1258. Iida, K., Whitlow, M.B., Nussenzweig, V., (1989). Amastigotes of Trypanosoma cruzi escape destruction by the terminal complement components. Journal of Experimental Medicine 169, 881-891. Jacobs, R.T., Plattner, J.J., Nare, B., Wring, S.A., Chen, D., Freund, Y., Gaukel, E.G., Orr, M.D., Perales, J.B., Jenks, M., Noe, R.A., Sligar, J.M., Zhang, Y.K., Bacchi, C.J., Yarlett, N., Don, R., (2011). Benzoxaboroles: a new class of potential drugs for human African trypanosomiasis. Future Medicinal Chemistry 3, 1259-1278. Jakes, K.A., O'Donoghue, P., Munro, M., Adlard, R., (2001a). Hemoprotozoa of freshwater turtles in Queensland. J Wildl Dis 37, 12-19. Jakes, K.A., O'Donoghue, P.J., Adlard, R.D., (2001b). Phylogenetic relationships of Trypanosoma chelodina and Trypanosoma binneyi from Australian tortoises and platypuses inferred from small subunit rRNA analyses. Parasitology 123, 483- 487. Jefferies, R., Ryan, U.M., Irwin, P.J., (2007). PCR-RFLP for the detection and differentiation of the canine piroplasm species and its use with filter paper-based technologies. Veterinary Parasitology 144, 20-27. Joiner, K.A., Dasilva, W.D., Rimoldi, M.T., Hammer, C.H., Sher, A., Kipnis, T.L., (1988). Biochemical characterization of a factor produced by trypomastigotes of Trypanosoma cruzi that accelerates the decay of complement c-3 convertases. Journal of Biological Chemistry 263, 11327-11335. Kobayashi, A., Tizard, I.R., Woo, P.T., (1976). Studies on the anemia in experimental African trypanosomiasis. II. The pathogenesis of the anemia in calves infected with Trypanosoma congolense. The American Journal of Tropical Medicine and Hygiene 25, 401-406. Koffi, M., De Meeus, T., Sere, M., Bucheton, B., Simo, G., Njiokou, F., Salim, B., Kabore, J., MacLeod, A., Camara, M., Solano, P., Belem, A.M.G., Jamonneau, V., (2015). Population genetics and reproductive strategies of African trypanosomes: revisiting available published data. PLoS Neglected Tropical Diseases, 1-21. Kohl, L., Gull, K., (1998). Molecular architecture of the trypanosome cytoskeleton. Molecular and Biochemical Parasitology 93, 1-9. Kollien, A.H., Schaub, G.A., (1998). The development of Trypanosoma cruzi (trypanosomatidae) in the reduviid bug Triatoma infestans (insecta): Influence of starvation. Journal of Eukaryotic Microbiology 45, 59-63. Krautz, G.M., Kissinger, J.C., Krettli, A.U., (2000). The targets of the lytic antibody response against Trypanosoma cruzi. Parasitology Today 16, 31-34. 175

La Greca, F., Magez, S., (2011). Vaccination against trypanosomiasis can it be done or is the trypanosome truly the ultimate immune destroyer and escape artist? Human Vaccines 7, 1225-1233. Lai, D.H., Wang, Q.P., Li, Z., Lukes, J., Lun, Z.R., (2009). Evolution of the serum resistance-associated SRA gene in African trypanosomes. Chinese Science Bulletin 54, 1275-1278. Langousis, G., Hill, K.L., (2014). Motility and more: the flagellum of Trypanosoma brucei. Nature reviews. Microbiology 12, 505-518. Larkin, M.A., Blackshields, G., Brown, N.P., Chenna R., McGettigan, P., McWilliam, H., Valentin, F., Wallace, I.M., Wilm, A., Lopez, R., Thompson, J.D., Gibson, T.J., Higgins, D.G., (2007). ClustalW and ClustalX version 2 (2007). Bioinformatics 23, 2947-2948. Latif, A.A., Bakheit, M.A., Mohamed, A.E., Zweygarth, E., (2004). High infection rates of the tick Hyalomma anatolicum anatolicum with Trypanosoma theileri. Onderstepoort Journal of Veterinary Research 71, 251-256. Lee, J.Y., Ryan, U.M., Jefferies, R., McInnes, L.M., Forshaw, D., Friend, J.A., Irwin, P.J., (2009). Theileria gilberti n. sp. (Apicomplexa: Theileriidae) in the Gilbert's potoroo (Potorous gilbertii). The Journal of Eukaryotic Microbiology 56, 290- 295. Levine, N.D., Corliss, J.O., Cox, F.E., Deroux, G., Grain, J., Honigberg, B.M., Leedale, G.F., Loeblich, A.R., 3rd, Lom, J., Lynn, D., Merinfeld, E.G., Page, F.C., Poljansky, G., Sprague, V., Vavra, J., Wallace, F.G., (1980). A newly revised classification of the protozoa. The Journal of Protozoology 27, 37-58. Ley, V., Andrews, N.W., Robbins, E.S., Nussenzweig, V., (1988). Amastigotes of Trypanosoma cruzi sustain an infective cycle in mammalian cells. The Journal of experimental medicine 168, 649-659. Lukes, J., Jirku, M., Dolezel, D., Kralova, I., Hollar, L., Maslov, D.A., (1997). Analysis of ribosomal RNA genes suggests that trypanosomes are monophyletic. Journal of Molecular Evolution 44, 521-527. Lun, Z.R., Reid, S.A., Lai, D.H., Li, F.J., (2009). Atypical human trypanosomiasis: a neglected disease or just an unlucky accident? Trends in Parasitology 25, 107- 108. Machado, E.M.M., Fernandes, A.J., Murta, S.M.F., Vitor, R.W.A., Camilo, D.J., Pinheiro, S.W., Lopes, E.R., Adad, S.J., Romanha, A.J., Dias, J.C.P., (2001). A study of experimental reinfection by Trypanosoma cruzi in dogs. American Journal of Tropical Medicine and Hygiene 65, 958-965. Mackerras, I.M., Mackerras, M.J., (1960). Taxonomy of the common short-nosed marsupial bandicoot of eastern Queensland. Australian Journal of Science 23, 51-53. Mackerras, I.M., Mackerras, M.J., Sandars, D.F., (1953). Parasites of the bandicoot, Isoodon obesulus. Proceedings of the Royal Society of Queensland 63, 61-63. Mackerras, M.J., (1959). The haematozoa of Australian mammals. Australian Journal of Zoology 7, 105-135. Mallah, H.S., Brown, M.R., Rossi, T.M., Block, R.C., (2010). Parenteral fish oil- associated burr cell anemia. The Journal of pediatrics 156, 324-326 e321. Maraghi, S., Molyneux, D.H., (1989). Studies on cross-immunity in Herpetosoma trypanosomes of Microtus, Clethrionomys and Apodemus. Parasitology Research 75, 175-177. Maslov, D.A., Simpson, L., (1995). Evolution of parasitism in kinetoplastid protozoa. Parasitology Today 11, 30-32. Maxwell, S., Burbidge, A.A., Morris, K. 1996. The 1996 action plan for Australian marsupials and monotremes. 176

McBride, W.J., Hanson, J.P., Miller, R., Wenck, D., (2007). Severe spotted fever group rickettsiosis, Australia. Emerging Infectious Diseases 13, 1742-1744. McCallum, H., (2012). Disease and the dynamics of extinction. Philosophical transactions of the Royal Society of London. Series B, Biological sciences 367, 2828-2839. McInnes, L.M., Gillett, A., Hanger, J., Reid, S.A., Ryan, U.M., (2011). The potential impact of native Australian trypanosome infections on the health of koalas (Phascolarctos cinereus). Parasitology 138, 873-883. McInnes, L.M., Gillett, A., Ryan, U.M., Austen, J., Campbell, R.S.F., Hanger, J., Reid, S.A., (2009). Trypanosoma irwini n. sp (Sarcomastigophora: Trypanosomatidae) from the koala (Phascolarctos cinereus). Parasitology 136, 875-885. McInnes, L.M., Hanger, J., Simmons, G., Reid, S.A., Ryan, U.M., (2010). Novel trypanosome Trypanosoma gilletti sp. (: Trypanosomatidae) and the extension of the host range of Trypanosoma copemani to include the koala (Phascolarctos cinereus). Parasitology 138, 59-70. McMillian, B., Bancroft, J., (1974). On the morphology of Trypanosoma binneyi Mackerras 1959 from the platypus Ornithorhynchus anatinus. International Journal for Parasitology 4, 441-442. Milner, J.D., Hajduk, S.L., (1999). Expression and localization of serum resistance associated protein in Trypanosoma brucei rhodesiense. Molecular and Biochemical Parasitology 104, 271-283. Molina-Portela Mdel, P., Lugli, E.B., Recio-Pinto, E., Raper, J., (2005). Trypanosome lytic factor, a subclass of high-density lipoprotein, forms cation-selective pores in membranes. Molecular and Biochemical Parasitology 144, 218-226. Molyneux, D.H., (1969a). The attachment of Trypanosoma lewisi in the rectum of its vector flea Nosopsyllus fasciatus. Transactions of the Royal Society of Tropical Medicine and Hygiene 63, 117. Molyneux, D.H., (1969b). The fine-structure of the epimastigote forms of Trypanosoma lewisi in the rectum of the flea, Nosopsyllus fasciatus. Parasitology 59, 55-66. Monis, P.T., Andrews, R.H., Saint, C.P., (2002). Molecular biology techniques in parasite ecology. International Journal for Parasitology 32, 551-562. Moreira, D., Lopez-Garcia, P., Vickerman, K., (2004). An updated view of kinetoplastid phylogeny using environmental sequences and a closer outgroup: proposal for a new classification of the class Kinetoplastea. International Journal of Systematic and Evolutionary Microbiology 54, 1861-1875. Morris, J.C., Drew, M.E., Klingbeil, M.M., Motyka, S.A., Saxowsky, T.T., Wang, Z., Englund, P.T., (2001). Replication of kinetoplast DNA: an update for the new millennium. International Journal for Parasitology 31, 453-458. Navarro, M.C., De Lima, A.R., Askue, J., Contreras, V.T., (2003). Morphological comparison of axenic amastigogenesis of trypomastigotes and metacyclic forms of Trypanosoma cruzi. Memorias Do Instituto Oswaldo Cruz 98, 83-91. Nishimura, K., Shima, K., Asakura, M., Ohnishi, Y., Yamasaki, S., (2005). Effects of heparin administration on Trypanosoma brucei gambiense infection in rats. The Journal of Parasitology 91, 219-222. Noyes, H.A., Stevens, J.R., Teixeira, M., Phelan, J., Holz, P., (1999). A nested PCR for the ssrRNA gene detects Trypanosoma binneyi in the platypus and Trypanosoma sp. in wombats and kangaroos in Australia. International Journal for Parasitology 29, 331-339. Ogedengbe, J.D., Hanner, R.H., Barta, J.R., (2011). DNA barcoding identifies Eimeria species and contributes to the phylogenetics of coccidian parasites (Eimeriorina, Apicomplexa, Alveolata). International Journal for Parasitology 41, 843-850.

177

Owen, W.J., ca. 1934-5., (undated). Trypanosoma in the Monotreme. Publication of the Australian Institute of Anatomy, Canberra, ACT. Paparini, A., Irwin, P.J., Warren, K., McInnes, L.M., de Tores, P., Ryan, U.M., (2011). Identification of novel trypanosome genotypes in native Australian marsupials. Veterinary Parasitology 183, 21-30. Pentreath, V.W., Kennedy, G.E., 2004. Pathogenesis of human African trypanosomiasis, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The trypanosomiases. CABI Publishing, Wallingford, pp. 283-301. Pound, C.J., (1905). On Trypanosoma and their presence in the blood of Brisbane rats. Proceedings of the Royal Society Queensland 19, 33-38. Power, M.L., Richter, C., Emery, S., Hufschmid, J., Gillings, M.R., (2009). Eimeria trichosuri: Phylogenetic position of a marsupial coccidium, based on 18S rDNA sequences. Experimental Parasitology 122, 165-168. Reid, S.A., Husein, A., Partoutomo, S., Copeman, D.B., (2001). The susceptibility of two species of wallaby to infection with Trypanosoma evansi. Australian Veterinary Journal 79, 285-288. Richardson, L.R., (1968). Trypanosomes in the crop of an Haemadipsid leech. The Australian Journal of Science 30, 374–375. Rickman, L.R., Robson, J., (1970). The testing of proven Trypanosoma brucei and T. rhodesiense strains by the blood incubation infectivity test. Bulletin of the World Health Organization 42, 911-916. Rifkin, M.R., (1978). Identification of trypanocidal factor in normal human-serum - high-density lipoprotein. Proceedings of the National Academy of Sciences of the United States of America 75, 3450-3454. Roberts, F.H.S., 1970. Australian ticks Melbourne : Commonwealth Scientific and Industrial Research Organization. Robson, J., Rickman, L.R., Allsopp, R., Scott, D., (1972). The composition of the Trypanosoma brucei subgroup in nonhuman reservoirs in the Lambwe Valley, Kenya, with particular reference to the distribution of T. rhodesiense. Bulletin of the World Health Organization 46, 765-770. Rodak, B.F., Fritsma, G.A., Keohane, E.M., 2012. Hematology: clinical principles and applications., Fourth edition Edition. Elsevier Saunders St. Louis, Missouri. Rozsa, L., Reiczigel, J., Majoros, G., (2000). Quantifying parasites in samples of hosts. Journal of Parasitology 86, 228-232. Saiki, R.K., Gelfand, D.H., Stoffel, S., Scharf, S.J., Higuchi, R., Horn, G.T., Mullis, K.B., Erlich, H.A., (1988). Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239, 487-491. Samanovic, M., Molina-Portela, M.P., Chessler, A.D., Burleigh, B.A., Raper, J., (2009). Trypanosome lytic factor, an antimicrobial high-density lipoprotein, ameliorates Leishmania infection. PLoS Pathogens 5, e1000276. Schenkman, S., Guther, M.L.S., Yoshida, N., (1986). Mechanism of resistance to lysis by the alternative complement pathway in Trypanosoma cruzi trypomastigotes - effect of specific monoclonal-antibody. Journal of Immunology 137, 1623-1628. Schneider, C.A., Rasband, W.S., Eliceiri, K.W., (2012). NIH Image to ImageJ: 25 years of image analysis. Nature Methods 9, 671-675. Sehgal, R.N., Jones, H.I., Smith, T.B., (2001). Host specificity and incidence of Trypanosoma in some African rainforest birds: a molecular approach. Molecular Ecology 10, 2319-2327. Seifi, H.A., (1995). Clinical trypanosomosis due to Trypanosoma theileri in a cow in Iran. Tropical Animal Health and Production 27, 93-94.

178

Shastri, U.V., Deshpande, P.D., (1981). Hyalomma-anatolicum-anatolicum (koch, 1844) as a possible vector for transmission of Trypanosoma theileri, laveran, 1902 in cattle. Veterinary Parasitology 9, 151-155. Silva, R.A.M.S., Herrera, H.M., Domingos, L.B.d.S., Ximenes, F.A., Darvila, A.M.R., (1995). Pathogenesis of Trypanosoma evansi infection of dogs and horses: heamatological and clinical aspects. Ciencia Rural Santa Maria 25, 233-238. Sinclair, E.A., (1998). Morphological variation among populations of the quokka, Setonix brachyurus (Macropodidae : Marsupialia), in Western Australia. Australian Journal of Zoology 46, 439-449. Sinclair, E.A., (2001). Phylogeographic variation in the quokka, Setonix brachyurus (Marsupialia : Macropodidae): implications for conservation. Animal Conservation 4, 325-333. Sinclair, E.A., Danks, A., Wayne, A.F., (1996). Rediscovery of Gilbert's Potoroo, Potorous tridactylus, in Western Australia. Australian Mammalogy 19, 69-72. Singh, N., Pathak, K.M., Kumar, R., (2004). A comparative evaluation of parasitological, serological and DNA amplification methods for diagnosis of natural Trypanosoma evansi infection in camels. Veterinary Parasitology 126, 365-373. Slapeta, J.R., Modry, D., Votypka, J., Jirku, M., Obornik, M., Lukes, J., Koudela, B., (2001). Eimeria telekii n.sp (Apicomplexa : Coccidia) from Lemniscomys striatus (Rodentia : Muridae): morphology, pathology and phylogeny. Parasitology 122, 133-143. Smith, A., Telfer, S., Burthe, S., Bennett, M., Begon, M., (2005). Trypanosomes, fleas and field voles: ecological dynamics of a host-vector-parasite interaction. Parasitology 131, 355-365. Sogin, M.L., Elwood, H.J., Gunderson, J.H., (1986). Evolutionary diversity of eukaryotic small-subunit rRNA genes. Proceedings of the National Academy of Sciences of the United States of America 83, 1383-1387. Stephens, J.W.W., Fantham, H.B., (1910). On the peculiar morphologyof a trypanosome from a case of sleeping sickness and the possibility of its being a new species (T. rhodesiense). Proceedings of the Royal Society B85, 223. Stevens, J., Noyes, H., Gibson, W., (1998). The evolution of trypanosomes infecting humans and primates. Memorias Do Instituto Oswaldo Cruz 93, 669-676. Stevens, J., Rambaut, A., (2001). Evolutionary rate differences in trypanosomes. Infection, genetics and evolution : journal of molecular epidemiology and evolutionary genetics in infectious diseases 1, 143-150. Stevens, J.R., Brisse, S., 2004. Systematics of trypanosomes of medical and veterinary importance, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The trypanosomiases. CABI Publishing, Wallingford, pp. 1-23. Stevens, J.R., Noyes, H., Dover, G.A., Gibson, W.C., (1999a). The ancient and divergent origins of the human pathogenic trypanosomes, Trypanosoma brucei and T. cruzi. Parasitology 118, 107-116. Stevens, J.R., Noyes, H.A., Schofield, C.J., Gibson, W., (2001). The molecular evolution of Trypanosomatidae. Advances in Parasitology 48, 1-56. Stevens, J.R., Teixeira, M.M.G., Bingle, L.E.H., Gibson, W.C., (1999b). The taxonomic position and evolutionary relationships of Trypanosoma rangeli. International Journal for Parasitology 29, 749-757. Tamura, K., Dudley, J., Nei, M., Kumar, S., (2007). MEGA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Molecular Biology and Evolution 24, 1596-1599.

179

Tamura, K., Stecher, G., Peterson, D., Filipski, A., Kumar, S., (2013). MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Molecular Biology and Evolution 30, 2725-2729. Tanowitz, H.B., Gumprecht, J.P., Spurr, D., Calderon, T.M., Ventura, M.C., Raventos- Suarez, C., Kellie, S., Factor, S.M., Hatcher, V.B., Wittner, M., (1992). Cytokine gene expression of endothelial cells infected with Trypanosoma cruzi. The Journal of Infectious Diseases 166, 598-603. Taylor, K., Authie, E.M.L., 2004. Pathogenesis of animal trypanosomiasis, In: Maudlin, I., Holmes, P.H., Miles, M.A. (Eds.) The trypanosomiases. CABI Publishing, Wallingford, pp. 331-353. Teixeira, A.R., Hecht, M.M., Guimaro, M.C., Sousa, A.O., Nitz, N., (2011). Pathogenesis of Chagas' disease: parasite persistence and autoimmunity. Clinical Microbiology Reviews 24, 592-630. Telleria, J., Barnabe, C., Hide, M., Banuls, A.L., Tibayrenc, M., (2004). Predominant clonal evolution leads to a close parity between gene expression profiles and subspecific phylogeny in Trypanosoma cruzi. Molecular and Biochemical Parasitology 137, 133-141. Thekisoe, O.M.M., Honda, T., Fujita, H., Battsetseg, B., Hatta, T., Fujisaki, K., Sugimoto, C., Inoue, N., (2007). A trypanosome species isolated from naturally infected Haemaphysalis hystricis ticks in Kagoshima Prefecture, Japan. Parasitology 134, 967-974. Thompson, C.K., Botero, A., Wayne, A.F., Godfrey, S.S., Lymbery, A.J., Thompson, R.C., (2013). Morphological polymorphism of Trypanosoma copemani and description of the genetically diverse T. vegrandis sp. nov. from the critically endangered Australian potoroid, the brush-tailed bettong (Bettongia penicillata (Gray, 1837)). Parasites & Vectors 6, 121. Thompson, C.K., Godfrey, S.S., Thompson, R.C., (2014a). Trypanosomes of Australian mammals: A review. International Journal for Parasitology. Parasites and Wildlife 3, 57-66. Thompson, C.K., Wayne, A.F., Godfrey, S.S., Thompson, R.C., (2014b). Temporal and spatial dynamics of trypanosomes infecting the brush-tailed bettong (Bettongia penicillata): a cautionary note of disease-induced population decline. Parasites & Vectors 7, 169. Tobler, U., Borgula, A., Schmidt, B.R., (2012). Populations of a susceptible amphibian species can grow despite the presence of a pathogenic chytrid fungus. PLoS One 7, e34667. Tomlinson, S., Jansen, A.M., Koudinov, A., Ghiso, J.A., Choi-Miura, N.H., Rifkin, M.R., Ohtaki, S., Nussenzweig, V., (1995). High-density-lipoprotein- independent killing of Trypanosoma brucei by human serum. Molecular and Biochemical Parasitology 70, 131-138. Tomlinson, S., Raper, J., (1998). Natural human immunity to trypanosomes. Parasitology Today 14, 354-359. Truc, P., Buscher, P., Cuny, G., Gonzatti, M.I., Jannin, J., Joshi, P., Juyal, P., Lun, Z.R., Mattioli, R., Pays, E., Simarro, P.P., Teixeira, M.M., Touratier, L., Vincendeau, P., Desquesnes, M., (2013). Atypical human infections by animal trypanosomes. PLoS Neglected Tropical Diseases 7, e2256. Turner, A.W., Murnane, D., (1930). Trypanosomes in the blood of Victorian animals. 1. A preliminary note on the occurrence of Trypanosoma theileri in the blood of cattle. 2. On the presence of Trypanosoma melophagium in the blood of Victorian sheep, and its transmission by the sheep ''tick'', Melophagus ovinus. Journal of the Council for Scientific and Industrial Research, Australia 3, 120- 122. 180

Turner, C.M., McLellan, S., Lindergard, L.A., Bisoni, L., Tait, A., MacLeod, A., (2004). Human infectivity trait in Trypanosoma brucei: stability, heritability and relationship to sra expression. Parasitology 129, 445-454. Tyler, K.M., Engman, D.M., (2001). The life cycle of Trypanosoma cruzi revisited. International Journal for Parasitology 31, 472-481. Uche, U.E., Jones, T.W., (1992). Pathology of experimental Trypanosoma evansi infection in rabbits. Journal of Comparative Pathology 106, 299-309. Uilenberg, G. 1998. A field guide for diagnosis, treatment and prevention of African animal trypanosomosis (Food and Agriculture Organization of the United Nations, Rome. ). Urbina, J.A., (1999). Chemotherapy of Chagas' disease: the how and the why. Journal of Molecular Medicine 77, 332-338. Valentin, G., (1841). Ueber ein Entozoon im Blute von Salmo fario. Archiv für Anatomie, Physiologie und Wissenschaftliche Medicin, 435. Van de Peer, Y., De Wachter, R., (1994). TREECON for Windows: a software package for the construction and drawing of evolutionary trees for the Microsoft Windows environment. Computer Applications in the Biosciences 10, 569-570. Vanhollebeke, B., Truc, P., Poelvoorde, P., Pays, A., Joshi, P.P., Katti, R., Jannin, J.G., Pays, E., (2006). Human Trypanosoma evansi infection linked to a lack of apolipoprotein L-I. The New England journal of Medicine 355, 2752-2756. Vassella, E., Straesser, K., Boshart, M., (1997). A mitochondrion-specific dye for multicolour fluorescent imaging of Trypanosoma brucei. Molecular and Biochemical Parasitology 90, 381-385. Verma, A., Manchanda, S., Kumar, N., Sharma, A., Goel, M., Banerjee, P.S., Garg, R., Singh, B.P., Balharbi, F., Lejon, V., Deborggraeve, S., Singh Rana, U.V., Puliyel, J., (2011). Trypanosoma lewisi or T. lewisi-like infection in a 37-day- old Indian infant. American Journal of Tropical Medicine and Hygeine 85, 221- 224. Vickerman, K., (1985). Developmental cycles and biology of pathogenic trypanosomes. British Medical Bulletin 41, 105-114. Vickerman, K., (1994). The evolutionary expansion of the trypanosomatid flagellates. International Journal for Parasitology 24, 1317-1331. Woodruff, A.W., Prankerd, T.A., Ormerod, W.E., (1973). Mechanisms involved in anemia associated with infection and splenomegaly in tropics. Transactions of the Royal Society of Tropical Medicine and Hygiene 67, 313-328. Wyatt, K.B., Campos, P.F., Gilbert, M.T., Kolokotronis, S.O., Hynes, W.H., DeSalle, R., Ball, S.J., Daszak, P., MacPhee, R.D., Greenwood, A.D., (2008). Historical mammal extinction on Christmas Island (Indian Ocean) correlates with introduced infectious disease. PLoS One 3, e3602. Xong, H.V., De Baetselier, P., Pays, E., Magez, S., (2002). Selective pressure can influence the resistance of Trypanosoma congolense to normal human serum. Experimental Parasitology 102, 61-65. Xong, H.V., Vanhamme, L., Chamekh, M., Chimfwembe, C.E., Van Den Abbeele, J., Pays, A., Van Meirvenne, N., Hamers, R., De Baetselier, P., Pays, E., (1998). A VSG expression site-associated gene confers resistance to human serum in Trypanosoma rhodesiense. Cell 95, 839-846. Yang, R., Brice, B., Bennett, M.D., Eliott, A., Ryan, U., (2013). Novel Eimeria sp isolated from a King's skink (Egernia kingii) in Western Australia. Experimental Parasitology 133, 162-165. Yang, R., Fenwick, S., Potter, A., Elliot, A., Power, M., Beveridge, I., Ryan, U., (2012). Molecular characterization of Eimeria species in macropods. Experimental Parasitology 132, 216-221. 181

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