Unlocking Archived Ticks to Investigate a Novel Putative Pathogen

UNLOCKING ARCHIVED TICKS TO INVESTIGATE A NOVEL PUTATIVE PATHOGEN

Anna-Sheree Krige

A. S. KRIGE

This thesis is presented for the degree of Honours in Molecular Biology at Murdoch University 2017 Declaration

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

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Anna-Sheree Krige

Abstract

Ticks, as vectors of disease-causing bacteria, represent a significant threat to human and animal health. Of particular concern is the genus Borrelia, which contains several recognised tick-borne pathogens of global health importance. In 2015, the microbiomes within modern Australian ticks were profiled using next- generation sequencing (NGS). Complex communities of bacterial species were revealed, including a novel Borrelia sp. that was identified within a tick parasitising an Australian echidna, Tachyglossus aculeatus. This bacterium, named ‘Candidatus Borrelia tachyglossi’, after its association with the echidna tick, Bothriocroton concolor, forms the fourth known Borrelia group. Following this first report of Borrelia in native Australian ticks, additional questions have emerged concerning whether this bacterium resides in other echidna-biting ticks, and if it has a historic presence in Australia.

Around 1,725 tick specimens from 89 registered echidna hosts were collected between 1928 and 2013. Of these, 850 ticks were selected for examination in this study. A total of eight species from three tick genera were morphologically identified. The most common species was B. concolor (89.2%). Interestingly, Amblyomma fimbriatum (22, 2.6%) and Amblyomma triguttatum (19, 2.2%) ticks were recorded for the first time from echidnas. A NGS survey of the bacterial communities within 66 echidna ticks, targeting the V4 region of the 16S rRNA gene, revealed a diverse range of preserved bacteria. Genera of interest included Borrelia (6, 9.1%), Coxiella (19, 28.8%), Ehrlichia (2, 3.0%), Francisella (5, 7.6%), and Rickettsia (23, 34.8%). While NGS of archived echidna tissue biopsies were negative for Borrelia, genus-specific PCR assays amplified the flaB (378 bp) locus in two skin biopsies (2/34; 5.9%) and nine ticks (9/160; 5.6%), with a 99.5-100% similarity to ‘Candidatus B. tachyglossi’ genotypes B and C. Of these positive samples, six ticks and both skin biopsies (6/9, 66.7%; 2/2, 100%) amplified the longer 16S rRNA (1,087 bp) locus, with a 98.2-99.6% similarity to ‘Candidatus B. tachyglossi’ genotype B. Phylogenetic positioning was consistent with previous reports, suggesting ‘Candidatus B. tachyglossi’ formed the fourth Borrelia clade.

This is the first morphological audit of echidna ticks and NGS survey of the archived tick microbiome. This study has confirmed ‘Candidatus B. tachyglossi’ in Australian echidna ticks in the recent past and for the first time, revealed Borrelia in echidnas. However, whether echidnas are a reservoir for this bacterium could not be established. Additional tick transmission analyses and the examination of echidna blood samples are necessary for confirmation.

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Acknowledgements

I would first like to thank my supervisor, Dr. Charlotte Oskam, for your support, advice, and guidance over the course of this research project.

There are many people whom I am grateful to for their help and encourgagement. To former research assistant, Annachiara Codello, for ‘showing me the ropes’ in the lab and your kindness. To Kimberly Loh, for introducing me to tick identification and pointing out that ‘those are not eyes’. Also, for your help with Geneious software and providing feedback on one of my earliest drafts, I thank you. To Alexander Gofton, your knowledge and assistance with my library preparation and NGS run was invaluable. Thank you for your time and perseverance with the challenges experienced using this technology. A further thank you goes to the rest of the team at the Vector and Water-Borne Pathogen Research Group for your kind support and advice throughout the year. And to Murdoch University, thank you for granting me a Murdoch University Academic Excellence Award.

Finally, I would like to thank my family for their unwavering support and encouragement throughout my honours year. No matter what direction I have decided to pursue, Mum and Dad have always been there to reassure and inspire. And to my two adoring fans and most precious companions, Toby and Lulu, I am so thankful for your company during the many long hours in front of the computer typing up my thesis. Your presence has always given me comfort and to you I dedicate my work.

Table of Contents

Declaration…………………………………………………………………………………………………………i Abstract…………………………………………………………………………………………………………….ii Acknowledgements…………………………………………………………………………………………...iv Table of Contents……………………………………………………………………………………………….v List of Figures……………………………………………………………………………………………………ix List of Tables…………………………………………………………………………………………………….xi List of Abbreviations………………………………………………………………………………………...xii

CHAPTER ONE: INTRODUCTION………………………………………………………………………1 1. Introduction………………………………………………………………………………………………....2 1.1 Ticks……………………………………………………………………………………………………...4 1.1.1 Tick morphology……………………………………………………………………………..5 1.1.2 Tick life cycle…….…………………………………………………………………………….7 1.1.3 Ticks in Australia…………………………………………………………………………….8 1.1.4 Tick hosts………………………………………………………………………………………..9 1.1.4.1 Echidnas and echidna-biting ticks…………………………………………9 1.2 Tick-borne pathogens and the tick microbiome……………………………………..10 1.2.1 Tick-borne diseases (TBDs) – a global perspective …………………………12 1.2.2 TBDs in Australia…………………………………………………………………………..13 1.2.3 The genus Borrelia………………………………………………………………………...14 1.2.4 Borrelia in Australia………………………………………………………………………16 1.2.4.1 Borrelia in Australian animals……………………………………………..17 1.2.4.2 Borrelia in Australian ticks………………………………………………….19 1.3 Detection methods of tick-borne pathogens and the tick microbiome………………………………………………………………………………………….20 1.3.1 Polymerase Chain Reaction (PCR)………………………………………………….21 1.3.2 Sanger and next-generation sequencing (NGS) – revolutionary methods of DNA detection……………………………………..22 1.4. Museum specimens – an unexplored ‘gold mine’…………………………………...24 1.4.1 Ancient DNA (aDNA)……………………………………………………………………..25 1.4.1.1 aDNA and NGS – a ‘perfect’ pair…………………………………………..27 1.4.2 Using museum specimens to investigate pathogens………………………..28 v

1.4.2.1 Using museum specimens to detect tick-borne pathogens…………………………………………………………………………..28 1.4.3 Non-destructive sampling – a method for historical preservation………………………………………………………………………………….30 1.5. Research Aims and Hypotheses………………………………...……………………...... 31

CHAPTER TWO: MATERIALS AND METHODS………………………………………………...33 2. Overview…………………………...………………………………………………...………………………34 2.1 Tick collection and preparation…………………………...………………………………..35 2.1.1 Archived ticks with correlating host tissue…………………………...………...36 2.2 Tick identification…………………………...……………………………………………………36 2.3 Sample collection…………………………...…………………………………………………….37 2.4 DNA extraction…………………………...………………………………………………………..37 2.4.1 Archived echidna ticks…………………………...……………………………………...38 2.4.2 Archived echidna tissue…………………………...…………………………………….39 2.5 Library preparation and NGS…………………………...…………………………………...39 2.6 NGS analysis…………………………...……………………………………………….…………...41 2.7 PCR assays…………………………...………………………………………………………………42 2.7.1 16S Mam qPCR assay of echidna tissue…………………………...……………...44 2.7.1.1 16S Mam conventional PCR assay of correlating echidna tissue ticks…………………………...…………………………….….44 2.7.2 Borrelia-specific PCR assays of echidna tissue and ticks…………………..45 2.7.2.1 flaB nested-PCR assay…………………………...……………………………45 2.7.2.2 Borrelia-specific 16S nested-PCR assay…………………………...…..46 2.8 Gel electrophoresis…………………………...………………………………………………….47 2.9 PCR purification…………………………...………………………………………………………47 2.10 Sanger sequencing and BLAST…………………………...……………………………….48 2.11 Phylogenetic analysis…………………………...………………………………………….…49 2.12 Percentage of Borrelia-infected echidna ticks………………………………………50

CHAPTER THREE: RESULTS……………………………………………………………………………51 3. Results…………………………………………………………………………………………………………52 3.1 Morphological identification of archived echidna-biting ticks………………...52 3.1.1 Sample selection for DNA extraction and analyses…………………………..54 vi

3.2 NGS………….…………………………………………………………………………………………..56 3.3 PCR assays………..………………………………………………………………………………….62 3.3.1 16S Mam qPCR assay of echidna tissue………………………..……..…………..62 3.3.1.1 16S Mam PCR assay of correlating tissue ticks…………………..…64 3.3.2 flaB nested-PCR assay……………………………………….……………….…………..66 3.3.2.1 The flaB gene in echidna tissue…………………………………………...68 3.3.2.2 The flaB gene in echidna ticks……………………………………………..70 3.3.3 Borrelia-specific 16S nested-PCR assay………………………………………..…72 3.4 Sanger sequencing and BLAST…………………………………..………………………….72 3.5 Phylogenetic analysis………………………..………………………………………………….73 3.6 Percentage of Borrelia-infected echidna ticks….………………………………….….76

CHAPTER FOUR: DISCUSSION AND CONCLUSION…………………………………………..79 4. Discussion……………………………………………………………………………………………………80 4.1 Morphological identification of archived echidna-biting ticks………………...80 4.2 NGS bacterial profiling of archived echidna-biting ticks…………………………82 4.3 Detecting a novel Borrelia species in archived echidna-biting ticks………...85 4.4 Future directions………………………………………….………………………………………88 4.5 Conclusion………………………………………….………………………………………………..89

REFERENCES…………………………………………………………………………………………………..90 References……………………………………………………………………………………………………....91

APPENDICES…………………………………………………………………………………………………114

Appendix A…………………………………………………………………………………………………….115 Table A.1. Metadata spreadsheet of archived echidna ticks………………………..115 Table A.2. Sub-sample of archived echidna ticks for DNA extraction…………..144 Table A.3. Metadata spreadsheet of archived echidna tissue biopsies…………155 Appendix B…………………………………………………………………………………………………….157 Table B.1. Comparative character matrix for the genera Amblyomma, Bothriocroton, and Ixodes………………………………………………………….157 Appendix C…………………………………………………………………………………………………….159

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Table C.1. NGS mapping file……………………………………………………………………...159 Table C.2. Legend for taxonomic assignment of bacterial phyla in QIIME………………………………………………………………………………………167 Table C.3. Legend for taxonomic assignment of bacterial genera in QIIME………………………………………………………………………………………168 Figure C.1. Bacterial diversity in echidna tissue and tick species according to instar, generated in QIIME…………………………………...169 Appendix D…………………………………………………………………………………………………....170 Figure D.1. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples of 1:10 dilution ratio…………………………………………………………………..170 Figure D.2. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples at neat concentration…………………………………………………………………………171 Figure D.3. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples at neat concentration………………………………………………………………………….172 Figure D.4. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples at 1:10 dilution ratio…………………………………………………………………………..173

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List of Figures

Figure 1.1. Phylogeny of the families and subfamilies for the order Ixodida……………………………………………………………………………………………..4 Figure 1.2. Morphological features of (A) ixodid (hard) ticks and (B) argasid (soft) ticks…………………………………………………………………………….6 Figure 1.3. Life cycle of a three-host ixodid tick…………………………………………………...8 Figure 1.4. Transmission cycles of a tick……………………………………………………………12 Figure 1.5. The timeline of Borrelia-related discoveries and reported observations in Australia.………………………………………………………………..17 Figure 2.1. A methodological overview of the current study……………………………….34 Figure 3.1. Larval pool of ticks recorded as Bothriocroton concolor…………………….52 Figure 3.2. Nymph Bothriocroton concolor tick…………………………………………………..53 Figure 3.3. Adult Bothriocroton concolor ticks (A) dorsal view of B. concolor female and male, and (B) ventral view of B. concolor female and male………………………………………………………………53 Figure 3.4. The number of reads (per million) generated during NGS that passed filter……………………………………………………………………………..57 Figure 3.5. Bacterial diversity in Australian echidna ticks and echidna tissue (T. aculeatus)………………………………………………………………………...60 Figure 3.6. Bacterial diversity in Australian B. concolor echidna ticks according to instar………………………………………………………………………….61 Figure 3.7. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples of 1:100 dilution ratio……………………………………………………………………………………………….63 Figure 3.8. Gel electrophoresis (1%) images (A, B, and C) of 16S Mam PCR products from neat echidna ticks sourced from correlating echidna hosts 1 to 7………………………………………………………………………...65 Figure 3.9. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples of 1:100 dilution ratio…………………….67 Figure 3.10. Gel electrophoresis (1%) image of flaB PCR products from echidna tissue of 1:100 dilution ratio originating from echidna hosts 1 to 8……………………………………………………………………………………69 ix

Figure 3.11. Gel electrophoresis (1%) image of flaB PCR products from neat echidna (n = 46) ticks originating from echidna hosts 1 to 7…………………………………………………………………………………………….71 Figure 3.12. Phylogenetic analysis of 378 bp flaB gene consensus sequence………….……………………………………………………………………………74 Figure 3.13. Phylogenetic analysis of 1,087bp Borrelia 16S rRNA gene consensus sequence………………………………………………………………………75 Figure 3.14. A comparison of the Australian geographical location of Borrelia positive B. concolor ticks, between the current study and the investigation by Loh et al. (2016)………………………………………78

List of Tables

Table 2.1. Number of echidna host registrations and an estimated number of ticks sourced from natural history and museum collections according to Australian state and territory locality…………………………….35 Table 2.2. Number of ticks according to instar removed from frozen archived echidna specimens, sourced from the Queensland Museum collection……………………………………………………………………………36 Table 2.3 Primers used for NGS and PCR amplification. Primer sequences, annealing temperatures, and expected amplicon sizes are included……………………………………………………………………………………………43 Table 3.1. Number of instars for eight echidna-biting tick species morphologically identified from Australian archived collections………...54 Table 3.2. Tick species with total number of host registrations according to state and territory location, including the number of hosts selected for tick DNA extraction, and subsequent number of tick extractions per species locality………………………………………………………….55 Table 3.3. Ticks selected for DNA extraction.……………………………………………………..56 Table 3.4. NGS sequencing statistics for bacterial 16S rRNA reads……………………...58 Table 3.5. Summary of the QLD B. concolor ticks according to instar that were positive for Borrelia genes in terms of percentage, using nested-PCR (with Sanger sequencing) and NGS technologies……………..77

List of Abbreviations

A Adenine BLASTaDNA BasicAncient local DNA alignment search tool BIC Bayesian Information Criterion bp Base pairs C CytosineBase pair DNA Deoxyribonucleic acid dNTP Deoxynucleotide triphosphate EDTA Ethylenediaminetetraacetic acid etbp al. and others EXT  Extraction blank control flaB Flagellin gene G Guanine GTRg General Gram time reversible Kb Kilobase s LBh LymeHour borreliosis mg Milligram min kb MinuteKilobase mL Millilitre mm Millimetre μg Microgram μL microlitre μm Micrometre MgCl 2 Magnesium chloride nμM Number Micromolar of samples nmol Nanomol NCBI National Centre for Biotechnology Information NTC No -template control PCR NGS Polymerase Next-generation chain sequencing reaction qPCR OTU QuantitativeOperational taxonomic PCR unit rDNA Ribosomal DNA RNA Ribonucleic acid rRNA REP RibosomalReptile-associated RNA borreliae rpm Revolutions per minute s RFB SecondRelapsing fever borreliae sp./spp. Species T SLO Thymine Spirochaete -like object TAE Tris acetate EDTA buffer Taq Thermos aquaticus DNA polymerase TBDs Tick-borne diseases UV Ultra violet light w/v U WeightUnit of enzymaticof solute per activity volume of solvent x Times 16S 16S bacterial gene 5’ Phosphate -terminus of DNA molecule ~ 3’ ApproximatelyHydroxyl-terminus of DNA molecule & And > Greater than

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< Less than - Negative % Percentage °C Degrees Celsius  Present  Absent



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CHAPTER ONE: INTRODUCTION

1 1. Introduction

Ticks (Acari: Ixodida) are obligate haematophagous (blood-feeding) ectoparasites of a number of vertebrate hosts including mammals, birds, reptiles, and occasionally amphibians (Sonenshine & Roe, 2013). To date, more than 850 tick species have been discovered, from which approximately 10% are determined as capable of transmitting disease-causing agents (Barker & Murrell, 2004; Jongejan & Uilenberg, 2004). Ticks pose a major health threat to humans, companion animals, livestock, and wildlife, because of their ability to vector a diverse range of pathogenic microorganisms that include protozoa, fungi, viruses, and bacteria (Parola & Raoult, 2001; Jongejan & Uilenberg, 2004).

Recent profiling of the bacterial communities in modern Australian ticks using next-generation sequencing (NGS), revealed a novel bacterium within the genus Borrelia in a tick removed from an Australian echidna, Tachyglossus aculeatus (Gofton et al., 2015a). A subsequent study of native Australian Bothriocroton concolor ticks removed exclusively from echidna hosts determined the presence of this novel Borrelia sp. in echidna ticks (Loh et al., 2016). This bacterium has since been named ‘Candidatus Borrelia tachyglossi’ after its association with the echidna tick B. concolor and proposed vertebrate host (Loh et al., 2017). This was the first report of Borrelia in native Australian ticks. Further questions have consequently emerged concerned at whether this bacterium resides in other echidna-biting ticks and therefore has an association with echidnas. In addition, the past presence of this putative pathogen in Australia remains unknown. As museum archives house a typically extensive and readily available resource, utilising historic specimens can offer a vast repository of genetic data to further address biological questions including systematics, origin, occurrence, and past biogeography. Hence, ticks archived in museum collections provide an avenue for investigating the likely source and former prevalence of pathogens and novel microorganisms.

This chapter reviews the literature pertaining to ticks of the Ixodidae family, commonly referred to as hard ticks, due to their role in transmitting disease- causing pathogens in humans and animals of medical and veterinary significance.

2 Structurally, the following sections review:

1.1 Ticks. This includes tick morphology, and the ixodid tick life cycle, with a focus prescribed to the presence of native and introduced tick species in Australia. Tick hosts are briefly discussed with specific mention made concerning the Australian echidna (T. aculeatus) and associated tick ectoparasites.

1.2 Tick-borne pathogens and the tick microbiome. Tick-borne diseases (TBDs) in Australia and their causative agents are discussed. The genus Borrelia is described in regards to current phylogeny and associated diseases, with a concise synopsis of spirochaete morphology. While mention is made regarding the various Borrelia pathogens and related predominant diseases: Lyme borreliolis (LB) and relapsing fever borreliae (RF), a detailed discussion of these diseases is beyond the scope of this review. Emphasis will be placed on reviewing the literature concerning the detection and identification of Borrelia in Australia, highlighting current Borrelia findings in Australian ticks.

1.3 Detection methods of tick-borne pathogens and the tick microbiome. A chronological order of the principal molecular tools and sequencing technologies used to profile bacteria in contemporary studies will be evaluated. Specifically, techniques used to identify Borrelia are examined. Additionally, though many different platforms have been developed for NGS, highlighting them all in terms of their various advantages and disadvantages is beyond the scope of this review, though a number of excellent reviews are available by Mardis (2013), van Dijk et al. (2014), and Goodwin et al. (2016) for further reference.

1.4 Museum specimens – an underutilised ‘gold mine’. Ancient DNA (aDNA) applications and the advantage of incorporating NGS in aDNA retrieval and analysis is explained. An overview of the potential for museum specimens to be used in disease investigation is proposed, with a specific review of studies detecting tick-borne pathogens in archived specimens. Finally a concise report of non-destructive sampling methods for specimen preservation is mentioned.

3 1.1 Ticks

As arthropods, ticks belong to the largest and most diverse phylum, representing approximately 80% of all known animals, and occur in almost every region of the world (Parola & Raoult, 2001). Ticks belong to the subphylum Chelicerata, within the class of arachnids (Arachnida) together with mites, scorpions, and spiders. Ticks are further classified into the subclass Acari, and suborder Ixodida (Figure 1.1), which includes three tick families: Ixodidae (hard ticks) with 12 extant genera and more than 700 species, Argasidae (soft ticks) with four genera and over 190 species, and Nuttalliellidae, which shares characteristics of both Ixodidae and Argasidae and is represented by a single genus confined to Southern Africa (Black & Piesman, 1994; Parola & Raoult, 2001; Barker & Murrell, 2004; Guglielmone et al., 2010).

Figure 1.1. Phylogeny of the families and subfamilies for the order Ixodida. Revised from Barker and Murrell (2004) and Nava et al. (2009).

Traditionally, the relationship among tick genera was based on the tick evolution work by Hoogstraal, who observed ticks morphologically and devised a phylogeny illustrating the relationship between tick lineages (Hoogstraal & Aeschlimann, 1982). Much of Hoogstraal’s original work regarding tick systematics remains,

4 however in recent years several revisions have been proposed following DNA sequencing and phylogenetic analyses. A recent amendment concerned the genus Aponomma, which was identified as polyphyletic by Kaufman in 1972. Based initially on morphological differences (Kaufman, 1972), this taxon was later resolved by molecular evidence including DNA sequence disparities, and distinct groups were identified within the genus Aponomma; the genus Amblyomma for the typical more primitive Aponomma, and a new genus termed Bothriocroton, representing the genetically distinctive ‘indigenous Australian’ Aponomma (Dobson & Barker, 1999; Klompen et al., 2000; Klompen et al., 2002).

1.1.1 Tick morphology

The external body of a tick consists of three anatomical regions: the capitulum, the idiosoma or body region, and the podosoma or legs (Sonenshine & Roe, 2013). The capitulum resembles a head and contains the tick’s mouthparts, sensory organs, and the hypostome that acts like a barb and with recurved teeth, anchors the tick to the hosts skin (Parola & Raoult, 2001). Although clearly visible in hard ticks it is less so in soft ticks (Sonenshine & Roe, 2013). The basal portion of the capitulum is used in the morphological identification of tick species and is based on structures that project from the posterior corners of the dorsal side (cornua) or ventral side (auriculae) (Hillyard, 1996). Other key characteristics which aid in identifying ixodid ticks to the genus and species level include the length of the mouthparts (particularly the palpal articles), the presence or absence of eyes and of festoons, colour and markings on their dorsal surface, as well as shape and orientation of their anal groove (Sonenshine & Roe, 2013; Barker & Walker, 2014). Additionally, numerous taxonomic keys are available for identifying tick species inhabiting different regions of the world (Hoogstraal, 1956; Roberts, 1970; Hillyard, 1996). On the dorsal surface, ixodid ticks contain a shield or scutum, which acts as a major discrepancy between hard ticks and soft ticks, as shown in Figure 1.2. Moreover, though many ticks lack eyes they do possess a variety of sensory organs such as the Haller’s organ and external hair-like structures, deemed significant in enabling ticks to locate their host (Waladde, 1987; Sonenshine & Roe, 2013).

5 For copyright purposes please refer to Figures 2 and 3 in Barker and Walker (2014)

Figure 1.2. Morphological features of (A) ixodid (hard) ticks and (B) argasid (soft) ticks. Hard ticks are distinguished from soft ticks by the presence of a hardened shield called a scutum that covers part of the dorsal region of female hard ticks and the entire dorsal region of male hard ticks. In addition, hard ticks contain a prominent capitulum (head and mouthparts), which project forward from the body. In comparison, soft ticks lack a hardened scutum and their capitulum is positioned beneath the body. As female and male soft ticks are morphologically similar, a female tick diagram is sufficient representation of the argasid family. Sourced from Barker and Walker (2014).

Internally, like other arthropods, ticks have an open circulatory system filled with haemolymph, a circulating fluid that bathes all the various internal organs and tissues. Consequently, this open system enables microorganisms and putative pathogens harboured within the midgut (blood meal storage organ) of ticks to

6 potentially invade and multiply in organs and fluids such as the tick’s haemolymph and saliva (Parola & Raoult, 2001; Sonenshine & Roe, 2013; Qiu et al., 2014). This is a significant factor when considering the transmission route of infectious agents between a tick and their vertebrate host.

1.1.2 Tick life cycle

Ticks have three instars: larva, nymph, and mature adult. As obligate blood feeders at all life stages ticks are deemed parasitic (Walker et al., 2003). Each active stage feeds only once in its life, however female ticks have been known to reattach and continue feeding if forcibly removed from a host (Sonenshine & Roe, 2013). In addition, hard ticks have a life cycle of approximately 1-6 years and are generally less able to survive without a blood meal compared to their soft tick counterparts (Anderson & Magnarelli, 1993a; Anderson & Magnarelli, 2008; Sonenshine & Roe, 2013).

Ticks can be three-, two-, or one-host arthropods, though as most Ixodidae require three different hosts to complete their life cycle (Parola & Raoult, 2001), this review will focus on three-host ticks, as illustrated in Figure 1.3. Initially, the questing six-legged sexually indistinct larva that emerges from the egg climbs onto a host and using its mouthparts, cuts the epidermis and dermis of the hosts skin and inserts its feeding organ, the hypostome, to initiate feeding. This can continue for approximately 3-5 days before the larva detaches from the host to moult to the nymphal stage (Sonenshine & Roe, 2013). After metamorphosis, the tick acquires an extra pair of legs. The now eight-legged nymph, also sexually indistinct, will attach to a second host (same or different species), and take a blood meal (Parola & Raoult, 2001). After approximately 4-8 days, the nymph drops from the host and moults into the adult stage. At this final evolved stage, the sexually distinct and mature adult tick infests a third host (same or different species) feeding for anywhere between 2-15 days depending on various factors including tick species and gender, host species, and site of attachment (Parola & Raoult, 2001; Sonenshine & Roe, 2013).

7 Regarding reproduction, after inseminating a female, which apart from Ixodes species usually takes place on the host, the male tick dies (Barker & Walker, 2014). Female ticks typically feed for longer periods than males, and only after ovipositing several thousand eggs, eventually die (Parola & Raoult, 2001). Within four weeks, 2000 or more eggs hatch into larvae, primed to quest for blood (Hillyard, 1996; Sonenshine & Roe, 2013; Barker & Walker, 2014).

For copyright purposes please refer to Figure 5 in Barker and Walker (2014)

Figure 1.3. Life cycle of a three-host ixodid tick. Example is Rhipicephalus sanguineus, the brown dog tick. Adapted from Barker and Walker (2014).

1.1.3 Ticks in Australia

Seventy-one native and introduced tick species have been described in Australia (Roberts, 1970; Barker et al., 2014; Ash et al., 2017). The tick Ixodes woyliei n. sp. is the most recent description to date (Ash et al., 2017). Based on external features, 57 species are recorded as hard ticks (family Ixodidae) and 14 as soft ticks (family Argasidae). Sixty-six of these species are endemic to Australia and comprise ticks of the subfamilies Amblyomminae, Argasinae, Bothriocrototinae, Haemaphysalinae, Ixodinae, and Ornithodorinae. The remaining five species identified are the poultry tick Argas persiscus, the bush tick Haemaphysalis

8 longicornis, the spinose ear tick Otobius megnini, and two tick species of the introduced genera Rhipicephalinae, which are Rhipicephalus sanguineus and Rhipicephalus (Boophilus) australis, the brown dog tick and Australian cattle tick respectively (Barker & Walker, 2014). Currently there are 55 Australian tick species that are known to parasitise wild mammals, birds, and reptiles (Barker & Walker, 2014; Barker et al., 2014; Ash et al., 2017).

1.1.4 Tick hosts

Ticks and associated pathogens have coevolved with many wildlife hosts and are believed to generally reside in a benign co-existence, however when in contact with humans, companion animals, or livestock, they can serve as reservoirs for tick-transmitted diseases (Jongejan & Uilenberg, 2004). Throughout the world many small mammals act as reservoirs or amplifying hosts of bacteria and other pathogens (Mills & Childs, 1998; Durden, 2006). Considered important tick hosts, more than 87 tick species are known to parasitise small mammals within the order Rodentia (Horak et al., 2002; Kolonin, 2007). Similarly, although insectivores such as hedgehogs, shrews, and moles, as well as the monotremes: platypus and echidna, are not as frequently parasitised, species of Ixodidae have been documented as present on these small mammal hosts, with some host specific (Durden, 2006; Kolonin, 2007; Barker & Walker, 2014).

1.1.4.1 Echidnas and echidna-biting ticks

The echidna belongs to the order Monotremata and is part of the family Tachyglossidae (Augee et al., 2006). Classified as a myrmecophage (ant and termite specialist), though commonly referred to as a spiny anteater, the echidna is a medium-sized mammal with a dorsal covering of stout spines among a fur coat, ranging from light brown to black in colour (Collins 1973; Nicol & Andersen, 2007). There are two extant species of echidna: the short-beaked echidna (T. aculeatus) and the long-beaked echidna (Zaglossus bruijini). The short-beaked species exclusively inhabits mainland Australia and Tasmania (Augee et al., 2006), while the long-beaked echidna populates New Guinea (Augee et al., 2006). As indigenous wildlife that are almost ubiquitous in their distribution in Australia

9 (Nicol & Andersen, 2007), the short-beaked echidna has been observed in recent studies concerning bacteria associated with native Australian ticks (Gofton et al., 2015a; Loh et al., 2016; 2017).

Host specific parasites are typical of Australian monotremes (Kolonin, 2007). In particular, echidnas can carry a number of ectoparasites that include fleas, ticks, and mites (Roberts, 1970; Augee et al., 2006). Australian short-beaked echidnas are documented as parasitised by two dominant tick genera: Bothriocroton and Amblyomma (Roberts, 1970; Booth, 1994). There are six confirmed species of Bothriocroton in Australia, with the native tick species B. concolor and Bothriocroton tachyglossi commonly found on wild and captive echidnas (Barker & Walker, 2014). Bothriocroton undatum and Bothriocroton hydrosauri have also been reported on echidnas, with B. hydrosauri, the southern reptile tick, the only species of Bothriocroton documented to bite humans (Roberts, 1970; Booth, 1994; Barker & Walker, 2014). Other tick species recorded on echidnas include Amblyomma australiense, Amblyomma echidnae, and Amblyomma moyi with respect to Australian echidna species, whilst Amblyomma papuanum is found on New Guinea echidnas. Furthermore, ticks within the Ixodes genus: Ixodes holocyclus and Ixodes tasmani have also been observed, though these species appear opportunistic on echidna hosts as opposed to host-specific (Roberts, 1970; Booth, 1994; Kolonin, 2007; Gofton et al., 2015a).

1.2 Tick-borne pathogens and the tick microbiome

Ticks depend on vertebrate hosts for their survival and are therefore deemed obligatory parasites. Moreover, as vectors, ticks serve as reservoirs for microorganisms that are dependent on the tick for completion of their own life cycle (Bogtish et al., 2013). The obligate microorganisms that exist within a host organism’s body are commonly known as the microbiome, and refer to a community of symbiotic, commensal, and pathogenic microorganisms (Bäckhed et al., 2005; Turnbaugh et al., 2007). The microbiome of ticks are significant in studies concerning tick-borne pathogens because of the potential for transmission of disease-causing microorganisms to vertebrate hosts (Jongejan & Uilenberg, 2004; Ahantarig et al., 2013). For instance, in 1893 it was revealed that arthropods are capable of acting as biological vectors of a pathogenic organism, when Smith 10 and Kilbourne discovered that the tick Rhipicephalus (Boophilus) anulatus transmitted the agent of Texas cattle fever, the protozoan Babesia bigemina (Parola & Raoult, 2001). This was the first historic demonstration of the ability for ticks to transmit pathogens.

Ticks are a significant threat to the health of humans, companion animals, livestock, and wildlife because of their role in transmitting bacterial, fungal, protozoan, and viral disease-causing pathogens (Parola & Raoult, 2001; Jongejan & Uilenberg, 2004; Fritz, 2009). An important point to understand is that ticks do not transmit disease; instead they transmit pathogens that can cause the clinical manifestations that are defined as a particular disease (Barker & Walker, 2014). Ticks are efficient vectors of numerous pathogens because of their interactions with several different vertebrate hosts during their life cycle. Consequently, ticks are able to acquire a vast array of microorganisms present in the blood of infected hosts and hence, transmit more than one putative pathogen at a time (Jongejan & Uilenberg, 2004; Gern, 2008; Ahantarig et al., 2013).

During feeding, a tick’s salivary glands function as osmoregulators that return most of the fluid and ion content of the blood meal to the host via salivation into the feeding site. Therefore, as salivary glands are the site for most pathogen development within ticks, injection of the saliva into the host makes it a route of pathogen transmission (Ribeiro et al., 1987; Bowman & Sauer, 2004; Francischetti et al., 2009). Other pathogens can be found residing within the midgut, only introduced into the host when the tick regurgitates digestive enzymes (Sonenshine, 2005). A tick is only considered a competent vector for a specific pathogen if it: 1) feeds on an infected vertebrate host, 2) can acquire the pathogen during feeding, 3) is able to maintain the pathogen throughout its life cycle which is known as trans-stadial or ‘vertical’ transmission, and 4) has the ability to transfer the pathogen to another host when feeding, termed ‘horizontal’ transmission (Kahl et al., 2002; Nicholson et al., 2009). Another form of transmission can occur when an infected female tick lays eggs. Here, if a pathogen is capable of transferring into the eggs and survives in a subsequent generation, it is coined transovarial transmission, also considered a form of ‘vertical’ transmission (Burgdorfer & Varma, 1967; Sonenshine & Roe, 2013). The

11 differences between trans-stadial and transovarial transmission routes are depicted in Figure 1.4.

Figure 1.4. Transmission cycles of a tick. Example is the three-host tick I. holocyclus (paralysis tick), which is a known vector of Rickettsia australis, a bacterium causing Queensland tick typhus. Host records for I. holocyclus in Roberts (1970) include the short-beaked echidna (T. aculeatus), northern brown bandicoot (Isoodon macrourus), red-necked wallaby (Macropus rufogriseus), and humans (Homo sapiens). As humans are unable to propagate the bacterium in nature they are considered a dead-end host (Moulder, 1989). The question mark associated with individual hosts indicates the uncertainty surrounding the specific host for each tick life stage shown, and should therefore be regarded with caution.

1.2.1 Tick-borne diseases (TBDs) – a global perspective

The number of recognised infectious diseases has increased over the decades, with the emergence of more than 300 human infectious diseases since the 1940s (Daszak et al., 2008). Around 60% of these diseases are zoonotic (transferred between animals and humans), of which 72% are believed transmissible from wildlife (Daszak et al., 2008; Institute of Medicine, 2011).

12 Ticks are thought to surpass all other arthropods in the variety of infectious agents they transmit (Sonenshine & Roe, 2013). Concerning bacterial agents, ticks have been linked with the transmission of a diverse array of infectious bacteria, with those of significant concern belonging to the genera Anaplasma, Borrelia, Coxiella, Ehrlichia, Francisella, Neoehrlichia, and Rickettsia, all of which contain recognised pathogens of humans and animals (Munderloh et al., 2005; Ahantarig et al., 2013).

Approximately 30% of emerging infectious diseases are vector-borne which include tick-borne diseases (TBDs) (Institute of Medicine, 2011). Presently, there are in excess of 16 TBDs of humans and more than 19 TBDs of companion animals, livestock, and wildlife recognised worldwide (Nicholson et al., 2009). TBDs have been specifically attributed to various human and animal activities that disrupt ecosystems. These include increases in human populations and shifts in geographical distribution of humans, wildlife, vectors, and pathogens throughout the world (Parola & Raoult, 2001; Sonenshine & Roe, 2013). Such alterations have generated more opportunities for humans and wildlife to interact at more interfaces, thus increasing the prospect for the transmission of TBDs and emergence of novel infectious diseases worldwide (Parola & Raoult, 2001; Jongejan & Uilenberg, 2004; Grogan et al., 2014). However, despite this understanding there is currently incomplete knowledge regarding TBDs in wildlife (Taylor et al., 2001), a major concern considering wildlife and their tick ectoparasites act as reservoirs and potential amplifiers of pathogenic microorganisms (Cooper et al., 2013; Tozer et al., 2014).

1.2.2 TBDs in Australia

Three prevalent bacterial zoonotic TBDs are recognised in Australia and have been linked to native tick species. The two main TBDs associated with tick bites are Queensland tick typhus and Flinders Island Spotted Fever, which are correspondingly caused by the bacterial agents Rickettsia australis and Rickettsia honei, and are associated with I. holocyclus (Australian paralysis tick), the marsupial tick I. tasmani, and the southern reptile tick B. hydrosauri (Campbell & Domrow, 1974; Sexton et al., 1991; Stenos et al., 2003; Graves & Stenos, 2009). The third TBD, although not typically associated with tick bites, is Query (Q) fever,

13 discovered in Australia by Derrick in 1935 (Smith & Derrick, 1940). Q fever is almost ubiquitous in global distribution and is caused by the pathogen Coxiella burnetii, which is found in many species of wildlife (Smith & Derrick, 1940; Cutler et al., 2007; Cooper et al., 2013; Tozer et al., 2014). In Australia, C. burnetii is vectored by Haemaphysalis humerosa and Amblyomma triguttatum, the bandicoot tick and ornate kangaroo tick respectively (Smith & Derrick, 1940; McDiarmid et al., 2000; Bennett et al., 2011).

Protozoa and other bacterial pathogens are also the cause of TBDs in Australian wildlife. TBDs caused by haemoprotozoan pathogens: Babesia spp., Theileria spp., and Trypanosoma spp., include babesiosis (Jefferies et al., 2003), theileriosis (Islam et al., 2011; Hammer et al., 2015), and multiple clinical manifestations for Trypanosoma spp. (Austen et al., 2011; McInnes et al., 2011; Thompson et al., 2014; Barbosa et al., 2016). Tick-borne bacterial pathogens include Rickettsia spp. and Bartonella spp., and are established as correspondingly causing fever and malaise, and cat scratch disease in Australian wildlife (Breitschwerdt & Kordick, 2000). Despite wildlife being reservoirs for ticks and infectious agents, the significance of wildlife ticks concerning putative pathogens is presently overlooked, and although ticks are generally well studied because of their impact on human health, novel microbial associations remain to be described.

1.2.3 The genus Borrelia

Borrelia is a group of helical-shaped, motile, obligate arthropod-borne parasites of vertebrates within the phylum Spirochaetes and family Spirochaetaceae (Barbour & Hayes, 1986; Gupta et al., 2013; Burgdorfer & Rosa, 2015). As a spirochaete, Borrelia is considered a large bacterium with a length between 10-30 micrometres (μm) and width less than 1 μm. Moreover, Borrelia spirochaetes consist of a flat, wavy shaped body and flagella that assists in locomotion, enabling the motile bacteria to efficiently move through blood and tissue (Barbour & Hayes, 1986; Burgdorfer & Rosa, 2015). Borrelia spirochaetes reproduce by binary fission (Fritzsche, 2005), and with an obligate parasitic lifestyle, are unable to survive outside a host, whether tick or vertebrate (Parola & Raoult, 2001).

14 The genus Borrelia consists of approximately 37 recognised species (Gupta et al., 2013; Burgdorfer & Rosa, 2015) that traditionally comprise two major lineages: Borrelia burgdorferi sensu lato (B. burgdorferi s.l.) and the relapsing fever borreliae (Geller et al., 2012; Wu et al., 2013; Burgdorfer & Rosa, 2015). The B. burgdorferi s.l. complex contains several species that are causative agents of Lyme borreliosis (LB), or more commonly Lyme disease. A TBD found in the temperate regions of the Northern Hemisphere, LB is conventionally viewed as transmitted by hard ticks of the genus Ixodes (Anderson & Magnarelli, 1993b; Stanek et al., 2012). The LB group is considered a heterogenous bacterial complex, currently comprising more than 20 species of which six are currently recognised as pathogenic: Borrelia burgdorferi senso stricto (s.s), Borrelia afzelii, Borrelia garinii, Borrelia bavariensis, Borrelia spielmanii, and more recently a novel species called Borrelia mayonii (Jungnick et al., 2015; Pritt et al., 2016). The major human pathogenic borreliae are associated with various clinical multi-organ manifestations of LB, which include dermatologic, neurologic, and rheumatologic symptoms (Tilly et al., 2008; Samuels & Radolf, 2010; Halperin, 2011a; b; Stanek et al., 2012). LB typically presents as a chronic disease and therefore has associated health and economic burdens (Fingerle et al., 2008). Furthermore, since it’s official 1970s recognition in Northern America (Scrimenti, 1970; Steere et al., 1977; Steere et al., 1978), LB has become one of the most commonly diagnosed TBDs in the Northern Hemisphere (Institute of Medicine, 2011) with approximately 86,000 people acquiring a LB infection annually, and an estimated 77% of LB incidences reported in Europe, 19% in North America, 4% in Asia, and around 1% in northern parts of Africa (Hubálek, 2009).

The second group, the tick-borne relapsing fever (RF) borreliae, typically cause disease characterised by recurring episodes of fever (Barbour, 2014), and includes several species: Borrelia hermsii, Borrelia turicatae, Borrelia parkeri, Borrelia duttonii, Borrelia crocidurae, Borrelia anserina which causes avian spirochaetosis, and Borrelia theileri and Borrelia coriaceae which are the agents of bovine borreliosis (McNeil et al., 1949; LeFebvre & Perng, 1989). Another RF inclusion is Borrelia miyamotoi, which was discovered in Asia in 1995 (Fukunaga et al., 1995) and only in 2011 gained recognition in Russia as a human pathogen (Platonov et al., 2011). RF borreliae are established in North and South America, Europe, Africa,

15 and Asia, and are commonly louse and soft tick-borne; mainly soft ticks of the genus Ornithodoros (Cutler, 2015), with the exception of B. miyamotoi and B. theileri, which have been found in hard tick species (Scoles et al., 2001; Fraenkel et al., 2002; Bunikis et al., 2004; McCoy et al., 2014).

In 2003, the discovery of a novel Borrelia sp. in Turkey that was isolated from the hard tick, Hyalomma aegyptium, found parasitising a tortoise (Güner et al., 2003), has led to the support of a third Borrelia lineage named the reptile-associated (REP) Borrelia group (Takano et al., 2010). Later termed Borrelia turcica (Güner et al., 2003; Güner et al., 2004), this species as well as others that fall within the REP Borrelia phylogenetic group are genetically distinct from spirochaetes of the LB and RF groups, and appear to be associated with reptiles and their hard-bodied ectoparasites of the genera Amblyomma and Hyalomma (Takano et al., 2010). Furthermore, the discovery of a novel Borrelia sp. in the native echidna-biting tick, B. concolor, and which forms a unique cluster in the genus, appears to comprise of a fourth major Borrelia group, potentially associated with Australian monotremes (Gofton et al., 2015a; Loh et al., 2016).

1.2.4 Borrelia in Australia

The confirmation of LB in the United States in the late 20th century stimulated a renaissance in tick-borne pathogen research around the world, in particular regarding Borrelia spp. and their potential pathogenicity (Bowman & Nuttall, 2008). Australia has not been exempt from the search for these elusive organisms and Figure 1.5 illustrates the timeline entailing significant Borrelia-related studies.

Figure 1.5. The timeline of Borrelia-related discoveries and reported observations in Australia. Information compiled from Mulhearn (1946), Mackerras (1959), Carley and Pope (1962), Rothwell et al. (1989), Wills and Barry (1991), Russell et al. (1994), Petney et al. (2004), Gofton et al. (2015a), and Loh et al. (2016). Dates with a red ‘X’ symbol mark the studies that contain results which are either inconclusive for Borrelia spirochaetes, the species is undetermined due to an inability to replicate the investigation or all Borrelia isolates have been lost, contain unsubstantiated claims, or conclude the absence of Borrelia spirochaetes.

1.2.4.1 Borrelia in Australian animals

Two species of Borrelia, B. theileri and B. anserina, the global causative agents of bovine borreliosis and avian spirochaetosis respectively (Sharma et al., 2000; Ataliba et al., 2007), were introduced to Australia through agricultural routes in the 1900s (Mulhearn, 1946). Borrelia theileri is transmitted by the cattle tick R. (B.) australis, which has a geographical distribution along the northern and eastern coasts of Australia, and has been documented in cattle from the Australian states of Queensland and New South Wales (Mulhearn, 1946; Callow & Hoyte, 1961; Seddon & Albiston, 1967; Barker & Walker, 2014). Borrelia anserina, vectored by the soft tick Argas persicus sensu stricto (Petney et al., 2004), is observed throughout mainland Australia and has previously infected poultry originating from Victoria and the Northern Territory (Gorrie, 1950; Seddon & Albiston, 1967; Beveridge &

17 Hart, 1985; Swayne, 2013). Although R. (B.) australis is known to bite humans, there have been no reported cases of Borrelia-related infections from either B. theileri or B. anserina in humans (Barker & Walker, 2014).

In 1989, LB was claimed to be the cause of fever, anaemia, polyarthritis, and overall poor condition in two cows from New South Wales, diagnosed from the apparent presence of spirochaetes in synovial stroma cells, though a formal confirmation was never made (Rothwell et al., 1989). Additionally, as the possibility of these cases resulting from a B. theileri infection was not ruled out, the evidence in favour of LB in Australian cattle remains unsubstantiated and controversial to say the least.

Queensland has played host to numerous reporting’s of Borrelia spp. other than cattle-related (Mulhearn, 1946). These reports have concerned native animals such as bandicoots and kangaroos, with spirochaetes observed in blood films collected from both Australian natives (Mackerras, 1959). Although concluded as a novel Borrelia, the spirochaete’s phylogenetic place has remained unspecified in the Borrelia genus due to the limitations of molecular profiling technologies at the time (Mackerras, 1959). Similarly, in response to reports that the native rat, Rattus villosissimus, was succumbing to a mysterious infectious disease in north-western Queensland, Pope and Carley in 1956, autopsied 27 R. villosissimus specimens in an attempt to isolate the infectious agent (Pope & Carley, 1956). A novel spirochaete species was observed from one autopsied rat and subsequently named Borrelia queenslandica (Carley & Pope, 1962). Carley and Pope conducted additional studies in 1957 and 1958 to further examine this spirochaete’s potential pathogenicity, however attempts to inoculate a human volunteer, as well as prove a transmission link between mice via the soft tick Ornithodorus gurneyi, were both unsuccessful (Carley & Pope, 1957; 1958). Moreover, with the loss of all isolates from the studies of Pope and Carley (1956), no conclusion can be made regarding the phylogeny of B. queenslandica in relation to other Borrelia spp.

18 1.2.4.2 Borrelia in Australian ticks

Several studies to date have been conducted to investigate the presence of Borrelia, in particular the LB agent, B. burgdorferi s.l., in Australian ticks. The first study was piloted by Wills and Barry (1991), who at the time amassed a total of 167 I. holocyclus and H. longicornis ticks from the coastal regions of New South Wales. The ticks were dissected and their midguts cultured, with motile, rigid spirochaete-like objects (SLOs) observed in 44% and 35% of I. holocyclus and H. longicornis cultures respectively. Further investigations using various identification techniques were performed which were indicative of Borrelia spp., though neither the laboratory methods utilised in the investigations nor the organisms recovered were made available for reproduction, hence rendering Wills and Barry’s results as inconclusive. Additionally, cultures of another bacterium, Bacillus, have been identified as able to produce similar SLOs (Brem et al., 1999), adding further doubt to their claimed findings. Wills and Barry’s study was later challenged by Russell et al. (1994), who collected nearly 12,000 ticks from the coast of New South Wales, derived from natural habitats and native and domesticated animal hosts. The three main tick species were I. holocyclus, H. longicornis, and Haemaphysalis bancrofti. The midguts of these ticks were cultured and screened using microscopy, with 92 cultures showing SLOs, later determined as contaminant flagella aggregates. The study by Russell et al. (1994) had two main advantages over that of Wills and Barry. Firstly, Russell and colleagues had a larger sample size that was more representative of the tick population. Secondly, with the incorporation of Borrelia genus-specific primers, they were able to justify the likely absence of LB spirochaetes in Australian ticks. Finally, Russell and colleagues observed the cultured SLOs from the 1991 investigations by Wills and Barry and deduced that both studies had cultured identical SLOs, contradicting the findings of Wills and Barry and supporting the absence of LB spirochaetes formulated by Russell et al. (1994).

Prompted by the finding of a novel Borrelia sp. within a single I. holocyclus tick that had been removed from an echidna (Gofton et al., 2015a), Loh et al. (2016) investigated the presence of Borrelia in the native echidna-biting tick, B. concolor. A total of 97 B. concolor ticks were removed from 22 echidnas of the species T.

19 aculeatus, which originated from Queensland, New South Wales, and Victoria. Using Borrelia genus-specific Polymerase Chain Reaction (PCR) assays targeting the ubiquitous 16S rRNA (16S) and flaB (flagella encoding protein) genes, Loh and colleagues revealed that 38 of the 97 B. concolor ticks from Queensland and New South Wales contained the novel Borrelia DNA, confirming the observation by Gofton et al. (2015a). Furthermore, although genetically related to the REP group, Loh and colleagues concluded that the novel Borrelia sp., recently termed ‘Candidatus B. tachyglossi’ (Loh et al., 2017), was distinct, proposing a fourth unique clade within the Borrelia genus (Loh et al., 2016). Though the zoonotic potential and pathogenicity of ‘Candidatus B. tachyglossi’ remains unknown, the investigations initiated by Gofton et al. (2015a; b) and Loh et al. (2016; 2017) are ground breaking as they revealed for the first time the presence of Borrelia in native Australian ticks. Nonetheless, not yet explored is the presence of ‘Candidatus B. tachyglossi’ in other echidna-biting ticks, or whether it has an association with the Australian echidna, a potential host reservoir for the bacterium (Loh et al., 2016; 2017). In addition, the past presence of ‘Candidatus B. tachyglossi’ in Australian ticks and its occurrence over a greater geographical region are currently unknown. Further investigation using a larger and more diverse sample of museum-archived echidna-biting ticks can contribute to answering these fundamental questions.

1.3 Detection methods of tick-borne pathogens and the tick microbiome

Significant advances have been made regarding the detection and identification of microorganisms and pathogens in the world, with more than 50 emerging infectious agents recognised in the last 40 years (Dong et al., 2008), and almost 10% identified as bacteria (Woolhouse & Gaunt, 2009). This technological evolution over the decades can be attributed to the introduction of new tools enabling sequencing of genetic components and profiling characteristics at the molecular level (Houpikian & Raoult, 2002). These include PCR analysis (Mullis & Faloona, 1987), Sanger sequencing (Sanger et al., 1977), and NGS technologies (Anderson & Schrijver, 2010).

20 1.3.1 Polymerase Chain Reaction (PCR)

The development of the Polymerase Chain Reaction (PCR) in the late 1980s led to the rapid expansion in studies concerning microorganism detection. Whilst early studies investigating pathogens have often relied on traditional techniques including microscopy, serology, and cloning and culture (Houpikian & Raoult, 2002; Kahl et al., 2002), since the invention of the PCR, this technology has become one of the most commonly utilised molecular techniques to amplify sequences of nucleic acids (Li et al., 2000; Padmanabhan et al., 2013). The PCR is a highly sensitive technology when compared to conventional serology (Fournier et al., 1998), and has been incorporated in studies as a means of expanding the ability to detect difficult to culture and low quantity organisms such as intracellular bacteria (Houpikian & Raoult, 2002). The sensitivity of the PCR has enabled data to be obtained from small quantities of DNA and within a reduced timeframe compared to traditional methods (Hopwood, 1999; Glick & Pasternak, 2003). Moreover, this sensitivity has detected organisms of dormant or latent diseases, such as LB (Halperin, 2011a), and identified diseases in which the causative organism’s quantity in culture from tissue or body fluid is low (Fredericks & Relman, 1996).

The PCR can assist in establishing geographical distribution. For instance, B. burgdorferi s.l. was thought to only occur in Northern Hemisphere countries and their respective ticks of the Ixodes ricinus complex. However, in 2013 B. burgdorferi s.l. DNA was reported in ticks inhabiting Uruguay, South America by Barbieri et al. (2013). Moreover, though live Borrelia spirochaetes were unable to be cultivated in the study, Borrelia was detected using the sensitivity of the PCR (Barbieri et al., 2013). Similarly, in 2014 Schramm and colleagues detected B. burgdorferi s.l. DNA in the blood of king penguins, infested with Ixodes uriae and breeding in the Southern Indian Ocean, using serology and in vitro DNA amplification (Schramm et al., 2014). Interestingly, of the five penguins positive by serology, one of these birds was tick-free. This study indicates a possible reservoir role for penguins in the natural maintenance of this bacterium and suggests an association between I. uriae and the Borrelia found in king penguins. Furthermore, this study eludicates a potential association between other Borrelia spp., ticks, and their wildlife hosts.

21 The PCR is a well-utilised approach for direct detection of Borrelia infection in both tick vectors and reservoir hosts. To determine the species of Borrelia, a qualitative or conventional PCR analysis approach is used (Loh et al., 2016), whereas to quantify Borrelia DNA, a real-time or quantitative PCR (qPCR) method is necessary (Piesman et al., 2001; Jenkins et al., 2012). Both methodologies involve targeting genes such as flaB, OspA and OspC (encode for outer surface proteins A and C), and 16S genes for amplification (Persing et al., 1990; Johnson et al., 1992; Fingerle et al., 1998; Wang et al., 1999; Wang et al., 2003). The 16S gene is universal in distribution and also ubiquitous among microorganisms of the bacterial domain, with nine hyper-variable regions (V1-V9) separated by highly conserved regions, allowing for primer design and amplification of multiple bacterial taxa (Schabereiter-Gurtner et al., 2003; Clarridge, 2004; Radulović et al., 2010; Schloss, 2010). Mutations within the 16S gene have facilitated the study of microbial communities (Sinclair et al., 2015) and bacterial evolution and genetic divergence (Woese, 1987), with amplified sequences compared with reference- sequence databases to infer phylogenetic relationships (Mizrahi-Man et al., 2013; Di et al., 2014).

1.3.2 Sanger and next-generation sequencing (NGS) – revolutionary methods of DNA detection

Dideoxy Sanger sequencing, known as Sanger or first-generation sequencing, works by utilising the enzyme DNA polymerase by dideoxy chain termination (Sanger et al., 1977), and has previously been the predominant method for sequencing and characterising bacterial communities using 16S gene clone libraries (Tewari et al., 2011). A Sanger sequencing run involves only one sequence at a time and up to 1,000 base pairs (bp), though by merging overlapping sequences longer sequences can be obtained. Furthermore, Sanger sequencing has a small amount of DNA read with each sequencing reaction at 80-100 kilobases (Kb) per hour (Morey et al., 2013), due in part to template preparation and time to carry out the enzymatic reaction. However, Sanger sequencing has been optimised to reduce run times, now incorporating automation and fluorescently labelled dideoxy-nucleotides that act as chain terminators (Atkinson et al., 1969; Smith et al., 1986). The benefits from Sanger sequencing have been complemented with the

22 continuous advancements in sequencing technologies since the year 2000, and although Sanger sequencing offers longer sequences and is routinely used, many contemporary studies have incorporated the more efficient second-generation sequencing technologies, termed next-generation sequencing (NGS) platforms (Morozova & Marra, 2008; Metzker, 2010; van Dijk et al., 2014).

NGS is a type of DNA sequencing technology that processes multiple small fragments of DNA in a parallel fashion (Rizzo & Buck, 2012). There are two main types of NGS: 1) de novo or shot-gun sequencing, which is used for whole genome sequencing, and 2) amplicon or metabarcoding, which is a targeted approach where unique indices are incorporated into amplicons, allowing multiple samples to be pooled together and sequenced simultaneously. The indices are used for subsequent sample identification (Schatz et al., 2010; Pareek et al., 2011; Fantini et al., 2015). Compared to Sanger sequencing, NGS results in more DNA sequenced at a reduced timeframe and overall cost (Anderson & Schrijver, 2010; Mardis, 2011; 2013; Niedringhaus et al., 2011). Hence, the primary advantage of NGS platforms is the significantly larger amount of DNA read with each sequencing reaction, termed a high-throughput (Morey et al., 2013). For instance, the most commonly used platform for 16S rRNA metabarcoding is the Illumina MiSeq (Claesson et al., 2010; Burke & Darling, 2016), where the 16S gene is amplified by PCR and tagged with unique indices for amplicon library pooling. With the library loaded onto the machine, the amplicons are clonally amplified on the flowcell ranging from 1-30 million sequences. The unique indices on each sequence allow assignment back to an individual sample (Caporaso et al., 2012; Vences et al., 2016). Data is then analysed using various bioinformatic pipelines such as Geneious, USEARCH, and QIIME (Caporaso et al., 2010; Edgar, 2010; Kearse et al., 2012). Consequently, the ability to massively sequence in a parallel manner has enabled shorter regions of the 16S gene to be sequenced at a greater depth and significantly lower cost (Tringe & Hugenholtz, 2008; Burke & Darling, 2016). This has permitted studies involving pathogen detection and sequencing the tick bacterial microbiome to be performed (Schopman et al., 2012; Gofton et al., 2015a; b), with results revealing high levels of bacterial diversity in individual ticks and specific tick species (Clay et al., 2008; Heise et al., 2010). Therefore, as it is now viable to study complete microbial communities, the need for isolating and culturing individual

23 microorganisms as well as prior knowledge of their genetic sequences are no longer a requirement (Capobianchi et al., 2013).

NGS is not without its pitfalls. Currently, NGS only sequences short DNA fragments and when unable to resolve taxonomy or phylogeny, PCR of longer amplicons with Sanger sequencing is often used to confirm NGS findings (Vogl et al., 2012; Morey et al., 2013). For example, Gofton et al. (2015a) detected a novel Borrelia sp. in a single I. holocyclus tick using NGS, and tentatively determined it was closer to the RF borreliae clade. However a subsequent investigation by Loh et al. (2016), using PCR assays targeting the flaB and longer 16S genes followed by Sanger sequencing, revealed the novel Borrelia sp. as more closely related to the REP Borrelia group, but forming a new fourth clade in the Borrelia genus. Consequently, to overcome the shortcomings of each sequencing technology, the incorporation of multiple technologies is often adopted to increase sequence length, limit errors, and reduce cost and laborious effort (Vayssier-Taussat et al., 2013; Gofton et al., 2015a; b; Loh et al., 2016).

1.4 Museum specimens – an unexplored ‘gold mine’

Globally, natural history collections and museums house an expansive array of dried and spirit-preserved specimens of a variety of taxa including current, endangered, or extinct species, as well as examples of pathological conditions (Barnes et al., 2000; Burrell et al., 2015). These biological archives have enabled evolutionary processes in populations to be measured over time (Mende & Hundsdoerfer, 2013), provided an insight into genetic variation among past and present individuals of a particular species (Besnard et al., 2016), allowed the presence of pathogens in past populations to be examined (Persing et al., 1990; Marshall et al., 1994; Matuschka et al., 1996; Hubbard et al., 1998; Movila et al., 2013), and facilitated an understanding of the effects pathogens can have on populations (Wyatt et al., 2008).

Museums are of vital importance for establishing the baseline of an organism’s diversity and past prevalence, though despite accommodating potentially significant sources of genetic data, especially from specimens collected prior to the

24 molecular era (Payne & Sorenson, 2003), these historical collections are currently underutilised. This is due in part to their variable DNA quality and quantity, as well as the destructive nature of traditional DNA sampling (Wandeler et al., 2007). However, the advent of high-throughput sequencing (Mardis, 2013) has revolutionised the ability to source these often low and fragmented quantities of genetic material (Wandeler et al., 2007), opening the doors for novel discoveries to be made through the exploration of these archived specimens.

1.4.1 Ancient DNA (aDNA)

Ancient DNA (aDNA) is defined broadly as the retrieval of nucleic acid sequences from biological archives such as museums, as well as archaeological finds and fossil remains (Pääbo et al., 2004; Druzhkova et al., 2015). Museum specimens have been used in studies concerning evolutionary genetics since the 1980s, when Higuchi and colleagues extracted aDNA from an extinct type of zebra, the Equus quagga quagga, which had been archived in a museum for more than 150 years (Higuchi et al., 1984). A year later, the retrieval of aDNA from mummified human specimens dating back thousands of years was achieved by Pääbo (1985), which paved the way for further use of museum specimens to investigate taxonomy, resolve phylogenetic relationships (Schaffner et al., 1989; Cooper et al., 1992), deduce past geographical ranges, and compare biodiversity over time (Thomas et al., 1990; Nielsen et al., 1997; Bouzat et al., 1998).

Initially, aDNA studies were few and far between due to the complexity and limited amplification success of aDNA using cloning techniques in vivo. However, the concurrent arrival of the PCR and Sanger sequencing has enabled DNA amplification to take place in vitro from much smaller quantities including shorter fragments of DNA. This has delivered an efficient and normally reproducible method of sequencing ancient genetic markers (Pääbo, 1989) that has ultimately led to an expansion in the aDNA field (Druzhkova et al., 2015). Though both technologies have diversified aDNA studies, problems still exist regarding the fragmented quality (Bär et al., 1988), low quantity, and therefore limited and potentially erroneous amplification of DNA retrieved from museum specimens (Wandeler et al., 2007). This is a particular issue for specimens that have either

25 been poorly preserved (Hall et al., 1997; Barnes et al., 2000; Zimmermann et al., 2008) or were collected decades to centuries ago, with extreme fragmentation of DNA limiting the type and number of amplifiable sequences using this method (Hofreiter et al., 2001; Pääbo et al., 2004; Wandeler et al., 2007).

Analytical challenges involving the interpretation of aDNA are primarily caused by the typical physical and chemical damage sustained following cell death, which increases over the course of time. Due to this oxidative damage, the expected sequence length for aDNA is at most 500 bp, meaning that larger sequences yielding more informative data cannot be obtained. Additionally, only multicopy genes, such as mitochondrial or ribosomal genes, tend to survive which limits the type of sequence that can be retrieved from ancient material (Hofreiter et al., 2001). This can affect the phylogenetic interpretation of aDNA data as some genes, depending on their level of sequence divergence, can suggest relationships on different ordinal levels (Dittmar et al., 2006). Therefore, in order to avoid misrepresentation and to construct a more meaningful phylogenetic context, more than one gene should be used. By using different genes in a combined analysis, more genetic information becomes available which optimises the phylogenetic explanatory power (Wiens, 2003; Dittmar et al., 2006).

Finally, when amplifying aDNA, the potential for contamination with modern or exogenous DNA is common (Pääbo, 1989; Cooper & Poinar, 2000; Druzhkova et al., 2015). For instance, if modern DNA is co-extracted with the fragmented aDNA from a museum specimen, the modern DNA may be amplified preferentially during the PCR (Pääbo, 1989). This was found to be the case with the apparent retrieval of aDNA from mummified humans by Pääbo et al. (1985) whose results were later dismissed due to contamination from modern DNA (Pääbo, & Wilson, 1988; Pozzo, & Guardiola, 1989; Knapp et al., 2015). Hence, contamination has remained a constant concern with aDNA research. Specific criteria to limit the possibility of modern DNA contamination in aDNA samples have been extensively proposed and reviewed numerously by researchers including Cooper and Poinar (2000), Hofreiter et al. (2001), Pääbo et al. (2004), and Gilbert et al. (2005), and is of paramount importance to any aDNA investigation.

26 1.4.1.1 aDNA and NGS – a ‘perfect’ pair

A new era in the study of aDNA began with the advent NGS which has allowed for two main advancements in molecular profiling: 1) billions of base pairs can be analysed simultaneously, and with metabarcoding and library capture, desired genomic sequences can be resolved (Druzhkova et al., 2015), and 2) there is now an ability to explore the aDNA of non-traditional substrates, such as eggshell (Allentoft et al., 2009). The limitation of DNA fragmentation has been mitigated with the features of NGS. For instance, NGS platforms require the DNA template to be short, which happens to be a main characteristic of the usually highly fragmented nature of aDNA (Mardis, 2013; Linderholm, 2016).

Despite the many benefits of using NGS for aDNA analysis, computational challenges have emerged. NGS generates billions of sequences from both species of interest and non-target species, resulting in a massive output of genetic data. The main issue is that to find the target species, one must first conduct a similarity search using the genome of a closely related species as well as explore a large database of sequences (Prüfer et al., 2010). With the typically low amount of endogenous DNA in aDNA studies, sequencing success is highly dependent on how well the endogenous sequences can be identified, and hence on the availability of a comparative genomic sequence, which may be currently absent for novel microorganisms. Further aDNA concerns include DNA base damage and fragmentation, which have the potential to result in the incorporation of wrong bases during amplification, which may be difficult to discern (Pääbo et al., 2004). Therefore, a close genomic reference sequence is crucial when validating aDNA sequences (Dittmar et al., 2006) and operating sequence alignment software such as Geneious (Kearse et al., 2012). Finally, sequencing the 16S gene permits the relative proportions of different microbial populations within complex communities to be determined. However, a major limitation of bacterial community analysis is that it can exaggerate groups of microorganisms by favouring the organisms with a high gene copy number and hence generate a skewed bacterial composition (Větrovský & Baldrian, 2013; Gofton et al., 2015a). Nevertheless, what is not normally exaggerated is the diversity of microorganisms within a community. One way to minimise these obvious aDNA and NGS pitfalls is

27 to analyse different genes using more specific sequencing technologies such as the PCR and Sanger sequencing, which when combined with NGS analysis, will provide the greatest possible explanatory power when resolving relationships in a phylogenetic context (Dittmar et al., 2006).

1.4.2 Using museum specimens to investigate pathogens

A unique branch in aDNA research involves the study of microbial pathogenic DNA within the bodies of infected host specimens (Donoghue & Spigelman, 2006). The first major study of pathogens in ancient remains began with Spigelman and Lemma who isolated Mycobacterium tuberculosis from 14-16th century skeletal remains (Spigelman & Lemma, 1993). Since then, molecular studies of archived specimens has provided an invaluable tool for investigating the evolution and past occurrence of specific pathogens (Raoult et al., 2000; Tsangaras & Greenwood, 2012; Burrell et al., 2015), including an insight into their potential host or environmental reservoir (Donoghue, & Spigelman, 2006). For example, in 1994 Marshall and colleagues obtained museum specimens of Peromyscus leucopus, the white-footed mouse, which were originally collected in the United States between the period of 1870 and 1919 (Marshall et al., 1994). In this study, Marshall and colleagues investigated whether the microbial pathogen B. burgdorferi was enzoonotic within the United States at the onset of the 20th century. Although only two of the 280 specimens analysed were positive for B. burgdorferi, their findings were significant as it suggested P. leucopus as a potential reservoir host for the bacterial pathogen (Marshall et al., 1994). Similarly, a previous study in 1990 by Persing and colleagues also identified B. burgdorferi, though in archived Ixodes dammini ticks (Persing et al., 1990; see Section 1.4.2.1). Accordingly, B. burgdorferi, the causative agent of LB, became one of the first tick-borne bacteria to be isolated from museum specimens.

1.4.2.1 Using museum specimens to investigate tick-borne pathogens

Modern tick specimens are a conventional source in studies concerning vectored putative pathogens, however they provide only indirect evidence of historical processes and a limited temporal insight into a pathogens emergence over time

28 (Besnard et al., 2016). Testing archived material sourced from museum and natural history collections can clarify the history of infectious agents (Hofreiter et al., 2001) and provide an avenue to explore past occurrence (Besnard et al., 2016).

While in the early 1980s, Burgdorfer was first to discover Borrelia DNA in the gut content of modern dissected ticks (Burgdorfer et al., 1982; Burgdorfer, 1984), subsequent research by Persing et al. (1990) determined the presence of this Borrelia DNA, B. burgdorferi, in archived tick specimens. Persing and colleagues examined 136 archived ticks originating from various geographical locations throughout the United States, and discovered that 13 specimens, derived from the 1940s, contained B. burgdorferi DNA with sequences similar to modern day isolates of the bacteria. This study by Persing et al. (1990) was monumental as it depicted the appearance of B. burgdorferi, the aetiological agent of LB, in ticks as preceding the formal 1970s recognition of the disease (Steere et al., 1977). Further studies into tick-borne pathogens pursued with the use of the PCR in the 1990s, including the identification of B. burgdorferi and B. garinii in archived European ticks of the species Ixodes ricinus by Matuschka and colleagues. This study concluded the presence of Borrelia spirochaetes in ticks dating back to 1884 (Matuschka et al., 1996). A subsequent study by Hubbard et al. (1998) investigated the distribution of B. burgdorferi in 1896 to 1994 archived ticks of eight species and various life stages from the European countries of England, Scotland, and Wales. Similarly, Hubbard and colleagues incorporated the sensitivity of the PCR to detect B. burgdorferi DNA, finding that all species contained the bacterium, including for the first time the presence of B. burgdorferi in the bat soft tick, Argas vespertilionis (Hubbard et al., 1998). Furthermore, by using archived material, studies by Matuschka and Hubbard have suggested an extensive range of host-tick relationships involving the Borrelia bacterium in early 20th century Europe (Matuschka et al., 1996; Hubbard et al., 1998).

More recently, Movila et al. (2013) investigated the prevalence of tick-borne pathogens in I. ricinus ticks archived from the eastern-European country Moldova, in the year 1960. The purpose of their study was to determine the occurrence of bacterial and protozoan agents in museum-archived ticks so as to explore the diversity of tick-borne microorganisms in recent Moldovan history. Through the

29 utilisation of pathogen-specific PCR screening, five different Borrelia spp. were discovered in 21 of the 126 ticks examined, with B. burgdorferi the most prevalent species identified. Rickettsia, Anaplasma, Neoehrlichia, and Babesia spp. were also detected, supporting the evidence for the occurrence of these putative pathogens in Moldova since at least the year 1960 (Movila et al., 2013).

Although PCR with Sanger sequencing has proved effective in detecting putative tick-borne pathogens harboured within archived specimens, it is the introduction of NGS coupled with PCR amplification that has facilitated profiling of entire bacterial communities in modern tick specimens (Gofton et al., 2015a). Nevertheless, bacterial profiling of archived ticks using the advanced features of NGS has yet to be performed. This would permit the exploration of historic and novel microbial associations, and could further assist in resolving an unknown bacteraemic reservoir host. In addition, by comparing microbiomes over time, further information concerning tick biology and how pathogens are transmitted may be revealed. Therefore, profiling archived ticks constitutes a significant knowledge gap in the investigation of past and emerging tick-borne pathogens.

1.4.3 Non-destructive sampling – a method for historical preservation

Ancient and archived biological specimens are valuable resources for the study of historical processes (Payne & Sorenson, 2003; Staats et al., 2013; Besnard et al., 2016). However, a major constraint exists other than the fragility and irreplaceability of such specimens when attempting to utilise archived material in aDNA research, which is the general destructive nature of the sampling procedure (Whitfield & Cameron, 1994; Mandrioli, 2008). Traditional DNA extractions require the grinding of part or whole specimen for successful nucleic acid retrieval. This destructive method is an issue when dealing with small museum specimens such as insects and other invertebrates, where even minor sampling destroys significant morphological characteristics (Whitfield & Cameron, 1994; Watts et al., 2007). Ideal sampling therefore would not require destruction of any material, but rather maintain the specimen’s original morphology for future studies (Hofreiter, 2012). To overcome this problem, a solution is to apply an extraction protocol that uses digestion buffers designed to recover nucleic acids without affecting the

30 morphological condition of the specimen (Gilbert et al., 2007). Such protocols have been developed for non-destructive sampling of archived specimens, which can still retrieve a suitable quantity of host DNA for PCR amplification (Gilbert et al., 2007; Thomsen et al., 2009; Porco et al., 2010; Tin et al., 2014). However, incorporating this extraction method to assess bacterial communities within archived specimens is a novel approach utilised by the following study.

1.5 Research Aims and Hypotheses

The advent of advanced molecular technologies has enabled in vitro amplification and high-throughput in-depth sequencing of the tick microbiome, allowing for the detection of novel microorganisms that can potentially cause disease (Gofton et al., 2015a; b). Nevertheless, incorporating NGS as a method of profiling the bacterial communities within museum-archived ticks to detect novel microorganisms has yet to be conducted, and consequently advocates a meaningful knowledge gap regarding historic existence and prevalence of past putative pathogens. Hence, the 1990 detection of Borrelia in archived ticks (Persing et al., 1990), the 2015 novel Borrelia finding (Gofton et al., 2015a) and subsequent identification of this bacterium in 40% of echidna-biting B. concolor ticks (Loh et al., 2016), and incorporating a non-destructive sampling method (Gilbert et al., 2007), collectively presents an opportunity to explore tick-borne microorganisms and their presence in recent Australian history, whilst ensuring the preservation of these archived specimens for future investigations.

The principal aim of this research project is to detect the novel Borrelia sp., ‘Candidatus B. tachyglossi’, as identified by Loh et al. (2016; 2017), in a larger and more diverse sample of echidna-biting ticks by utilising the available archives of Australian museums and natural history collections. The main purpose for this study is to provide further supporting evidence that ‘Candidatus B. tachyglossi’ is found within ticks that parasitise the echidna (T. aculeatus).

31 The objectives of this project are as follows: 1. Morphologically identify archived echidna-biting ticks from Australian museums and natural history collections using standard morphological keys. 2. Profile the bacterial communities in archived echidna ticks using NGS targeting the V4 hyper-variable region of the 16S rRNA gene, to determine whether Borrelia and other tick-borne bacteria can be retrieved from archived ticks. 3. Detect Borrelia in archived echidna ticks using genus-specific PCR assays that amplify the flaB and 16S rRNA genes, and further resolve phylogenic positioning with ‘Candidatus B. tachyglossi’ and other borreliae.

The hypotheses of this project are: 1. Using a readily available and larger sample size of museum specimens, morphological identification of archived echidna ticks will reveal B. concolor as the predominant echidna tick ectoparasite. 2. Archived ticks harbour a preserved cache of bacterial communities; therefore NGS bacterial profiling will detect Borrelia and other putative pathogens in echidna ticks. 3. Phylogenetic analysis of the flaB and 16S rRNA sequences within archived echidna ticks will be consistent with ‘Candidatus B. tachyglossi’, and support a fourth Borrelia clade. 4. ‘Candidatus B. tachyglossi’ will be found within different Australian tick species that parasitise echidnas, suggesting that echidnas may be a potential reservoir for this novel bacterium.

CHAPTER TWO: MATERIALS AND METHODS

33 2. Overview

The flow diagram in Figure 2.1 represents the methodology used in this study. Collected ticks were first identified using the morphological keys in the Australian tick identification guide (Roberts, 1970). Then, of the ticks identified, a sub-sample was selected based on species and Australian geographical location for further analyses. This included total genomic DNA extraction, which was followed by 16S rRNA (16S) bacterial metagenomic profiling using next-generation sequencing (NGS) to establish the presence of preserved bacterial communities, and more specifically Borrelia, in archived ticks. Finally, to confirm NGS results and further resolve phylogeny, nested Polymerase Chain Reaction (PCR) assays targeting Borrelia genus-specific flagellin (flaB) and 16S rRNA (16S) genes with Sanger sequencing were performed. Phylogenetic analysis concluded this investigation.

Figure 2.1. A methodological overview of the current study.

34 2.1 Tick collection and preparation

Archived tick specimens were sourced from the Australian National Insect Collection (ANIC) and national Australian museums including Museum Victoria, Queensland Museum, South Australian Museum, and the Western Australian Museum between July 2016 and October 2016. The estimated total number of ticks of a specific developmental stage (instar) collected from each Australian state and territory are recorded in Table 2.1. The ticks collected for this study were registered as removed from an echidna (Tachyglossus aculeatus) host, determined by a unique accession number. Each specimen selected for morphological identification was observed individually, and the geographical location according to state or territory and year of collection recorded. Additional notes regarding observed morphological condition, such as engorged or normal, and any signs of damage were also documented in Appendix A, Table A.1.

Table 2.1. Number of echidna host registrations and an estimated number of ticks sourced from natural history and museum collections according to Australian state and territory locality. State or Host Larvae Nymph Male Female Total Territory registrations (n) (n) (n) (n) (n) (n) NSW 13 0 16 11 35 62 NT 2 0 0 13 13 26 QLD 59 1 + *5x 148 + *2x 210 164 >1282 numerous numerous SA 10 0 179 + *1x 24.5** 16 >319.5** numerous TAS 4 2 2 0 6 10 VIC 6 0 10 1 14 25 Net Total 94 >503 >655 259.5 248 >1724.5** (n) *Numerous indicates one or more host registrations containing more than 100 ticks of a specific life stage or instar. For example, 1x numerous contains more than 100 (>100) ticks of a specific instar, and 2x numerous contains more than 200 (>200) ticks of a specific instar. ** 0.5 represents half a tick specimen

35 2.1.1 Archived ticks with correlating host tissue

Several frozen archived echidna specimens were located at the Queensland Museum. Ticks were removed from these archived echidna specimens (see Table 2.2) using gloves and sterilised forceps and placed into 5 ml vials containing 3 ml of 70% ethanol for preservation. Liver, spleen, kidney, and muscle tissue biopsies, including some skin samples (~2 to 3mm2) removed at the site of a tick bite, were retrieved from echidna specimens and placed into individual vials of 70% ethanol. Vials containing the tick and tissue specimens were stored at 4 °C. The geographical location of each echidna specimen and year of collection were documented, with biopsy type and tick location in relation to host site recorded and represented in Appendix A, Table A.3.

Table 2.2. Number of ticks according to instar removed from frozen archived echidna specimens, sourced from the Queensland Museum collection. Echidna Larvae Nymph Male Female Total specimen (n) (n) (n) (n) (n) (n) 1 0 25 13 5 43 2 0 3 20 4 27 3 0 0 1 6 7 4 0 2 12 0 14 5 0 3 10 20 33 6 0 0 6 3 9 7 1 2 0 1 4 Net Total (n) 1 35 62 39 137

2.2 Tick identification

Ticks were removed from their original collection tubes and examined in separate petri dishes using a Wild M3 Heerbrugg light microscope with a magnification power between 6.4 X to 40 X. The ticks were counted, with instar stage and species identified, and recorded into a spreadsheet (see Appendix A, Table A.1). Forceps and all other instruments used to handle the ticks were cleaned with 10% bleach and DNA AWAY™ (Molecular Bio-Products Inc., San Diego, USA) between samples, and work surfaces were sterilised with 10% bleach and 70% ethanol before and

36 after each examination. Following morphological identification, ticks of the same host registration were stored according to instar in new collection tubes containing 70% ethanol and at 4 °C. Species identification of the ticks was based on the morphological features outlined in the Australian tick morphology dichotomous key by Roberts (1970) and a genus reference key produced for the purpose of this study (see Appendix B, Table B.1) using additional information sourced from Barker and Walker (2014).

Ticks were photographed with an Olympus SC30 digital camera (Olympus, Center Valley, USA) and analysis getIT software (Olympus, Center Valley, USA) at a magnification ranging between 0.67 X to 4.5 X. An Olympus SZ61 stereomicroscope (Olympus, Center Valley, USA) was used with a SCHOTT KL 1500 LED (SCHOTT AG, Mainz, Germany) light source.

2.3 Sample selection

All sample handling and accompanying procedures were performed in a dedicated ancient DNA (aDNA) clean laboratory facility at Murdoch University, Western Australia, following stringent procedures to prevent contamination from modern DNA. Identified tick species were segregated according to Australian state and territory locality (see Section 3.1.1, Table 3.2). In addition, the total number of host registrations per state and territory location for the different tick species was used as the basis for sampling the ticks according to instar stage, for the purpose of selection for DNA extraction.

2.4 DNA extraction

Total genomic DNA was extracted from individual adult male and female ticks. Where more than one nymph or larva was present for a particular echidna host, the ticks were pooled to increase DNA yield. The number of ticks selected from each host for DNA extraction is represented in Appendix A, Table A.2.

37 2.4.1 Archived echidna ticks

The following sample preparation, digestion, and extraction methods were performed in a clean laboratory environment, as stipulated by Cooper and Poinar (2000). Prior to DNA extraction, ticks were surface sterilised with 10% bleach for 1 min followed by a 5 s vortex, before being submerged in 70% ethanol, vortexed, and rinsed in DNA-free water. Each tick was then moved to a clean petri dish where an ultra-violet (UV) light sterilised pin was used to puncture the ventral surface of the tick between the genital pore and anus (in adults) or an area on the ventral surface not affecting morphological identification (in nymphs and larvae). A fresh UV sterilised pin was used for each tick specimen. Ticks were placed in 2 mL safelock tubes (EppendorfTM, Germany) in preparation for digestion. A QIAGEN DNeasy® Blood & Tissue Kit (QIAGEN, Germany) and the supplementary protocol “Purification of total DNA from insects using the DNeasy® Blood & Tissue Kit” were used as the foundation for the following extraction method:

For unengorged adults, nymph, and larval ticks, 180 μL ATL and 20 μL Proteinase K was added to the safelock tubes containing the prepared ticks. For engorged adults, nymphs, and larval ticks (engorgement determined by their globular swollen appearance and decreased depth of festoon and scutum grooves), 360 μL ATL and 40 μL Proteinase K were added, with the amount recorded for subsequent referral. Safelock tubes with the solution and tick were vortexed to mix and spun lightly on a mySPINTM Mini Centrifuge to collect the solution and tick at the bottom of each tube. Tubes were placed in a shaking heating block to be incubated at 56 °C while shaking at 700 rpm, for 16 to 18 h. Safelock tubes were removed from the heating block and centrifuged (Hettich MIKRO 220 Centrifuge, UK) at 15,000 rpm for 1 min. For unengorged ticks, 200 μL of supernatant was removed and added to 1.5 mL tubes containing 400 μL of 1:1 AL/ethanol solution. For engorged ticks, 400 μL of supernatant was removed and added to tubes containing 800 μL of 1:1 AL/ethanol solution. Tubes containing the solution were vortexed and spun briefly. Using a pipette, 600 μL of the solution in each 1.5 mL tube was transferred into a spin column, which was placed in a collection tube, and spun at 8,000 rpm for 1 min, with flow-through discarded. For engorged samples, this process was repeated with the remaining 600 μL of solution. Next, 500 μL AW1 wash buffer

38 was added to each spin column, centrifuged at 8,000 rpm for 1 min, and flow- through discarded. 500 μL AW2 wash buffer was then added to each spin column, centrifuged at 15,000 rpm for 3 min, with flow-through discarded. Spin columns were placed into sterilised 1.5 mL tubes (EppendorfTM, Germany) with lids removed. AE buffer was directly added to the spin column membrane, with 60 μL AE buffer for unengorged samples, and 100 μL AE buffer for engorged samples. Samples were subsequently incubated at room temperature for 4 min, before being spun at 8,000 rpm for 1 min. A second elution was performed with the flow- through pipetted back over the spin column membrane and incubated again for 4 min, followed by a centrifuge at 8,000 rpm for 1 min. The flow-through in the 1.5 ml tubes containing the DNA was added to individually labeled 2 mL EppendorfTM DNA LoBind tubes (EppendorfTM, Germany) and stored in a freezer at -20 °C. Extraction reagent controls were run in parallel with the DNA extractions in order to assess cross contamination and provide a background bacterial profile of the clean laboratory. Finally, ticks subject to DNA extraction were placed into new collection tubes containing 3 mL of 70% ethanol and labeled with a modified accession number for future reference.

2.4.2 Archived echidna tissue

The QIAGEN DNeasy®Blood & Tissue Kit (QIAGEN, Germany) protocol for “tissue” was used for sample preparation, digestion, and DNA extraction. The DNA was eluted in 100 μL AE buffer and stored in a freezer at -20 °C until further analysis. Extraction reagent controls were run in parallel with the DNA extractions in order to assess cross contamination and provide a background bacterial profile of the laboratory.

2.5 Library preparation and NGS

Using the V4 hyper-variable region of the bacterial 16S gene, due to its short length (~300 bp), high bacterial diversity, and success in tick microbiome assays (Sperling et al., 2017), the echidna tissue (n = 11) and tick samples (n = 66) were PCR amplified using the primers 515F and 806R (see Section 2.7, Table 2.3). Each sample was amplified with primers that contained an Illumina MiSeq adapter. All

39 community profiling PCRs were carried out in 25 μL reactions containing 0.5 U KAPA Taq DNA polymerase and 2.5 μL 10 X KAPA Taq Buffer + dye

(KAPABiosystems, Massachusetts, USA), 1.0 μL MgCl2, 0.25 mM dNTPs (FisherBiotec, Australia), 0.4 μM of each 515F and 806R primer (Integrated DNA Technologies, California, USA), including 1.0 μL BSA (FisherBiotec, Australia) and 2.0 μL of DNA. No-template controls (NTCs) and extraction reagent blanks (EXTs) were included in every PCR run and were incorporated into the sequencing libraries. All PCR amplifications were performed on a BioRAD T100TM conventional PCR machine with the following thermal conditions: initial denaturation at 95 °C for 5 min followed by 40 cycles of denaturation at 95 °C for 30 s, annealing at 55 °C for 30 s, and extension at 72 °C for 30 s. Thermal cycling was followed by a final extension at 72 °C for 5 min. Amplicon samples were subsequently PCR indexed with a unique forward and reverse primer in 25 μL volumes which consisted of 12.5 μL KAPA HiFi HotStart 2 X mix, 2.5 μL of Nextera XT Index 1 primer (N7XX) and 2.5 μL of Nextera XT Index 2 primer (S5XX), and 1.5 μL of amplicon DNA. Thermal cycling followed the protocol (Illumina 16S Metagenomic Sequencing Library Preparation) with the exception of 15 cycles performed.

Amplicon library preparation was executed according to the recommended protocol (Illumina 16S Metagenomic Sequencing Library Preparation) with the following alterations: uniquely indexed samples were electrophoresed on a 2% (w/v) gel (see Section 2.8) to maximise band separation, and a 100 bp ladder used as a reference for amplicon band size. The DNA bands were visualised using a UV transillumination and an AlphaDigiDoc transillumination system (BioRad, Hercules, USA), with images retrieved, photographed, and saved using a Cannon camera and AlphaDigiDoc software. Indexed samples of similar band intensity were combined in equal volumes to make up four libraries labeled A, B, C, and D. NTCs and EXTs were also included in the libraries to establish background bacterial populations that would be deducted in later sequence analyses. Libraries were run on a 2% (w/v) agarose gel electrophoresis and the correct band for each library excised using a sterile scalpel and placed into it’s respective labelled 1.5 mL tube (EppendorfTM, Germany). The QIAquick® Gel Extraction Kit protocol (QIAGEN, Germany) was used to purify the DNA within each library (see Section 2.9).

40 Libraries were subsequently quantified using the Qubit® 2.0 Fluorometer (FisherBiotec, Australia) and each diluted to an equimolar concentration before being pooled in equimolar amounts to create the final library. A minimum of 5% PhiX library, used as an internal control for the Illumina platform to increase sequence diversity within the NGS run, was combined with the final amplicon library. This uniquely indexed final library was then loaded onto an Illumina MiSeq using 500-cycle V2 chemistry (250 bp paired-end reads) following the manufacturer’s recommendations. All pre-PCR and post-PCR procedures were performed in physically separated laboratories to minimise amplicon contamination.

2.6 NGS analysis

Sequences were originally subjected to quality control procedures as formerly described (Gofton et al., 2015a) with the following exceptions: paired-end reads were merged using USEARCH (version 8) (Edgar, 2010) with a minimum overlap length of 50 bp and no gaps allowed in the merged alignments. Primer sequences and distal bases were trimmed from the ends of reads in Geneious (version 8.1.9) (Kearse et al., 2012) and reads shorter than the minimum previously reported length of the bacterial 16S V4 amplicon (< 250 bp) were removed. Singleton sequences (per sample) and sequences with a > 1% error rate were removed from the dataset using USEARCH (version 8) (Edgar, 2010). Operational taxonomic units (OTUs) were created by clustering sequences at 97% similarity with the UPARSE algorithm (Edgar, 2013), and taxonomy was assigned to OTUs in QIIME (Caporaso et al., 2010) by aligning to the GreenGenes 16S database (DeSantis et al., 2006) using the UCLUST algorithm (Edgar, 2010) with default parameters. OTUs taxonomically assigned to the genus-level were used for further analysis. OTUs that were present in EXT and NTCs were deducted from all samples in order to eliminate potential contaminating and background bacterial sequences.

Following OTU analysis to assign genus level taxonomy to 16S sequences, the Basic local alignment search tool (BLAST) (Altschul et al., 1990) was used to resolve the species identity of bacterial genera that have medical or veterinary significance, or contain members that are known, or proposed, tick pathogens. Species-level taxonomy was only inferred when the query matched 16S sequences from one

41 species with a ≥ 99% pairwise identity over ≥ 99% the length of the query sequence. Bacterial genera that were deemed not of medical or veterinary significance, or otherwise previously associated with ticks, are not discussed.

2.7 PCR assays

A universal 16S mammalian-specific primer set was initially used on all samples and performed using a dilution series to evaluate efficiency of PCR assays. To confirm NGS results and gain further information on phylogenetic positioning with other Borrelia spp., genus-specific PCR assays targeting the flaB and longer 16S genes, for a combined analysis, were used on echidna-biting ticks and echidna tissue. The PCR assays and Borrelia-specific primers used in this study have been confirmed previously in the laboratory to reliably amplify Borrelia burgdorferi sensu lato (B. burgdorferi s.l.), relapsing fever (RF) borreliae, and ‘Candidatus Borrelia tachyglossi’ (Gofton et al., 2015a; Loh et al., 2016).

All pre-PCR sample handling was performed in a dedicated aDNA clean laboratory facility at Murdoch University, Western Australia, to reduce contamination from modern DNA and PCR amplicons. NTCs, EXT and positive controls were included in all PCR assays and observed to verify consistency.

42 Table 2.3 Primers used for NGS and PCR amplification. Primer sequences, annealing temperatures, and expected amplicon sizes are included. Gene Primer Sequence (5’ – 3’ Annealing Expected Reference orientation) temperature amplicon (°C) size (bp)

16S Bac 515F* GTGCCAGCMGCCGCGGTA A

806R* GGACTACHVGGGTWTCT 55 °C 291 Caporaso AAT et al., 2011

16S Mam Mam1 CGGTTGGGGTGACCTCGG A

Mam2 GCTGTTATCCCTAGGGTA 57 °C 90-100 Taylor, ACT 1996

16S External Borrelia Bor-16 F TGCGTCTTAAGCATGCAA GT

Bor-1360R 51 °C 1,344 Loh et al., GTACAAGGCCCGAGAACG 2016 TA

Internal

Bor-27 F CATGCAAGTCAAACGGAA TG

Bor-1232R ACTGTTTCGCTTCGCTTT 51 °C 1,205 Loh et al., GT 2016 flaB External

flaB_280F GCAGTTCARTCAGGTAAC GG

flaB_RL 52 °C 645 Barbour et GCAATCATAGCCATTGCA al., 1996; GATTGT Clark et al., 2013 Internal

flaB_LL ACATATTCAGATGCAGAC AGAGGT

flaB_737R GCATCAACTGTRGTTGTA 55 °C 407 Barbour et ACATTAACAGG al., 1996; Clark et al., 2013 * For NGS an Illumina MiSeq adapter was added to the primer sequence

43 2.7.1 16S Mam qPCR assay of echidna tissue

The successful extraction of DNA from the tissue biopsies was assessed using a mammalian-specific quantitative PCR (qPCR) assay targeting the ubiquitous 16S gene. The 16S Mam primer set (see Section 2.7, Table 2.3) was also used to detect inhibition in extracted DNA samples of echidna tissue at neat and dilution ratios of 1:10 and 1:100 DNA/DNA-free water. The qPCR assay was performed in 25 μL volumes, and consisted of 12.5 μL Power SYBR® Green PCR Master Mix (Applied Biosystems by Thermo Fisher, UK), 0.4 μM each of Mam1 and Mam2 primer (Integrated DNA Technologies, California, USA), and 2.0 μL of neat or diluted DNA. Extracted DNA (n = 14) from skin, liver, spleen, kidney, and muscle echidna tissue deriving from echidna host number 1, 3 and 5 (see Appendix A, Table A.3) were screened using the StepOneTM Real-Time qPCR machine (version 2.1, Applied Biosystems, Foster City, USA). Thermal cycling included an initial hold cycle of 95 °C for 5 min, followed by 30 cycles of denaturation at 95 °C for 36 s, annealing at 57 °C for 30 s, and extension at 72 °C for 30 s, with a final replication on all templates at 72 °C for 7 min.

2.7.1.1 16S Mam conventional PCR assay of correlating echidna tissue ticks

The 16S Mam primer set (see Section 2.7, Table 2.3) was used to determine the presence of mammalian tissue in the echidna ticks (n = 46) removed from seven archived echidna specimens (see Appendix A, Table A.2; A.3). The PCR assay was carried out in 25 μL volumes, consisting of 0.5 U KAPA Taq DNA polymerase and 2.5 μL 10 X KAPA Taq Buffer + dye (KAPABiosystems, Massachusetts, USA), 0.5 μL

MgCl2, 0.25 mM dNTPs (FisherBiotec, Wembley, Australia), 0.4 μM of each Mam1 and Mam2 primer (Integrated DNA Technologies, California, USA), and 2.0 μL of DNA. Thermal cycling (see Section 2.7.1) included 35 cycles in total.

44 2.7.2 Borrelia-specific PCR assays of echidna tissue and ticks

Two Borrelia genus-specific nested-PCR assays were performed targeting the flaB and longer 16S genes. The purpose of amplifying the flaB gene was to detect and further confirm the presence of Borrelia in archived echidna tissue and echidna- biting ticks. Amplifying an extended region of the 16S gene was necessary to establish a more accurate phylogenetic position in relation to other borreliae, and in particular the relationship with ‘Candidatus B. tachyglossi’ (Loh et al., 2016; 2017).

2.7.2.1 flaB nested-PCR assay

A preliminary qPCR assay using the flaB gene external primer set flaB_280F and flaB_RL (see Section 2.7, Table 2.3) was performed on echidna tissue (n = 8) at neat, 1:10, and 1:100 dilution ratios to determine inhibition and hence the best DNA concentration for future analyses. The assay was divided into two groups: A) echidna tissue DNA at neat, 1:10 and 1:100 dilutions, and B) echidna tissue DNA at neat, 1:10, and 1:100 dilution spiked with a confirmed flaB positive. The qPCR assay was completed in 26 μL volumes for both group A and group B, and consisted of 12.5 μL Power SYBR® Green Master Mix (Applied Biosystems by Thermo Fisher, UK), 0.4 μM of primers flaB_280F and flaB_RL (Integrated DNA Technologies, California, USA), and 2.0 μL of neat or diluted DNA. Group B incorporated 1.0 μL of flaB positive DNA as a spike test (this was replaced as 1.0 μL of DNA-free water in group A). The prepared DNA samples from both groups were screened in parallel using the StepOneTM Real-Time qPCR machine (version 2.1, Applied Biosystems, Foster City, USA). The qPCR set-up followed an initial hold cycle of 95 °C for 5 min, with 35 cycles of denaturation at 95 °C for 30 s, annealing at 52 °C for 30 sec, and extension at 72 °C for 1 min. Thermal cycling was concluded by a final extension at 72 °C for 5 min.

Extracted DNA from echidna tissue (n = 34) of a 1:10 dilution ratio, were amplified on a BioRAD T100TM conventional PCR machine using the flaB external primers flaB_280F and flaB_RL (Loh et al., 2016). A conventional PCR assay was carried out in 25 μL volumes, consisting of 0.5 U KAPA Taq DNA polymerase and 2.5 μL 10 X

45 KAPA Taq Buffer + dye (KAPABiosystems, Massachusetts, USA), 0.5 μL MgCl2, 0.25 mM dNTPs, 0.4 μM of primers flaB_280F and flaB_RL (Integrated DNA Technologies, California, USA), and 2.0 μL of DNA. The primary PCR set-up was an initial hold cycle of 95 °C for 5 min, followed by 35 cycles of denaturation at 95 °C for 30 s, annealing at 52 °C for 30 s, and extension at 72 °C for 1 min. A final extension of 72 °C for 5 min completed the primary PCR. For the secondary PCR set-up the flaB internal primers flaB_LL and flaB_737R (see Section 2.7, Table 2.3; Loh et al., 2016) with the same reagent volumes described above were used, and the annealing temperature modified to 55 °C. All other PCR temperatures and times remained unchanged.

Extracted DNA from ticks (n = 160) of a neat concentration were amplified on a BioRAD T100TM conventional PCR machine using the flaB external primers (flaB_280F and flaB_RL) and internal primers (flaB_LL and flaB_737R) with the same reagent volumes and primary and secondary PCR set-up conditions as described for conventional PCR above.

2.7.2.2 Borrelia-specific 16S nested-PCR assay

A Borrelia-specific 16S nested-PCR assay was performed on the echidna tissue and tick samples that were positive at the flaB locus. For the external primers Bor-16 F and Bor-1360R and subsequent internal primers Bor-27 F and Bor-1232R (see Section 2.7, Table 2.3; Loh et al., 2016), 25 μL volumes were carried out using the same reagent quantities described for the flaB conventional nested PCR assay (see Section 2.7.2.1). Thermal cycling was conducted on a BioRAD T100TM conventional PCR machine, and involved identical temperatures and times for both the primary PCR and secondary PCR set-up: initial hold at 95 °C for 5 min, followed by 35 cycles of denaturation at 95 °C for 30 s, annealing at 51 °C for 40 s, and extension at 72 °C for 2 min, with a one cycle final extension of 72 °C for 5 min to complete.

46 2.8 Gel electrophoresis

PCR products were run on a 1- 2% (w/v) agarose gel electrophoresis. The agarose (FisherBiotech Multi-Purpose Grade, Wembley, Australia) was dissolved in 1 X TAE buffer and stained with 1 X Gel Red (FisherBiotech, Wembley, Australia). A 100 bp molecular weight ladder (Promega, Madison, USA) was used to determine the size of the flaB gene PCR products, and a 1 Kb ladder (Promega, Madison, USA) used to establish the size of the 16S gene PCR products. The DNA bands were visualized using a UV transillumination and an AlphaDigiDoc transillumination system (BioRad, Hercules, USA), with images retrieved and saved using a Cannon camera and AlphaDigiDoc software. DNA bands of interest were excised from the agarose gel using a sterilised scalpel and stored in a post-PCR freezer at -20 °C, in labeled 1.5 mL tubes (EppendorfTM, Germany).

2.9 PCR purification

The QIAquick® Gel Extraction Kit protocol (QIAGEN, Germany) was used for purifying the DNA obtained subsequent to gel electrophoresis, with the following modifications: each tube (EppendorfTM, Germany) containing a DNA sample had 450 μL of QG buffer added prior to incubation at 50 °C for 10 min. This timeframe was extended in the case of a gel slice not being fully dissolved. The addition of 150 μL of isopropanol to each DNA sample proceeded, and a QIAquick® spin column (QIAGEN, Germany) inserted into a 2 mL collection tube. The centrifugal method outlined in the QIAquick® Gel Extraction Kit protocol was performed (Hettich MIKRO 220 Centrifuge, UK), with 8,000 rpm for 1 min after the addition of the sample to the spin column, and later the 500 μL QG buffer, and 750 μL PE buffer. After the addition of PE buffer, the tubes were incubated at room temperature for 4 min and centrifuged at 35,000 rpm for 1 min. Lastly, 30 μL of EB buffer was added to elute the DNA. The purified DNA was stored in a post PCR freezer at -20 °C in labeled 1.5 mL tubes (EppendorfTM, Germany).

47 2.10 Sanger sequencing and BLAST

Purified PCR products were sequenced with both forward and reverse secondary PCR primers for flaB and 16S Borrelia genes (see Section 2.7, Table 2.3) on an ABI 373096 Capillary Sequencer using a Big Dye version 3.1 Terminator Cycle Sequencing Kit (Applied Biosystems, Massachusetts, USA). The sequencing reactions were performed in 10 μL volumes and consisted of 3.2 μL of primer (forward or reverse), with 1.5 μL of 5 X reaction buffer (Applied Biosystems, Massachusetts, USA), 1 μL of Big Dye version 3.1 (Applied Biosystems, Massachusetts, USA), and 4.5 μL of DNA for flaB gene sequencing; and 1 μL of 5 X reaction buffer, 2 μL Big Dye version 3.1, and 4 μL of DNA for 16S gene sequencing. PCR conditions included a preliminary step of 96 °C for 2 min, followed by 25 cycles of 96 °C for 10 s, 55 °C (flaB) or 51 °C (16S) for 5 s, and 60 °C for 4 min, with a final hold temperature of 14 °C. Post reaction purification required each 10 μL reaction to be transferred into individual 0.5 mL tubes (EppendorfTM, Germany) for the following: 1 μL (125 mM) EDTA, 1 μL (3 M) Sodium Acetate, and 25 μL of 100% ethanol added to each tube and mixed by pipetting. Tubes were left to precipitate for 20 min at room temperature followed by a 30 min microfuge (Hettich MIKRO 220 Centrifuge, UK) at 14,000 rpm. The supernatant was removed using a vacuum and discarded, leaving the DNA pellet. The pellet was washed with 125 μL of 70% ethanol and spun for 5 min at 14,000 rpm. The supernatant was again removed by vacuum and the DNA pellet left to air dry in the dark for 15 min. Samples were placed in a freezer at -20 °C until sequenced.

Sanger sequences generated in this study were aligned and primers trimmed using Geneious (version 8.1.9) (Kearse et al., 2012). Individual flaB and 16S gene sequences followed by the derived consensus sequence for flaB (378 bp) and 16S (1,087 bp) were compared against the NCBI GenBank nucleotide database using BLAST (Altschul et al., 1990) in an attempt to resolve species level taxonomy. Sequences were only assigned to a species if the query sequence matched a reference sequence with a pairwise identity match ≥ 99% with ≥ 99% query coverage.

48 2.11 Phylogenetic analysis

Phylogenetic analyses were conducted on the trimmed flaB (378 bp) and 16S (1,087 bp) sequences generated in this study, together with other Borrelia spp. sequences retrieved from GenBank. Sequences were aligned in Geneious (version 8.1.9) (Kearse et al., 2012) and the gapped alignment was refined using MUSCLE (Edgar, 2004). The software MEGA (version 7.0.21) (Tamura et al., 2013) was used to determine the most suitable nucleotide substitution model and selected based on the Bayesian Information Criterion (BIC). The general time reversible (GTR) model was chosen for the flaB sequences. Due to the short gene region, a phylogenetic distance model was deemed not suitable for the flaB sequences. Instead, the phylogenetic tree was generated using FastTree software (version 2.1.5) to infer an approximate-maximum-likelihood (Price et al., 2010), with optimised 20 rate categories of site. Bracyspira pilosicoli strain NR998 (GenBank: LC259310) (Ogawa et al., 2017) was used as the outgroup, so as to compare the phylogenetic positioning in this study with the previous flaB phylogeny produced by Loh et al. (2017). With the longer 16S gene suitable to infer genetic distance, the Borrelia 16S phylogenetic tree was generated using the HKY model. The 16S tree was constructed using the computing efficient clustering algorithm of the neighbour-joining Geneious Tree Builder software (version 8.1.9) (Kearse et al., 2012), with a Bootstrap resampling of 500 replicates to give a measure of confidence in the nodes. The outgroup used was Spirochaeta americana strain ASpG1 (GenBank: AF373921) (Hoover et al., 2003), for direct comparison to the previous study by Loh et al. (2016). Overall, two different outgroups as representatives of the phylum Spirochaetes were used to provide a more reliable inference for determining monophyly of the ingroups.

49 2.12 Percentage of Borrelia-infected echidna ticks

The percentage of echidna ticks (if any) found infected with Borrelia in the total sample of ticks (n = 160) analysed in this study, was determined using the following formulae:

As two detection techniques were used in this study, any duplicate Borrelia positive ticks are only recorded and formulated once.

CHAPTER THREE: RESULTS

51 3. Results

3.1 Morphological identification of archived echidna-biting ticks

A total of 850 echidna-biting ticks (see Table 3.1) retrieved from 89 registered echidna hosts, sourced from Australian natural history and museum collections, were morphologically identified in this study. Eight species from the genera Amblyomma, Bothriocroton, and Ixodes were recognised. Four species of Amblyomma; Amblyomma australiense (n = 15), Amblyomma echidnae (n = 1), Amblyomma fimbriatum (n = 22), and Amblyomma triguttatum (n = 19); three species of Bothriocroton; Bothriocroton concolor (n = 758), Bothriocroton hydrosauri (n = 12), and Bothriocroton tachyglossi (n = 13); and one species of Ixodes; Ixodes tasmani (n = 10), corresponded with the descriptions by Roberts (1970) and Barker and Walker (2014). The number of instars, for the eight tick species morphologically identified, are represented in Table 3.1. Furthermore, these species were documented according to locality and instar (see Appendix A, Table A.1), and instar photographs of the most prevalent tick species, B. concolor, are presented in Figures 3.1 – 3.3.

Figure 3.1. Larval pool of ticks recorded as B. concolor.

Figure 3.2. Nymph B. concolor tick.

Figure 3.3. Adult B. concolor ticks, with (A) dorsal view of B. concolor female and male, and (B) ventral view of B. concolor female and male.

53 Table 3.1. Number of instars for eight echidna-biting tick species morphologically identified from Australian archived collections. Tick species Larvae (n) Nymphs (n) Males (n) Females (n) Total (n)

Amblyomma australiense 0 1 6 8 15

Amblyomma echidnae 0 0 1 0 1

Amblyomma fimbriatum 0 0 10 12 22

Amblyomma triguttatum 0 0 12 7 19

Bothricroton concolor 2 342 223 191 758

Bothriocroton hydrosauri 0 4 7 1 12

Bothriocroton tachyglossi 3 0 0 10 13

Ixodes tasmani 2 2 0 6 10

Net Total (n) 7 349 259 235 850

3.1.1 Sample selection for DNA extraction and analyses

The eight tick species identified, as shown in Table 3.1, were separated based on their Australian state and territory locality. The ticks were further segregated according to the total number of host registrations for each species, recorded in Table 3.2. From this information a number of hosts were selected, and their ticks (of various instars) cleaned and prepared for total genomic DNA extraction (see Appendix A, Table A.2).

54 Table 3.2. Tick species with total number of host registrations according to state and territory location, including the number of hosts selected for tick DNA extraction, and subsequent number of tick extractions per species locality. Tick species State/Territory Total host Hosts selected Total tick location registrations (n) for tick DNA extractions (n) extraction (n) A. australiense QLD 4 3 5 NT 1 1 4 A. echidnae QLD 1 1 1 A. fimbriatum NT 1 1 6 A. triguttatum QLD 1 1 1 SA 1 1 2

B. concolor QLD 45 31 94

NSW 11 6 12 SA 9 6 12 VIC 6 3 5 B. hydrosauri QLD 2 2 9 B. tachyglossi QLD 3 2 2 I. tasmani TAS 4 4 7 Total (n) 89 62 160

Of the 850 ticks morphologically identified, a total of 160 of these ticks were selected for DNA extraction. Table 3.3 represents these ticks (n = 160) according to species and instar. Larvae were specifically not selected for so as to maximise the number of adults and nymphs analysed in this study and to compare results with the investigation by Loh et al. (2016). Also, as adult ticks require two blood meals compared to one for nymphs, the likelihood of retrieving a putative pathogen such as a Borrelia bacterium from infected hosts increases. Hence, more adults were selected for extraction and where possible, a select number of nymphs from the same host were pooled to increase DNA yield.

55 Table 3.3. Ticks selected for DNA extraction. Tick species Larvae (n) Nymph pool Male (n) Female (n) Total (n) (n) A. australiense 0 1 3 4 8 A. echidnae 0 0 1 0 1 A. fimbriatum 0 0 3 3 6 A. triguttatum 0 0 2 1 3 B. concolor 1 30 37 56 124 B. hydrosauri 0 1 7 1 9 B. tachyglossi 0 0 0 2 2 I. tasmani 0 1 0 6 7 Net Total (n) 1 33 53 73 160

3.2 NGS

Of the 20,185,992 sequences generated on the Illumina MiSeq NGS platform, only 1,180,566 sequences passed quality filtering, represented in Table 3.4. Figure 3.4 was used to judge the number of reads by quality score (QScore) that passed filter, set at Q30 which is indicative of a 0.1% chance of an incorrect base call during sequencing. As only 45.4% of reads had a Qscore of 30 or higher, the NGS run was deemed poor in regards to overall performance.

Figure 3.4. The number of reads (per million) generated during NGS that passed filter. Green bars represent reads with a QScore above 30, indicative of an acceptable 0.1% chance of an incorrect base call during sequencing. The high number of reads with a QScore below 30 (in blue) is suggestive of a poor run performance.

Following further NGS quality control filtering, 25,485 sequences from 66 echidna ticks, 11 echidna tissue biopsies, and seven extraction blank (EXT and no- template control (NTC) samples were used for analysis. A total of 102 of the 110 operational taxonomic units (OTUs) were assigned, and 18 bacterial genera found in the EXT and NTCs were removed from the dataset as background bacteria. These bacteria were either ubiquitous environmental or human-associated commensal bacterial genera that to date have not been associated with tick-borne human or veterinary disease.

57 Table 3.4. NGS sequencing statistics for bacterial 16S rRNA reads.

Sequencing statistics Total

Initial number of reads 20,185,992 Number of reads that passed filter 1,180,566 Merged number of reads 480,985 Filtered (by primer binding, size, unique barcode, and removal of singletons and chimeras) number of reads 25,485 Minimum reads per sample 2 Average reads per sample 303 Maximum reads per sample 5037 Samples 84 OTUs 110 Assigned OTUs 102

Most of the bacterial organisms identified in the eight echidna tick species (see Section 3.1.1, Table 3.3) consisted of environmental or commensal bacteria within the phylum Actinobacteria, Bacteriodetes, Cyanobacteria, Firmicutes, Planctomycetes, Proteobacteria, and Verrucomicrobia. Of the environmental genera identified, Lysinibacillus, Facklamia, Flavobacterium, Methylobacterium, Pseudomonas, and Stenotrophomones are considered ubiquitous in the environment and associated with soil and moist leaf litter environments in which ticks spend the majority of their life cycle. In addition, species of the genus Staphylococcus, which typically lives as commensal organisms on vertebrate skin, was identified in B. concolor and B. tachyglossi ticks, as well as echidna (Tachyglossus aculeatus) tissue samples.

Spirochaetes of the genus Borrelia were found in 6.3% of the B. concolor tick species, illustrated in Figure 3.5. Of these, 9.7% were female and 3.5% were male B. concolor ticks (see Figure 3.6). No nymphs of this tick species were Borrelia positive. In addition, no Borrelia was identified in the echidna tissue biopsies using NGS. In order to identify the Borrelia (OTU 6) present in B. concolor at species level, the most abundant unique sequence within OTU 6 was queried against NCBI GenBank sequences in Geneious (version 8.1.9), and a 99% identity match was

58 made to Borrelia sp. Aus_A and Aus_B (GenBank: KU954112; KU954113), present in modern B. concolor ticks (Loh et al., 2016) and renamed ‘Candidatus Borrelia tachyglossi’ (Loh et al., 2017). Concerning other bacterial genera of medical and veterinary significance, a Coxiella species (OTU 2 and OTU 5) was identified in 66.5% of B. concolor ticks, with Coxiella burnetii, the agent of Query (Q) fever, the most abundant sequence detected (GenBank: CP018005) (Walter & Frangoulidis, 2017). Species of the genus Ehrlichia (OTU 21) and Rickettsia (OTU 3) were also identified in B. concolor ticks at 0.2% and 19.5% respectively. In total, 2.4% of A. triguttatum ticks also contained an Ehrlichia sp., whereas Rickettsia was most common in 88.7% of I. tasmani and 18.2% of B. tachyglossi ticks. Echidna tissue analyses revealed that 0.2% contained Rickettsia spp. The genus Francisella (OTU 8) was also identified in 96% of A. triguttatum and 45% of A. australiense ticks. However, the Ehrlichia, Francisella, and Rickettsia sequences generated using NGS were not distinguishable at species level.

Figure 3.5. Bacterial diversity in Australian echidna ticks and echidna tissue (T. aculeatus). Generated in QIIME (Caporaso et al., 2010). In brown: phylum Spirochaetes, genus Borrelia. The most abundant phylum is Proteobacteria, represented in green-blue (aqua), and contains environmental bacteria as well as bacteria of medical and veterinary significance. Red symbolises taxonomically unassigned bacteria. The legend for the remaining bacterial phyla is included in Appendix C, Table C.2. 60

Figure 3.6. Bacterial diversity in Australian B. concolor echidna ticks according to instar. Generated in QIIME (Caporaso et al., 2010). In pale green: genus Borrelia. Other bacteria of interest are labelled. Ticks of a given instar positive for a particular bacterial genus are represented as a total percentage (2 decimal places). The legend for additional bacterial genera is included in Appendix C, Table C.3. For the bacterial diversity in the other echidna ticks and echidna tissue represented in this study, see Appendix C, Figure C.1.

61 3.3 PCR assays

3.3.1 16S Mam qPCR assay of echidna tissue

A qPCR assay targeting the mammalian 16S gene (see Section 2.7, Table 2.3) was used to determine successful DNA extraction, and to detect inhibition in extracted DNA samples of echidna tissue at neat and dilution ratios of 1:10 and 1:100 DNA/DNA-free water. A random sample (n = 14) consisting of skin, liver, spleen, kidney, and muscle DNA extracts from echidna host 1, 3, and 5 were subject to real-time qPCR analysis. Amplification was observed as most efficient in the 1:100 dilution ratio as shown with the consistent and synchronised amplification curve produced in Figure 3.7. An EXT and NTC were included, and their negative amplification ruled out contamination. The 1:10 dilution ratio, shown in Appendix D, Figure D.1, similarly amplified reliably, although slight variation in amplification curve suggested a poorer efficiency compared with the 1:100 dilution ratio. In comparison, the neat DNA samples represented in Appendix D, Table D.2 showed either no amplification or the presence of unstable and variable amplification curves, indicative of DNA inhibition.

Figure 3.7. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples of 1:100 dilution ratio. Amplification curves for the extraction blank and no-template control are labelled.

63 3.3.1.1 16S Mam PCR assay of correlating tissue ticks

The 16S Mam primer set (see Section 2.7, Table 2.3) was used to determine the presence of mammalian tissue in the echidna ticks (n = 46) at neat concentration, removed from seven archived echidna specimens (see Appendix A, Table A.2). A conventional PCR assay was performed with the results displayed in Figure 3.8.

Figure 3.8. Gel electrophoresis (1%) images (A, B, and C) of 16S Mam PCR products from neat echidna ticks sourced from correlating echidna hosts 1 to 7 (see Appendix A, Table A.2). Lane 1 in images A, B, and C contain the 100 bp DNA ladder. All five controls are labelled. PCR bands representing ticks e18, e19, e22, e23, e24, e25, e32, and e39 are enclosed in a yellow square.

65 As represented in Figure 3.8, mammalian DNA was determined as present in approximately 34 echidna ticks (34/46; 74%), illustrated by the positive banding. PCR bands for ticks of the extraction identification number e18, e19, e22, e23, e24, e25, e32, and e39 (see Appendix A, Table A.2) are highlighted as they correlate with the flaB positive ticks in Figure 3.11.

3.3.2 flaB nested-PCR assay

A qPCR assay using the flaB gene external primer set FlaB_280F and FlaB_RL (see Section 2.7, Table 2.3) was conducted on echidna tissue (n = 8) at neat, 1:10, and 1:100 dilution ratios to establish inhibition and consequently the optimum DNA concentration for subsequent conventional PCR assays. The assay was divided into two groups: A) echidna tissue DNA at neat, 1:10 and 1:100 dilutions, and B) echidna tissue DNA at neat, 1:10, and 1:100 dilution spiked with a confirmed flaB positive. Figure 3.9 represent the 1:100 DNA concentrations, which with the clean and consistent curves was deemed optimal for further analyses. For the neat and 1:10 dilution amplification plots, refer to Appendix D, Figure D.3 and D.4 respectively.

Figure 3.9. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples of 1:100 dilution ratio.

67 Amplification plots A) without positive spike DNA generated by the flaB qPCR assay reveal the likely absence of the flaB gene in the eight analysed samples, indicated by the observed flat-line and lack of exponential curve generated in amplification plots at neat, 1:10, and 1:100 DNA concentrations. However the amplification plots B) with positive spike DNA suggest that echidna tissue DNA dilutions of 1:100 dilution contain the least inhibition as shown by the improved amplification curve for plot B in Figures 3.9 (see Appendix D for related amplification plots). Therefore the 1:100 dilution ratio is most effective for proceeding flaB conventional PCR analysis on the echidna tissue samples.

3.3.2.1 The flaB gene in echidna tissue

A nested conventional PCR for the flaB gene was performed on echidna tissue DNA (n = 34) of 1:100 dilution ratio. Four controls were incorporated including an EXT, NTC, and two positive controls. The resulting PCR products are illustrated in Figure 3.10 as an imaged gel electrophoresis.

Figure 3.10. Gel electrophoresis (1%) image of flaB PCR products from echidna tissue of 1:100 dilution ratio originating from echidna hosts 1 to 8 (see Appendix A, Table A.3). Lane 1 in images A, B, and C contain the 100 bp DNA ladder. All controls are labelled. Positive tissue samples are identified.

69 Of the echidna tissue samples tested (n = 34) of 1:100 dilution ratio from the eight echidna hosts, two samples were positive for the flaB gene as revealed in Figure 3.10. These positive DNA bands for echidna tissue 13 and echidna tissue 23 (see Appendix A, Table A.3) correlate with both positive controls. Similarly, the absence of PCR product in the EXT NTC lanes is suggestive of no cross contamination and therefore validates the findings. These tissue samples positive for the flaB gene were subsequently analysed using Sanger sequencing (see Section 2.10).

3.3.2.2 The flaB gene in echidna ticks

A nested conventional PCR for the flaB gene was initially conducted on echidna ticks (n = 46) (see Appendix A, Table A.2) at neat concentration, which were retrieved from the seven archived echidna specimens. Included also were spleen tissue samples of 1:100 dilution ratio from echidna hosts 1, 2, 3, 4, 5, and 7, and a muscle sample of 1:100 dilution ratio from echidna host 6 (see Appendix A, Table A.3). The resulting gel electrophoresis is shown in Figure 3.11.

Figure 3.11. Gel electrophoresis (1%) image of flaB PCR products from neat echidna (n = 46) ticks originating from echidna hosts 1 to 7 (see Appendix A, Table A.2). Lane 1 in images A, B, and C contain the 100 bp DNA ladder. All four controls are labelled. Spleen tissue from echidna hosts 1, 2, 3, 4, 5, and 7, as well as muscle tissue from echidna host 6 at 1:100 dilution ratio are represented in image C, lanes 14 – 20.

71 The flaB gene was subsequently amplified in the remaining extracted ticks (n = 114) (see Appendix A, Table A.2) of a neat concentration using the same conventional nested-PCR assay. The resulting amplicons from the flaB positive tick samples were subsequently sequenced using Sanger sequencing (see Section 2.10).

3.3.3 Borrelia-specific 16S nested-PCR assay

A nested conventional PCR using the 16S Borrelia primer set (see Section 2.7, Table 2.3) was performed on the tissue (n = 2) and tick (n = 9) initial flaB positives (see Section 3.3.2). The positive tissues and ticks for the 16S Borrelia gene were then Sanger sequenced (see Section 2.10).

3.4 Sanger sequencing and BLAST

A partial fragment (378 bp) of the Borrelia flaB gene was successfully amplified in one male and eight female archived B. concolor ticks (9/160, 5.6%) and two echidna skin tissue biopsies (2/34, 5.9%). Both the B. concolor ticks and echidna tissue that were flaB positive originated from the state of Queensland, Australia. BLAST analysis showed that with 100% query coverage, these flaB sequences shared a 99.5-100% similarity with flaB sequences from novel Borrelia sp. isolates (GenBank: KX192144; KX192146; KX192147) (Loh et al., 2016). Further analyses of the flaB consensus sequence generated in this study revealed it most closely aligned with the flagellin ‘Candidatus B. tachyglossi’ genotype C (GenBank: KY586965) and genotype B (GenBank: KY586964) (Loh et al., 2017), with 99.7% identical sites and a 99.6% pairwise identity, and 99.5% identical sites and a 99.6% pairwise identity, respectively.

The Borrelia-specific 16S PCR assay was positive for six female B. concolor ticks and two echidna tissue biopsies (6/9, 66.7%; 2/2, 100%) of which were also positive at the flaB locus. BLAST analysis showed that with 97-100% query coverage, these 16S sequences (1,087 bp) produced in this study shared a 98.2- 99.6% similarity with the 16S sequence from Borrelia sp. Aus_B (GenBank: KU954113) (Loh et al., 2016), renamed ‘Candidatus B. tachyglossi’ genotype B (Loh et al., 2017).

72 3.5 Phylogenetic analysis

Trimmed flaB sequences (378 bp) and 16S sequences (1,087 bp) generated in this study were aligned using Geneious (version 8.1.9). The flaB phylogenetic tree is shown in Figure 3.12, and the Borrelia 16S phylogenetic tree is presented in Figure 3.13.

Figure 3.12. Phylogenetic analysis of 378 bp flaB gene consensus sequence. The maximum-likelihood tree was constructed using FastTree (version 2.1.5) (Price et al., 2010). Numbers at nodes represent branch support values. Branch length infers genetic distance. Bold type designates the consensus sequence obtained in the present study. * Represents the sequences from Loh et al. (2016) and are indicative of ‘Candidatus B. tachyglossi’ genotypes A, B, and C (Loh et al., 2017). Brachyspira pilosicoli used as outgroup. Rounded parentheses indicate GenBank accession numbers. 74

Figure 3.13. Phylogenetic analysis of 1,087 bp Borrelia 16S rRNA gene consensus sequence. The neighbour-joining tree was constructed using Geneious Tree Builder (version 8.1.9) (Kearse et al., 2012). Numbers at nodes represent branch support values. Branch length infers genetic distance. Bold type designates the consensus sequence obtained in the present study. * Represents the sequences from Loh et al. (2016) that are indicative of ‘Candidatus B. tachyglossi’ genotypes A, B, and C (Loh et al., 2017). Spirochaeta americana used as outgroup. Rounded parentheses indicate GenBank accession numbers.

75 BLAST results and phylogenetic reconstruction show that the flaB gene consensus sequence in this study is most similar to Borrelia sp. genotype C (‘Candidatus B. tachyglossi’ genotype C; 98.3%) followed by Borrelia sp. genotype B (‘Candidatus B. tachyglossi’ genotype B; 98.0%) (Loh et al., 2017). A high maximum-likelihood support value (0.981) at the node distinguishing the Novel Australian Group borreliae and Borrelia turcica, the single representative of the reptile-associated (REP) Group, suggests the two groups are related (85.5-87.9% sequence identity) yet distinctly separate clades (Figure 3.12). However, due to the short sequence length of the flaB gene, this tree lacks the accuracy necessary for producing a meaningful phylogenetic interpretation of genetic divergence, hence the longer Borrelia 16S gene has complemented it. The Borrelia 16S consensus sequence from this study clusters with the known genotypes of ‘Candidatus B. tachyglossi’ previously sequenced from B. concolor (Loh et al., 2016), and is most similar to Borrelia sp. Aus_B (‘Candidatus B. tachyglossi’ genotype B; 99.7%). The Novel Australian Group, which contains the 16S consensus sequence produced in this study, the ‘Candidatus B. tachyglossi’ genotypes, and Borrelia sp. NL230 (from an echidna parasitising Ixodes holocyclus tick), shows a high support value confidence (99.6%), which is suggestive of a monophyletic clade that does not group with the Lyme borreliosis (LB), REP, or relapsing fever (RF) borreliae (Figure 3.13).

3.6 Percentage of Borrelia-infected echidna ticks

Of the 160 museum-archived echidna ticks analysed in this study, a total of 10 ticks (6.25%) were found to harbour the novel Borrelia bacterium, ‘Candidatus B. tachyglossi’. A total of eight echidna tick species from six Australian states and territories were sampled and analysed using NGS and subsequent nested-PCR with Sanger sequencing. Nevertheless, only B. concolor ticks from Queensland, Australia, were positive for ‘Candidatus B. tachyglossi’. The six echidna ticks that were Borrelia positive using NGS were all positive at the flaB locus, however only two of these ticks were positively amplified using the longer 16S gene. Using nested-PCR with Sanger sequencing, a further three ticks were found to be Borrelia positive at the flaB locus, and were also positive at the longer 16S locus. All Borrelia positive ticks in this study were cross-validated using the technologies and associated gene regions represented in Table 3.5.

Table 3.5. Summary of the QLD B. concolor ticks according to instar that were positive for Borrelia genes in terms of percentage, using nested-PCR (with Sanger sequencing) and NGS technologies. NGS (n = 66) Nested-PCR (n = 160)

Instar Positive 16S Positive flaB Positive Borrelia 16S (n) (n) (n) Nymph 0 0% 0 0% 0 0%

Male 1 1.5% 1 0.6% 0 0%

Female 5 7.6% 8 5% 6 3.8%

Net Total (n) 6* 9.1% 9** 5.6% 6*** 3.8%

See Appendix A, Table A.2 for the following Borrelia positives: * ticks e23, e24, e25, e39, 30 and 31; ** ticks e18, e19, e23, e24, e25, e32, e39, 30 and 31; *** ticks e18, e19, e22, e23, e25, e32

A geographical comparison was made for the state of Queensland, between the Borrelia positive B. concolor ticks in this study and the Borrelia positive B. concolor ticks in the investigation by Loh et al. (2016). As depicted in Figure 3.14, in the current study, 10 Queensland B. concolor ticks (10/93; 10.8%) archived between the period of 1938 and 2011, and 35 Queensland B. concolor ticks (35/81; 43.2%) collected in 2016 by Loh and colleagues, contained ‘Candidatus B. tachyglossi’.

Figure 3.14. A comparison of the Australian geographical location of Borrelia positive B. concolor ticks, between the current study and the investigation by Loh et al. (2016). The frequency of Borrelia positive B. concolor ticks are expressed as a pie chart. Pie chart 1 represents QLD in the current study: Green = Borrelia positive ticks; Brown = total B. concolor ticks. Pie chart 2 represents QLD in Loh et al. (2016): Red = Borrelia positive ticks; Brown = total B. concolor ticks. Pie chart 3 represents NSW in Loh et al. (2016): Red = Borrelia positive ticks; Brown = total B. concolor ticks. A green symbol identifies the location of current study positive ticks; a red symbol identifies the location of Loh et al. (2016) positive ticks. Symbolises the location of the echidna with Borrelia positive skin biopsy and correlating positive B. concolor tick in current study.

CHAPTER FOUR: DISCUSSION AND CONCLUSION

79 4. Discussion

4.1 Morphological identification of archived echidna-biting ticks

The first objective of this study was to identify tick species that had been removed from echidnas (Tachyglossus aculeatus) and preserved in Australian museums and natural history collections. A total of 850 ticks were morphologically identified using the Australian tick identification key chartered by Roberts (1970) and the simplified tick genus key in Appendix B, produced using information sourced from Barker and Walker (2014). Of these ticks morphologically examined, 758 (89.2%) were of the species Bothriocroton concolor, confirming the first hypothesis, which was an unsurprising find considering B. concolor is a well-documented host of T. aculeatus (Roberts, 1970; Barker & Walker, 2014). Of the remaining seven species identified, four were previously documented echidna ectoparasites and include Amblyomma australiense, Amblyomma echidnae, Bothriocroton tachyglossi, and Ixodes tasmani (Roberts, 1960; Roberts, 1970; Andrews et al., 2006; Barker et al., 2014). An additional tick species identified, though loosely referenced by Roberts (1970) as a possible echidna ectoparasite, was Bothriocroton hydrosauri, aptly named the southern reptile tick with its main hosts recorded as lizards, snakes, and a terrestrial turtle (Roberts, 1970; Smyth, 1973). This record has remained questionable, as observations may be attributable to B. tachyglossi ticks instead, due to their similar external appearance (Andrews et al., 2006). Moreover, while some ticks from Australian museum and natural history collections came pre- identified as B. hydrosauri, upon further examination, many of these ticks were incorrectly identified. This misidentification was apparent when morphologically examining the tick hypostome of complete and undamaged specimens using a light microscope; the distinction in dentition was noticed between adult B. concolor, which have a 3 by 3 file of teeth on their hypostome, compared to adult B. hydrosauri, whose dentition represent a 2 by 2 file of teeth. Notably, not all specimens contained intact capitulum (head and mouthparts), making this discrepancy difficult and sometimes impossible based solely on the identification guide authored by Roberts (1970). In addition, nymph B. concolor ticks have a 2 by 2 dentition profile and can therefore also be mistaken for nymph B. hydrosauri ticks, especially with several undefined characteristics at this life stage.

80 Two tick species, Amblyomma trigutattum, the ornate kangaroo tick, and Amblyomma fimbriatum, the tropical reptile tick, were recorded from echidnas. These species were a novel discovery as A. triguttatum and A. fimbriatum had not been recorded to date as ectoparasites of the Australian monotreme, the echidna (T. aculeatus). Interestingly, as documented by Booth (1994), A. triguttatum has been retrieved from the only other extant Australian monotreme, the platypus, and may be suggestive of a possible host and perhaps mammalian order on which A. triguttatum might feed. However, as host-specific ticks are typically reluctant to feed on an unusual host (Sonenshine, 2005), concerning A. fimbriatum, a tick that is characteristically reptile-specific, a possible explanation for this exceptional finding on echidnas other than an accidental archived recording, may be attributed to opportunistic parasitism. Despite this, because it is unknown whether this species of tick had indeed fed on the echidna host it was removed from, it is unwise at this point in time to assume the echidna as a host of this species without further confirmation using blood-meal analyses (Léger et al., 2015). Lastly, previously recorded ticks Amblyomma moyi, Amblyomma papuanum, Bothriocroton undatum, and Ixodes holocyclus (Roberts, 1970) were not identified within the museum collections.

A noteworthy limitation of this study has been the accurate identification of tick species, and more specifically, tick instars, using the conventional techniques of microscopic examination. In particular, the morphology of larvae and nymphs was difficult to unequivocally determine using microscopy and the limited, outdated, and general lack of description in the Australian dichotomous tick key by Roberts (1970). More recently, advancements in molecular tools have supported species identification at the molecular level using the variation within the 18S and 16S genes, and enabled a more detailed insight into classification and phylogenetic relationships between tick genera (Lv et al., 2014). However, due to experimental constraints within this study, identification of ambiguous larvae and nymph tick specimens using a molecular approach could not be performed. Therefore, there remains a possibility for the inaccurate morphological identification of several larval and nymph tick specimens identified and subsequently analysed in this study.

81 4.2 NGS bacterial profiling of archived echidna-biting ticks

This study incorporated the use of next-generation sequencing (NGS) to survey the microbial communities in archived ticks found to parasitise echidnas. The principal aim of NGS bacterial profiling was to determine whether Borrelia and other tick-borne bacteria could be retrieved from non-destructively sampled archived ticks. In addition, 16S metagenomic analysis was used to better understand the diversity of microorganisms that make up the tick microbiome, important for future studies investigating the acquisition, transmission, and virulence of putative pathogens.

Firstly, this study has proven NGS to be a powerful tool for retrieving and identifying sequences of bacterial origin that are expected to contain degradation due to their age and prolonged storage in ethanol or at a frozen state. Furthermore, the use of a non-destructive DNA extraction technique that leaves the archived tick specimens morphologically intact has not impeded the retrieval of bacterial sequences, with this study amassing a total of 20,185,992 sequences, of which 25,485 were analysed subsequent to NGS quality control filtering.

Secondly, as hypothesised, archived ticks harbour a preserved cache of bacterial communities. Therefore, NGS bacterial profiling was able to detect the genus Borrelia in a total of six (6/66, 9.1%) archived ticks analysed in this study. Further analyses revealed this bacterium to share a 99% identical match to Borrelia sp. Aus_A and Borrelia sp. Aus_B (GenBank: KU954112; KU954113), the genotypes of the novel Borrelia sp., ‘Candidatus Borrelia tachyglossi’, recently described in modern echidna-biting B. concolor ticks (Loh et al., 2016; 2017). All Borrelia positive ticks in this study were of the species B. concolor, confirming an association between ‘Candidatus B. tachyglossi’ and B. concolor ticks, as previously determined by Loh et al. (2016; 2017). However, the absence of ‘Candidatus B. tachyglossi’ in other echidna-biting tick species was unexpected, especially since this Borrelia sp. was also found in a single I. holocyclus tick removed from an echidna (Gofton et al., 2015a), using the same NGS method as in the current study. Furthermore, as I. holocyclus is not a usual ectoparasite of echidnas, surveying the microbial communities of ticks that share a close association with echidnas such as A. echidnae and B. tachyglossi, was expected to reveal further Borrelia sp. isolates.

82 NGS bacterial profiling was also performed on several archived echidna skin and spleen biopsies. Skin and spleen specimens were analysed because Borrelia spp. typically enter the host via the skin and subcutaneous tissues, before accumulating and multiplying in various organs, including the spleen (Cook & Zumla, 2009). Furthermore, spleen tissue has been shown to frequently harbour Borrelia (Anderson et al., 1986). However, no Borrelia was detected in these tissue samples using NGS.

In total, 102 OTUs were assigned to bacterial taxa, of which 46 were identified to genus level in this study. In a recent NGS tick microbiome analysis, Gofton et al. (2015a) reported 199 OTUs, post-removal of environmental contaminants. The current study targeted a different hyper-variable region of the 16S gene, in addition to different tick species, however a lower number of OTUs were identified. This is likely attributed to the lower number of sequences that passed filtering during the NGS run. Specifically, only 45.4% of reads had a quality score (Qscore) of 30 or higher, indicative of an acceptable 0.1% chance of an incorrect base call during sequencing. Consequently, far fewer sequences were obtained than anticipated, and the run deemed poor in regards to overall performance. Technical issues with the MiSeq optical components that image the flow cell during the NGS run are believed responsible for this result. A poor run and the subsequent stringent quality filtering to remove sequencing artifacts, such as low-quality reads and contaminating reads (Gofton et al., 2015b), have compromised the extent of bacterial coverage obtained from the run, however a number of high quality sequences were still retrievable for analyses.

Expected bacterial genera detected in this study included Coxiella, Ehrlichia, Francisella, and Rickettsia. Together with known tick-borne pathogens, these bacterial genera also comprise of closely related endosymbiotic bacteria associated with ticks. Coxiella was sequenced from 17 B. concolor ticks, Ehrlichia was obtained from one A. triguttatum and one B. concolor tick, as well as an echidna spleen biopsy, and Francisella was found in two A. australiense and three A. triguttatum ticks, one of which also contained Ehrlichia. Rickettsia was detected in 20 B. concolor ticks followed by two I. tasmani ticks and one B. tachyglossi tick. Rickettsia was also detected in an echidna skin biopsy. Since bacterial

83 endosymbionts often dominate the microbial population within ticks and can therefore mask potential pathogens due to their high copy number (Gofton et al., 2015a; b), species level discrimination is of paramount importance to establishing potential pathogens.

An unanticipated discovery in B. concolor ticks was the pathogenic agent of Query (Q) fever, Coxiella burnetii, which until this study was not a reported bacterium in the echidna-specific B. concolor tick. Unfortunately, Ehrlichia, Francisella, and Rickettsia 16S sequences were unidentifiable at species level due to the high sequence homology (> 99%) between many of these species at the 16S locus analysed in this study. This finding was mirrored in the study by Gofton and colleagues who were unable to provide a species designation for Rickettsia sequences generated using NGS (Gofton et al., 2015a). Although not further analysed due to constraints on this current study, it is recommended that in future studies a genus- and species-specific PCR and Sanger sequencing of a more informative genetic marker are coupled with NGS to resolve the identity of the Ehrlichia, Francisella, and Rickettsia spp.

Sequencing of the 16S gene has allowed a determination of the different microbial populations within these complex tick microbiomes. Nevertheless, bacterial profiling using NGS has a significant limitation in that it can be biased towards the exaggeration of abundant taxonomic groups, typically belonging to bacterial endosymbionts, such as the high proportion Rickettsia sequences generated in this study. Although this can ultimately produce a skewed bacterial composition and false representation of proportions, favouring the abundant bacteria and high copy sequences, what it cannot exaggerate is the bacterial diversity within these various tick species. However, NGS can under represent diversity. Therefore, incorporating primers that block abundant endosymbiont taxa is required to increase diversity (Gofton et al., 2015a).

Finally, it is important to mention that although the external surface of all ticks was decontaminated with 70% ethanol and 10% hypochlorite solution prior to molecular analyses, common environmental and commensal bacteria were still prevalent among all tick microbiomes surveyed in this study. This is most likely

84 due to remnant bacterial DNA that survived the decontamination process, perhaps in bacterial plaques that may have accumulated in less accessible regions, such as leg joints or underneath the tick’s palps and between the barbs of the hypostome. Therefore, careful dissection of the tick’s main internal tissues (midgut, salivary glands, and sexual organ) may prove useful in distinguishing the microbiome of the internal tissues from environmental bacteria present on the tick’s external surfaces, including their exact location within the tick. Though this would unequivocally be at the cost of specimen preservation.

4.3 Detecting a novel Borrelia species in archived echidna-biting ticks

The main purpose of the current investigation was to provide further evidence for Borrelia in Australian echidna-biting ticks (Gofton et al., 2015a; Loh et al., 2016). This study incorporated a larger and more diverse sample of echidna-biting ticks than previously examined to date, which was made possible by accessing Australian museum and natural history archives. As mentioned (see Section 4.2), a NGS survey of echidna tick and tissue biopsies revealed ‘Candidatus B. tachyglossi’ in 9.1% of analysed samples, all of which were of the tick species B. concolor. Subsequent nested-PCR assays with Sanger sequencing, performed on a larger sample of archived tick and tissue biopsies, and targeting the Borrelia-specific flaB and 16S loci, revealed Borrelia in a total of 10 (10/160, 6.25%) ticks. Again, all ticks were of the species B. concolor. Six of these ticks were repeat positives formerly detected by NGS. Consequently, the ticks found to be Borrelia positive were all cross-validated by a combination of NGS and a nested-PCR at the flaB or 16S locus, with Sanger sequencing. Following phylogenic analyses, all 10 ticks were identified as positive for the species ‘Candidatus B. tachyglossi’, with 16S phylogenetic reconstructions indicating a close genetic similarity to ‘Candidatus B. tachyglossi’ genotype B (GenBank: KU954113) (Loh et al., 2016; 2017). Phylogenetic positioning at the flaB and 16S loci provided further evidence to suggest ‘Candidatus B. tachyglossi’ genotypes, inclusive of Borrelia sp. NL230 (echidna I. holocyclus isolate), and the consensus sequences generated in this study, form a monophyletic clade and hence support a fourth group in the Borrelia genus.

85 An unanticipated result from the nested-PCR assays was the first detection of ‘Candidatus B. tachyglossi’ in two archived echidna skin biopsies, one of which had a corresponding ‘Candidatus B. tachyglossi’ positive B. concolor tick that had been removed from the echidna skin prior to a separate DNA extraction. As the NGS survey did not detect Borrelia in any of the echidna tissue biopsies sequenced, this result, though unexpected, was not unjustifiable given the low number of reads generated, and that passed quality filtering, from the NGS run. These Borrelia positive skin biopsies originated from two different echidna hosts and both had amplified at the flaB and 16S loci. While ‘Candidatus B. tachyglossi’ was identified in both echidna tissue and tick specimens, it is unclear if the presence of ‘Candidatus B. tachyglossi’ was due to transmission by the infected feeding tick or whether the tick acquired the bacterium from a bacteraemic echidna host. Therefore, whether ticks obtain Borrelia from infected echidnas remains to be confirmed through additional tick analyses and the examination of echidna blood samples.

Despite the poor NGS run performed in this study, the overall prevalence of Borrelia was 9.1% by NGS, but only 6.25% by nested-PCR. Two explanations are provided for this discrepancy. First, for NGS the amplicon size was short, at approximately 300 bp, compared to the nested PCR amplicon, which for the flaB gene was marginally longer at 420 bp but significantly longer for the 16S gene, at 1.3 Kb. Therefore, it was anticipated that the shorter gene would amplify with a much greater efficiency (McPherson & Møller, 2006; Su et al., 2011). Second, NGS permits the detection of multiple, mixed sequences of low copy number and fragmented quality which are typically difficult or impossible to detect using Sanger sequencing (Su et al., 2011).

Regarding the nested-PCR assays, the reduced sensitivity of detecting Borrelia with the 16S PCR assay compared to the flaB PCR assay, may be explained by the reduced PCR efficiency that is typically observed when amplifying longer gene fragments (Loh et al., 2016). For this reason, PCR assays that target short gene fragments, such as the flaB assay used in this study, are recommended for sensitive detection of Borrelia spp. However, because short gene fragments are often inadequate for constructing meaningful phylogenetic interpretations, it is

86 suggested that these shorter genes are complimented by PCR assays that target and amplify longer gene sequences, albeit with less sensitivity, such as the 16S assay used in this study.

The investigation by Loh et al. (2016) revealed a higher percentage of Borrelia- infected echidna ticks at approximately 40%, compared to the 6.25% infected in the current study. Apart from a difference in sample size and range, the timeframe over which the archived ticks were initially collected and tick diversity, this discrepancy may be explained by the following hypotheses. Firstly, the occurrence of ‘Candidatus B. tachyglossi’ may have been limited in ticks prior to the 2015 discovery by Gofton et al. (2015a), as the bacterium may have had a restricted geography. Secondly, the nature of DNA fragmentation in archived specimens means that sequences of interest may be too degraded for detection, even with the aid of in-depth sequencing technologies such as NGS, therefore resulting in fewer detectable sequences overall.

Lastly, two surprising outcomes emerged from this study. First was the geographical distribution of ‘Candidatus B. tachyglossi’, which has previously been described in echidna ticks originating from Queensland and New South Wales by Loh et al. (2016). Approximately 43.2% of infected echidna B. concolor ticks originated from Queensland in the study by Loh and colleagues, therefore it was anticipated that the current study would reveal the majority of Borrelia positive ticks as originating from this Australian state. However, not expected was the absence of Borrelia-infected echidna ticks from neighbouring states including New South Wales. As host movement typically aids the geographical distribution of ticks (Soneneshine, 2005), it is curious that ‘Candidatus B. tachyglossi’, if indeed spread by bacteraemic echidnas, was not detected in echidna ticks, particularly of the species B. concolor, originating from nearby Australian regions. Echidnas have a ubiquitous distribution in Australia and are notorious for sharing home ranges that can extend several hectares (Augee et al., 2006). Hence, the absence of Borrelia in echidna-biting ticks from other Australian states is interesting and warrants further exploration.

87 The second interesting find resulting from this study was the detection of ‘Candidatus B. tachyglossi’ in more frozen echidna tissue and corresponding tick specimens compared with the ethanol preserved ticks and single tissue specimen. Although this may be coincidental, another thought is that it may potentially elucidate a downside to the quantity and quality of molecular data obtained from ethanol preserved samples (Dean & Ballard, 2001). However, without a direct comparison between the quality of DNA retrieved from each preservation method, the extent of DNA fragmentation and possible damage caused by storage within this chemical environment remains unknown.

4.4 Future directions

Throughout this investigation various techniques have been applied to gain a further understanding of the species of tick that parasitise the Australian echidna (T. aculeatus), and to reveal any microbial associations. However, there are areas of improvement that are recognised and are herein mentioned for future explorations in this particular field. Firstly, the tick key developed by Roberts has been the gold standard for morphological identification of Australian ticks since its conception in 1970. However, with ongoing modifications to tick names, their descriptions, and the discovery of new species (Ash et al., 2017), this key is now out-dated and lacks representation of recent findings. With advances in DNA technologies, molecular characterisation is an alternative method to distinguish these tick species at all life stages, something that has proven difficult particularly at the microscopically indistinct nymph and larval stages. Consequently, for future studies it is suggested that molecular tools are used to complement conventional techniques and hence lessen the possibility of misidentification between species of close resemblance. Secondly, surveying the bacterial communities within ticks using NGS is useful for establishing diversity. However, to greatly improve the understanding of species that colonise the tick microbiome, the incorporation of genus- and species-specific PCR with Sanger sequencing is vital for species level discrimination and ensuing phylogenetic interpretations. Lastly, this study analysed the most diverse range to date of archived echidna-biting ticks, with the main purpose to establish if ‘Candidatus B. tachyglossi’ could be found in different species of echidna ticks. This was an important investigatory step and the

88 successful retrieval of this bacterium from historic echidna ticks and correlating echidna skin samples has proven its past existence in the Australian ecosystem. However, the absence of this bacterium in all but one echidna-biting tick species currently restricts the proposal that this bacterium is vectored from infected echidna hosts. Consequently, a further analysis of a larger and more diverse sample of modern echidna ticks and their host blood is now needed to accompany this current study.

4.5 Conclusion

This study presents the first comprehensive morphological audit of echidna ticks. Two tick species, A. trigutattum and A. fimbriatum, were identified as new echidna ectoparasites. In addition, this was the first NGS survey of the archived tick microbiome. Identified genera of medical and veterinary significance included Borrelia, Coxiella, Ehrlichia, Francisella, and Rickettsia. Further analyses using Borrelia genus-specific PCR assays with Sanger sequencing revealed ‘Candidatus B. tachyglossi’ in archived B. concolor ticks and echidna skin. This is the first report of Borrelia in echidnas. Moreover, this study confirms its recent past presence in the Australian ecosystem. However, whether echidnas are a reservoir for this bacterium cannot be established at present, as it is unclear from these results if the presence of ‘Candidatus B. tachyglossi’ in echidna skin was due to transmission by an infected feeding tick or whether the tick obtained the bacterium from a bacteraemic echidna host. Therefore, whether ticks acquire this bacterium from infected echidnas remains to be confirmed through additional tick analyses and the examination of echidna blood samples. Nonetheless, the results generated in this study suggest that archived ticks harbour a preserved cache of bacterial communities that can offer new avenues for exploring past prevalence and the changes in microbiome composition over time.

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113

APPENDICES

114 Appendix A

Table A.1. Metadata spreadsheet of archived echidna ticks Sample Genus Species Life stage/Instar Location State Year Museum Host registration Other/description Tick registration ID # collected code code (if applicable) 1 Bothriocroton concolor Female Heathcote NSW 1993 WA T141308 Engorged n/a 2 Bothriocroton concolor Female Tooloom NSW 1961 ANIC 48001053 Grossly Engorged n/a 3 Bothriocroton concolor Nymph Tooloom NSW 1961 ANIC 48001053 Engorged n/a 4 Bothriocroton concolor Female Biddon NSW 1993 ANIC 48001072 Flattened - slight tear n/a 5 Bothriocroton concolor Female Biddon NSW 1993 ANIC 48001072 Flattened - slight tear n/a 6 Bothriocroton concolor Female Biddon NSW 1993 ANIC 48001072 Flattened - slight tear n/a 7 Bothriocroton concolor Male Biddon NSW 1993 ANIC 48001072 Normal n/a 8 Bothriocroton concolor Male Biddon NSW 1993 ANIC 48001072 Normal n/a 9 Bothriocroton concolor Male Biddon NSW 1993 ANIC 48001072 Normal n/a 10 Bothriocroton concolor Male Biddon NSW 1993 ANIC 48001072 Normal n/a 11 Bothriocroton concolor Male Batemans Bay NSW 1957 ANIC 48001049 Normal n/a 12 Bothriocroton concolor Male Batemans Bay NSW 1957 ANIC 48001049 Normal n/a 13 Bothriocroton concolor Nymph Batemans Bay NSW 1957 ANIC 48001049 Engorged n/a 14 Bothriocroton concolor Nymph Batemans Bay NSW 1957 ANIC 48001049 Normal n/a 15 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal n/a 16 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal n/a 17 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal n/a 18 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal n/a 19 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal n/a 20 Bothriocroton concolor Male Nanungo QLD 1968 ANIC 48000993 Normal n/a 21 Bothriocroton concolor Male Nanungo QLD 1968 ANIC 48000993 Normal n/a 22 Bothriocroton concolor Male Nanungo QLD 1968 ANIC 48000993 Normal n/a 23 Amblyomma triguttatum Male Bowen QLD 1997 ANIC 48000639 Normal n/a 24 Bothriocroton concolor Female Clermont QLD 1961 ANIC 48001055 Engorged n/a 25 Bothriocroton concolor Female Clermont QLD 1961 ANIC 48001055 Engorged n/a 26 Bothriocroton concolor Female Clermont QLD 1961 ANIC 48001055 Engorged n/a 27 Bothriocroton concolor Female Clermont QLD 1961 ANIC 48001055 Engorged n/a 28 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 29 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a

115 flattened 30 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 31 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 32 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 33 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 34 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 Normal - slightly n/a flattened 35 Bothriocroton concolor Nymph Uralla NSW 1958 ANIC 48001045 Engorged n/a 36 Bothriocroton concolor Female Uralla NSW 1958 ANIC 48001045 Engorged n/a 37 Bothriocroton concolor Female Uralla NSW 1958 ANIC 48001045 Engorged n/a 38 Bothriocroton concolor Nymph Uralla NSW 1958 ANIC 48001045 Engorged n/a 39 Bothriocroton concolor Nymph Uralla NSW 1958 ANIC 48001045 Normal n/a 40 Bothriocroton concolor Male Uralla NSW 1958 ANIC 48001045 Normal n/a 41 Bothriocroton concolor Male Uralla NSW 1958 ANIC 48001045 Normal n/a 42 Bothriocroton concolor Male Uralla NSW 1958 ANIC 48001045 Normal n/a 43 Bothriocroton concolor Male Uralla NSW 1958 ANIC 48001045 Normal n/a 44 Bothriocroton concolor Female Warwick QLD 1963 ANIC 48001057 Engorged n/a 45 Bothriocroton concolor Female Warwick QLD 1963 ANIC 48001057 Engorged n/a 46 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 47 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 48 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 49 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 50 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 51 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 52 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 53 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 54 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 55 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 56 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 57 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 58 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged n/a 59 Bothriocroton concolor Female Anakie QLD 1998 ANIC 48001065 Engorged n/a 60 Bothriocroton concolor Female Anakie QLD 1998 ANIC 48001065 Flattened n/a

116 61 Bothriocroton concolor Female Brisbane QLD 1954 ANIC 48001038 Engorged n/a 62 Bothriocroton concolor Female Brisbane QLD 1954 ANIC 48001038 Normal n/a 63 Bothriocroton concolor Female Brisbane QLD 1954 ANIC 48001038 Normal n/a 64 Bothriocroton concolor Female Boompa QLD 1948 ANIC 48001039 Normal n/a 65 Bothriocroton concolor Female Sydney NSW 1957 ANIC 48001048 Engorged n/a 66 Bothriocroton concolor Female Brisbane QLD 1941 ANIC 48001042 Normal n/a 67 Bothriocroton concolor Male Brisbane QLD 1941 ANIC 48001042 Normal n/a 68 Bothriocroton tachyglossi Female Woolooga QLD 2003 QLD 48001042 Grossly Engorged n/a 69 Bothriocroton tachyglossi Female Yepoon QLD 1967 QLD 588998 Grossly Engorged n/a 70 Bothriocroton tachyglossi Female Yepoon QLD 1967 QLD 588998 Engorged n/a 71 Bothriocroton tachyglossi Female Yepoon QLD 1967 QLD 588998 Engorged n/a 72 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Engorged n/a 73 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Grossly Engorged n/a 74 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Grossly Engorged n/a 75 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Grossly Engorged n/a 76 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Grossly Engorged n/a 77 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Grossly Engorged n/a 78 Bothriocroton concolor Male Gatton QLD 1954 ANIC 48001041 Normal n/a 79 Bothriocroton concolor Male Gatton QLD 1954 ANIC 48001041 Normal n/a 80 Bothriocroton concolor Female Gatton QLD 1954 ANIC 48001041 Engorged n/a 81 Bothriocroton concolor Female Gatton QLD 1954 ANIC 48001041 Engorged n/a 82 Bothriocroton concolor Male Queensland QLD 1971 QLD unregistered Normal n/a 83 Bothriocroton concolor Male Queensland QLD 1971 QLD unregistered Normal n/a 84 Bothriocroton concolor Female Maleny QLD 1954 QLD S21863 Grossly Engorged n/a 85 Bothriocroton concolor Nymph Maleny QLD 1954 QLD S21863 Engorged n/a 86 Bothriocroton concolor Nymph Maleny QLD 1954 QLD S21863 Normal n/a 87 Bothriocroton concolor Female Brisbane QLD no date QLD unregistered Engorged n/a 88 Bothriocroton concolor Female Brisbane QLD no date QLD unregistered Normal n/a 89 Bothriocroton concolor Male Brisbane QLD no date QLD unregistered Normal n/a 90 Bothriocroton concolor Female Whitlands VIC 1981 VIC 2016-17L Grossly Engorged n/a 91 Bothriocroton concolor Female Monbulk-Macelesfield VIC 1985 VIC 2016-17L Grossly Engorged n/a 92 Bothriocroton concolor Female Monbulk-Macelesfield VIC 1969 VIC 2016-17L Engorged n/a 93 Bothriocroton concolor Female Monbulk-Macelesfield VIC 1969 VIC 2016-17L Engorged n/a 94 Bothriocroton concolor Female Monbulk-Macelesfield VIC 1969 VIC 2016-17L Engorged n/a 95 Bothriocroton concolor Female Bathurst Teachers VIC 1966 VIC 2016-17L Grossly Engorged n/a College

117 96 Bothriocroton concolor Male Redlaud Bay QLD 1974 QLD S37045 Normal n/a 97 Bothriocroton concolor Male Redlaud Bay QLD 1974 QLD S37045 Normal n/a 98 Bothriocroton concolor Male Redlaud Bay QLD 1974 QLD S37045 Normal n/a 99 Bothriocroton concolor Male Redlaud Bay QLD 1974 QLD S37045 Normal n/a 100 Bothriocroton concolor Nymphs and larvae Redlaud Bay QLD 1974 QLD S37045 Normal n/a (numerous) 101 Bothriocroton concolor Female Rural Queensland QLD 1971 QLD S21871 Engorged n/a 102 Bothriocroton concolor Female Rural Queensland QLD 1971 QLD S21871 Engorged n/a 103 Bothriocroton concolor Female Rural Queensland QLD 1971 QLD S21871 Engorged n/a 104 Bothriocroton concolor Female Rural Queensland QLD 1971 QLD S21871 Engorged n/a 105 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 106 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 107 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 108 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 109 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 110 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 111 Bothriocroton concolor Nymph Rural Queensland QLD 1971 QLD S21871 Engorged n/a 112 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged - tear in n/a vetral side 113 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 114 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 115 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 116 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 117 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 118 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 119 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 120 Bothriocroton concolor Female Brisbane QLD 1942 QLD S37734 Engorged n/a 121 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 122 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 123 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 124 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 125 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 126 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal n/a 127 Bothriocroton concolor Female Mt. Abbott QLD 2007 QLD S37861 Normal n/a 128 Bothriocroton concolor Male Joyner QLD 2001 QLD S19939 Normal n/a 129 Bothriocroton concolor Female Maleny QLD 2000 QLD S15636 Engorged n/a

118 130 Bothriocroton concolor Female with eggs Maleny QLD 2000 QLD S19881 Engorged n/a 131 Bothriocroton concolor Female with eggs Maleny QLD 2000 QLD S19881 Engorged n/a 132 Bothriocroton concolor Larvae (numerous) Maleny QLD 2000 QLD S15629 n/a 133 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Normal n/a 134 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 135 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 136 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 137 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 138 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 139 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 140 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 141 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Engorged n/a 142 Bothriocroton concolor Male Mt. Ommaney QLD 2001 QLD S19459 Normal n/a 143 Bothriocroton concolor Male Mt. Ommaney QLD 2001 QLD S19459 Normal n/a 144 Bothriocroton concolor Male Mt. Ommaney QLD 2001 QLD S19459 Normal n/a 145 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S33975 Engorged n/a 146 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 147 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 148 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 149 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 150 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 151 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 152 Bothriocroton concolor Male Pinjarra Hills QLD 1997 QLD S19479 Normal n/a 153 Bothriocroton concolor Nymph (numerous) Pinjarra Hills QLD 1997 QLD S19479 Normal - some appear n/a engorged 154 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S26504 Engorged n/a 155 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S26504 Engorged n/a 156 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S26504 Normal n/a 157 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S26504 Normal n/a 158 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S33702 Engorged n/a 159 Amblyomma australiense Nymph Mt. Isa QLD 2002 QLD S33702 Normal n/a 160 Amblyomma australiense Male Mt. Isa QLD 2002 QLD S33702 Normal n/a 161 Amblyomma australiense Male Mt. Isa QLD 2002 QLD S33702 Normal n/a 162 Amblyomma australiense Male Mt. Isa QLD 2002 QLD S33702 Normal n/a 163 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 164 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a

119 165 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 166 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 167 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 168 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 169 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 170 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 171 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 172 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 173 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 174 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 175 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 176 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged n/a 177 Bothriocroton concolor Male Brisbane QLD 1971 QLD S37499 Normal n/a 178 Bothriocroton concolor Male Brisbane QLD 1971 QLD S37499 Normal n/a 179 Bothriocroton concolor Male Brisbane QLD 1971 QLD S37499 Normal n/a 180 Bothriocroton concolor Male Brisbane QLD 1971 QLD S37499 Normal n/a 181 Bothriocroton concolor Female Queensland QLD 1940 QLD unregistered Engorged n/a 182 Bothriocroton concolor Female Brisbane QLD 1959 QLD S21846 Engorged n/a 183 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 184 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 185 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 186 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 187 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 188 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 189 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged n/a 190 Bothriocroton concolor Male (Half) Kangaroo Island SA no date QLD S90972 Normal - half a tick n/a 191 Bothriocroton concolor Female Dayboro QLD 2006 QLD S73748 Engorged n/a 192 Bothriocroton concolor Female Dayboro QLD 2006 QLD S73748 Engorged n/a 193 Bothriocroton concolor Female Samford QLD 1938 QLD S21875 Normal n/a 194 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a 195 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a 196 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a 197 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a 198 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a 199 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal n/a

120 200 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Engorged n/a Arm 201 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 202 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 203 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 204 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 205 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 206 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 207 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 208 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 209 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 210 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 211 Bothriocroton concolor Nymph Running Creek, North QLD 2013 QLD S96574 Normal n/a Arm 212 Bothriocroton concolor Female Buccan QLD 2011 QLD S32257 Engorged n/a 213 Bothriocroton concolor Female Buccan QLD 2011 QLD S32257 Engorged n/a 214 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 215 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 216 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 217 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 218 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 219 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 220 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 221 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 222 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 223 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 224 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 225 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 226 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 227 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a

121 228 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 229 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal n/a 230 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Engorged n/a 231 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal n/a 232 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal n/a 233 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal n/a 234 Bothriocroton concolor Male Sethebrook QLD 1963 QLD S73159 Normal n/a 235 Bothriocroton concolor Male Sethebrook QLD 1963 QLD S73159 Normal n/a 236 Bothriocroton concolor Male Maleny QLD 2009 QLD S28705 Normal n/a 237 Bothriocroton concolor Male Maleny QLD 2009 QLD S28705 Normal n/a 238 Bothriocroton concolor Male Maleny QLD 2009 QLD S28705 Normal n/a 239 Bothriocroton concolor Male Boggom QLD 1996 QLD S73147 Normal n/a 240 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 241 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 242 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 243 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 244 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 245 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 246 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 247 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 248 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal n/a 249 Bothriocroton concolor Male Nerang QLD 1964 QLD S73036 Normal n/a 250 Bothriocroton concolor Male Nerang QLD 1964 QLD S73036 Normal n/a 251 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Flattened n/a 252 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 253 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 254 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 255 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 256 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 257 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 258 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 259 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 260 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 261 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 262 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a

122 263 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 264 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 265 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 266 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 267 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal n/a 268 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged n/a 269 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged n/a 270 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged n/a 271 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal n/a 272 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal n/a 273 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal n/a 274 Bothriocroton concolor Female Parkurst, Rockhampton QLD 1994 QLD S60871 Grossly Engorged n/a 275 Bothriocroton concolor Female Parkurst, Rockhampton QLD 1994 QLD S60871 Engorged n/a 276 Bothriocroton concolor Female Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 277 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 278 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 279 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 280 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 281 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 282 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 283 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 284 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 285 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 286 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 287 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 288 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 289 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 290 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 291 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 292 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 293 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 294 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 295 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 296 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 297 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a

123 298 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 299 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 300 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 301 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 302 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 303 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 304 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 305 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 306 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 307 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 308 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 309 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 310 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 311 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 312 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 313 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 314 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 315 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 316 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 317 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 318 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 319 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 320 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 321 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 322 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 323 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 324 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 325 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 326 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 327 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal n/a 328 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 329 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 330 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna

124 331 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 332 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 333 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 334 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 335 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 Engorged - From sick n/a echidna 336 Bothriocroton concolor Male Maleny QLD 2000 QLD S45426 Normal - From sick n/a echidna 337 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 338 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 339 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 340 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 341 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 342 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 343 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 344 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged n/a 345 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged n/a 346 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged n/a 347 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged n/a 348 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged n/a 349 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged n/a 350 Bothriocroton concolor Male Ormiston QLD 1964 QLD S73166 Normal n/a 351 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 352 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 353 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 354 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 355 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 356 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 357 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 358 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 359 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 360 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 361 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 362 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a

125 363 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 364 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 365 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 366 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 367 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 368 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 369 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 370 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 371 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 372 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged n/a 373 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Normal n/a 374 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 375 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 376 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 377 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 378 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 379 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 380 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 381 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 382 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 383 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 384 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 385 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 386 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 387 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 388 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 389 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 390 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 391 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 392 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 393 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 394 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 395 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 396 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 397 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a

126 398 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 399 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 400 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 401 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 402 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 403 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 404 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 405 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 406 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 407 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 408 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 409 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 410 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 411 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 412 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 413 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 414 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 415 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 416 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 417 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 418 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 419 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged n/a 420 Bothriocroton concolor Male Brisbane QLD 1971 QLD S21871 Normal n/a 421 Bothriocroton concolor Male Brisbane QLD 1971 QLD S21871 Normal n/a 422 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Grossly Engorged n/a Gairdner/Torrens Basin 423 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Grossly Engorged n/a Gairdner/Torrens Basin 424 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 425 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 426 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 427 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 428 Amblyomma triguttatum Female Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin

127 429 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 430 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 431 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 432 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 433 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Engorged n/a Gairdner/Torrens Basin 434 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 435 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 436 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 437 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 438 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 439 Amblyomma triguttatum Male Lake Hart, SA 1978 SA J7608-26 Normal n/a Gairdner/Torrens Basin 440 Bothriocroton concolor Female Straun, Lower South SA 1981 SA J7743-4 Normal n/a East 441 Bothriocroton concolor Female Straun, Lower South SA 1981 SA J7743-4 Normal n/a East 442 Bothriocroton concolor Female Snug Cove, Kangaroo SA 1988 SA J7758 Engorged n/a Island 443 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 444 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 445 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 446 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 447 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 448 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 449 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 450 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a

128 engorgement 451 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 452 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 453 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 454 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 455 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 456 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 457 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 458 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 459 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 460 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 461 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 462 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 463 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 464 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 465 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 466 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 467 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 468 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 469 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 470 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 471 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement

129 472 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 473 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 474 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 475 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 476 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 477 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 478 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 479 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 480 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 481 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 482 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 483 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 484 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 485 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 486 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 487 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 488 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 489 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 490 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 491 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 492 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 493 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a

130 engorgement 494 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 495 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 496 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 497 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 498 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 499 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 500 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 501 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 502 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 503 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 504 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 505 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 506 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 507 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 508 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 509 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 510 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 511 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 512 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 513 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 514 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement

131 515 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 516 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 517 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 518 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 519 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 520 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 521 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 522 Bothriocroton concolor Nymph Healesville Sanctuary SA 1989 SA J7771-5 Various degrees of n/a engorgement 523 Bothriocroton concolor Nymphs (numerous) Healesville Sanctuary SA 1989 SA J7770 Various degrees of n/a engorgement 524 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 525 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 526 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 527 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 528 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 529 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 530 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 531 Bothriocroton concolor Male Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 532 Bothriocroton concolor Nymph Birdwood, Adelaide, Mt SA 1979 SA J7760-8 Normal n/a Lofty Ranges 533 Bothriocroton concolor Female Bridgewater, Mt Lofty SA 1989 SA J7769 Engorged n/a Ranges 534 Bothriocroton concolor Female South Australia (no SA 1973 SA J7782-3 Engorged n/a locality data) 535 Bothriocroton concolor Male South Australia (no SA 1973 SA J7782-3 Normal n/a locality data) 536 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a

132 537 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a 538 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a 539 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a 540 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a 541 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged n/a 542 Bothriocroton concolor Male Victoria VIC 1974 SA J1798 Normal n/a 543 Bothriocroton concolor Female Victoria, Nowa Nowa VIC 1975 SA J1895 Grossly Engorged n/a 544 Bothriocroton concolor Female Victoria, Nowa Nowa VIC 1975 SA J1895 Grossly Engorged n/a 545 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 546 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 547 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 548 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 549 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 550 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 551 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 552 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 553 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 554 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged n/a 555 Bothriocroton concolor Male Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 556 Bothriocroton concolor Male Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 557 Bothriocroton concolor Male Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 558 Bothriocroton concolor Male Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 559 Bothriocroton concolor Female Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 560 Bothriocroton concolor Female Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 561 Bothriocroton concolor Female Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 562 Bothriocroton concolor Female Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 563 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 564 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 565 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a

133 East echidna 566 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 567 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 568 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 569 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 570 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 571 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 572 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 573 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 574 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 575 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 576 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 577 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 578 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 579 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 580 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 581 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 582 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 583 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 584 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 585 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 586 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna

134 587 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 588 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 589 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 590 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 591 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 592 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 593 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 594 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 595 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 596 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 597 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 598 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 599 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 600 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 601 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 602 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 603 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 604 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 605 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 606 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 607 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 608 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a

135 East echidna 609 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 610 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 611 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 612 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 613 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 614 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 615 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 616 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 617 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 618 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 619 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 620 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 621 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 622 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 623 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 624 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 625 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 626 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 627 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 628 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 629 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna

136 630 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 631 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 632 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 633 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 634 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 635 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 636 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 637 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 638 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 639 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 640 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 641 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 642 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 643 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 644 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 645 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 646 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 647 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 648 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 649 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 650 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 651 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a

137 East echidna 652 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 653 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 654 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 655 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 656 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 657 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 658 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 659 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 660 Bothriocroton concolor Nymph Millicent, Lower South SA 1981 SA J7745-57 Normal - from sick n/a East echidna 661 Ixodes tasmani Female Fingal TAS 1997 ANIC 48003382 Engorged n/a 662 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Engorged n/a 663 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Engorged n/a 664 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Normal n/a 665 Ixodes tasmani Female Maydena TAS 1959 ANIC 48003458 Normal n/a 666 Ixodes tasmani Female Maydena TAS 1959 ANIC 48003458 Normal n/a 667 Amblyomma australiense Female Bowen QLD 1928 ANIC 48000315 Grossly Engorged n/a 668 Bothriocroton hydrosauri Male Rockhampton QLD 1963 ANIC 48001159 Normal n/a 669 Bothriocroton hydrosauri Male Rockhampton QLD 1963 ANIC 48001159 Normal n/a 670 Amblyomma echidnae Male Townsville QLD 1979 ANIC 48000367 Normal n/a 671 Bothriocroton tachyglossi Larvae (numerous) Boompa QLD 1948 ANIC 48001176 Normal - bred from n/a paratype 672 Bothriocroton tachyglossi Larvae (numerous) Rockhampton QLD no date ANIC 48001182 Normal n/a 673 Bothriocroton tachyglossi Larvae (numerous) Warwick QLD 1937 ANIC 48001171 Normal n/a 674 Ixodes tasmani Larva Scamander TAS 1996 ANIC 48003343 Normal n/a 675 Ixodes tasmani Larva Scamander TAS 1996 ANIC 48003343 Normal n/a 676 Ixodes tasmani Nymph Scamander TAS 1996 ANIC 48003343 Normal n/a 677 Ixodes tasmani Nymph Scamander TAS 1996 ANIC 48003343 Normal n/a 678 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal n/a 679 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal n/a

138 680 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal n/a 681 Amblyomma australiense Female Darwin NT 1974 ANIC 48000312 Engorged n/a 682 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal n/a 683 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal n/a 684 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal n/a 685 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal n/a 686 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal n/a 687 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged n/a 688 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged n/a 689 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged n/a 690 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Normal n/a 691 Bothriocroton hydrosauri Female Rockhampton QLD 1966 ANIC 48001160 Engorged n/a 692 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged - missing n/a head 693 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged - missing n/a head 694 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged - missing n/a head 695 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 696 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 697 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 698 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 699 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 700 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 701 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 702 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Engorged n/a 703 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 704 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 705 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 706 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 707 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 708 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 709 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 710 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 711 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 712 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 713 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal n/a 139 Sample ID #* 714 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 715 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Engorged 1 Echidna 716 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 717 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 2Ta 718 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 1 Normal 3Ta 719 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 3Ta 720 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 1 Slightly engorged 4Ta 721 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 1 Engorged 4Ta 722 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 723 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 724 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Engorged 4Ta 725 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 726 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 11Ta 727 Bothriocroton concolor Nymph Forestdale QLD no date QLD 2 Normal 11Ta 728 Bothriocroton concolor Nymph Forestdale QLD no date QLD 2 Normal 11Ta 729 Bothriocroton concolor Nymph Forestdale QLD no date QLD 2 Engorged 12Ta 730 Bothriocroton concolor Female Forestdale QLD no date QLD 2 Grossly engorged 13Ta 731 Bothriocroton concolor Female Forestdale QLD no date QLD 2 Grossly engorged 14Ta 732 Bothriocroton concolor Female Forestdale QLD no date QLD 2 Grossly engorged 14Ta 733 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 734 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 735 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Engorged 22Ta** 736 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Engorged 23Ta** 737 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Engorged 24Ta 738 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Slightly engorged 24Ta 739 Bothriocroton concolor Male Karana Downs QLD 2007 QLD 3 Normal 24Ta 740 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 741 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 742 Bothriocroton concolor Nymph Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 743 Bothriocroton concolor Nymph Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 744 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 745 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 746 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 747 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta

140 748 Bothriocroton concolor Nymph Ipswich QLD 2011 QLD 5 Normal 41Ta 749 Bothriocroton concolor Nymph Ipswich QLD 2011 QLD 5 Normal 41Ta 750 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 43Ta** 751 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 43Ta** 752 Bothriocroton concolor Female Highvale QLD 2006 QLD 6 Engorged 51Ta 753 Bothriocroton concolor Female Highvale QLD 2006 QLD 6 Slightly engorged 51Ta 754 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 51Ta 755 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 51Ta 756 Bothriocroton concolor Female Malanda QLD 2003 QLD 7 Slightly engorged 61Ta 757 Bothriocroton concolor Nymph Malanda QLD 2003 QLD 7 Normal 61Ta 758 Bothriocroton concolor Nymph Malanda QLD 2003 QLD 7 Normal 61Ta 759 Bothriocroton concolor Larva Malanda QLD 2003 QLD 7 Normal 61Ta 760 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 761 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 762 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 763 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 764 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 765 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 766 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 767 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 768 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 769 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 770 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 771 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 772 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 773 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 774 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 1 Echidna 775 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 1 Engorged 4Ta 776 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 777 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 778 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 779 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 780 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 781 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 782 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta

141 783 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 784 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 785 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 786 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 787 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 788 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 789 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 790 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Normal 4Ta 791 Bothriocroton concolor Female Forestdale QLD no date QLD 2 Engorged 14Ta 792 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 793 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 794 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 795 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 796 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 797 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 798 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 799 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 800 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 801 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 802 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 803 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 804 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 805 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 806 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 807 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 808 Bothriocroton concolor Male Forestdale QLD no date QLD 2 Normal 14Ta 809 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Normal 24Ta 810 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 3 Normal 24Ta 811 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 812 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 813 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 814 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 815 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 816 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 817 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta

142 818 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 819 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 820 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 4 Normal 31Ta 821 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 822 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 823 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 824 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Engorged 41Ta 825 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 826 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 827 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 828 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 829 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 830 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 831 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 832 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 833 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 834 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 835 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 836 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 837 Bothriocroton concolor Female Ipswich QLD 2011 QLD 5 Normal 41Ta 838 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 839 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 840 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 841 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 842 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 843 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 844 Bothriocroton concolor Male Ipswich QLD 2011 QLD 5 Normal 41Ta 845 Bothriocroton concolor Nymph Ipswich QLD 2011 QLD 5 Normal 41Ta 846 Bothriocroton concolor Female Highvale QLD 2006 QLD 6 Engorged 51Ta 847 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 51Ta 848 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 51Ta 849 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 52Ta** 850 Bothriocroton concolor Male Highvale QLD 2006 QLD 6 Normal 52Ta** * Ticks removed from Queensland Museum archived echidna specimens with corresponding tissue samples (see Appendix A, Table A.3)

** Ticks removed with a skin biopsy (see Appendix A, Table A.3)

143 Table A.2. Sub-sample of archived echidna ticks for DNA extraction Host Archived Extraction ID Sample Life stage/ Registration Other/ Echidna # ID # Genus Species Instar Location State Year collected Museum code Description specimen # * e1 714 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 1 Echidna Normal 1 e2 715 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Echidna Engorged 1 e3 716 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 1 Echidna Normal 1 e4 717 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 2Ta Normal 1 e5 718 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 3Ta Normal 1 e6 719 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 3Ta Normal 1 Slightly e7 720 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 4Ta engorged 1 e8 721 Bothriocroton concolor Female South of Mooloolah QLD 1992 QLD 4Ta Engorged 1 e9 722 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 4Ta Normal 1 e10 723 Bothriocroton concolor Male South of Mooloolah QLD 1992 QLD 4Ta Normal 1 e11 724 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 4Ta Engorged 1 e12 725 Bothriocroton concolor Nymph South of Mooloolah QLD 1992 QLD 4Ta Normal 1 e13 726 Bothriocroton concolor Male Forestdale QLD no date QLD 11Ta Normal 2 e14 727 Bothriocroton concolor Nymph Forestdale QLD no date QLD 11Ta Normal 2 e15 728 Bothriocroton concolor Nymph Forestdale QLD no date QLD 11Ta Normal 2 e16 729 Bothriocroton concolor Nymph Forestdale QLD no date QLD 12Ta Engorged 2 Grossly e17 730 Bothriocroton concolor Female Forestdale QLD no date QLD 13Ta engorged 2 Grossly e18 731 Bothriocroton concolor Female Forestdale QLD no date QLD 14Ta engorged 2 Grossly e19 732 Bothriocroton concolor Female Forestdale QLD no date QLD 14Ta engorged 2 e20 733 Bothriocroton concolor Male Forestdale QLD no date QLD 14Ta Normal 2 e21 734 Bothriocroton concolor Male Forestdale QLD no date QLD 14Ta Normal 2 e22 735 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 22Ta** Engorged 3 e23 736 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 23Ta** Engorged 3

144 e24 737 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 24Ta Engorged 3 Slightly e25 738 Bothriocroton concolor Female Karana Downs QLD 2007 QLD 24Ta engorged 3 e26 739 Bothriocroton concolor Male Karana Downs QLD 2007 QLD 24Ta Normal 3 e27 740 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 31Ta Normal 4 e28 741 Bothriocroton concolor Male Queen Mary Falls QLD 2004 QLD 31Ta Normal 4 e29 742 Bothriocroton concolor Nymph Queen Mary Falls QLD 2004 QLD 31Ta Normal 4 e30 743 Bothriocroton concolor Nymph Queen Mary Falls QLD 2004 QLD 31Ta Normal 4 e31 744 Bothriocroton concolor Female Ipswich QLD 2011 QLD 41Ta Engorged 5 e32 745 Bothriocroton concolor Female Ipswich QLD 2011 QLD 41Ta Engorged 5 e33 746 Bothriocroton concolor Male Ipswich QLD 2011 QLD 41Ta Normal 5 e34 747 Bothriocroton concolor Male Ipswich QLD 2011 QLD 41Ta Normal 5 e35 748 Bothriocroton concolor Nymph Ipswich QLD 2011 QLD 41Ta Normal 5 e36 749 Bothriocroton concolor Nymph Ipswich QLD 2011 QLD 41Ta Normal 5 e37 750 Bothriocroton concolor Female Ipswich QLD 2011 QLD 43Ta** Engorged 5 e38 751 Bothriocroton concolor Male Ipswich QLD 2011 QLD 43Ta** Normal 5 e39 752 Bothriocroton concolor Female Highvale QLD 2006 QLD 51Ta Engorged 6 Slightly e40 753 Bothriocroton concolor Female Highvale QLD 2006 QLD 51Ta engorged 6 e41 754 Bothriocroton concolor Male Highvale QLD 2006 QLD 51Ta Normal 6 e42 755 Bothriocroton concolor Male Highvale QLD 2006 QLD 51Ta Normal 6 Slightly e43 756 Bothriocroton concolor Female Malanda QLD 2003 QLD 61Ta engorged 7 e44 757 Bothriocroton concolor Nymph Malanda QLD 2003 QLD 61Ta Normal 7 e45 758 Bothriocroton concolor Nymph Malanda QLD 2003 QLD 61Ta Normal 7 e46 759 Bothriocroton concolor Larva Malanda QLD 2003 QLD 61Ta Normal 7

Grossly n 1 2 Bothriocroton concolor Female Tooloom NSW 1961 ANIC 48001053 Engorged /a n 2 3 Bothriocroton concolor Nymph Tooloom NSW 1961 ANIC 48001053 Engorged /a

145 Flattened - slight n 3 4 Bothriocroton concolor Female Biddon NSW 1993 ANIC 48001072 tear /a n 4 7 Bothriocroton concolor Male Biddon NSW 1993 ANIC 48001072 Normal /a n 5 11 Bothriocroton concolor Male Batemans Bay NSW 1957 ANIC 48001049 Normal /a n 6 13 Bothriocroton concolor Nymph Batemans Bay NSW 1957 ANIC 48001049 Engorged /a n 6 14 Bothriocroton concolor Nymph Batemans Bay NSW 1957 ANIC 48001049 Normal /a n 7 15 Bothriocroton concolor Female Nanungo QLD 1968 ANIC 48000993 Normal /a n 8 20 Bothriocroton concolor Male Nanungo QLD 1968 ANIC 48000993 Normal /a n 9 23 Amblyomma triguttatum Male Bowen QLD 1997 ANIC 48000639 Normal /a n 10 24 Bothriocroton concolor Female Clermont QLD 1961 ANIC 48001055 Engorged /a Normal - slightly n 11 28 Bothriocroton concolor Male Clermont QLD 1961 ANIC 48001055 flattened /a n 12 35 Bothriocroton concolor Nymph Uralla NSW 1958 ANIC 48001045 Engorged /a n 13 36 Bothriocroton concolor Female Uralla NSW 1958 ANIC 48001045 Engorged /a n 14 40 Bothriocroton concolor Male Uralla NSW 1958 ANIC 48001045 Normal /a n 15 66 Bothriocroton concolor Female Brisbane QLD 1941 ANIC 48001042 Normal /a n 16 67 Bothriocroton concolor Male Brisbane QLD 1941 ANIC 48001042 Normal /a n 17 61 Bothriocroton concolor Female Brisbane QLD 1954 ANIC 48001038 Engorged /a Grossly n 18 69 Bothriocroton tachyglossi Female Yepoon QLD 1967 QLD 588998 Engorged /a n 19 72 Bothriocroton tachyglossi Female Cinnabar Goomeri QLD 1988 QLD 590000 Engorged /a n 20 78 Bothriocroton concolor Male Gatton QLD 1954 ANIC 48001041 Normal /a n 21 80 Bothriocroton concolor Female Gatton QLD 1954 ANIC 48001041 Engorged /a n 22 64 Bothriocroton concolor Female Boompa QLD 1948 ANIC 48001039 Normal /a n 23 85 Bothriocroton concolor Nymph Maleny QLD 1954 QLD S21863 Engorged /a

146 n 23 86 Bothriocroton concolor Nymph Maleny QLD 1954 QLD S21863 Normal /a n 24 87 Bothriocroton concolor Female Brisbane QLD no date QLD unregistered Engorged /a n 25 89 Bothriocroton concolor Male Brisbane QLD no date QLD unregistered Normal /a n 26 92 Bothriocroton concolor Female Monbulk-Macelesfield VIC 1969 VIC 2016-17L Engorged /a n 27 96 Bothriocroton concolor Male Redlaud Bay QLD 1974 QLD S37045 Normal /a n 28 101 Bothriocroton concolor Female Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 105 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 106 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 107 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 108 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 109 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 110 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 29 111 Bothriocroton concolor Nymph Rural QLD QLD 1971 QLD S21871 Engorged /a n 30 121 Bothriocroton concolor Male Mt. Abbott QLD 2007 QLD S37861 Normal /a n 31 127 Bothriocroton concolor Female Mt. Abbott QLD 2007 QLD S37861 Normal /a n 32 133 Bothriocroton concolor Female Mt. Ommaney QLD 2001 QLD S19459 Normal /a n 33 142 Bothriocroton concolor Male Mt. Ommaney QLD 2001 QLD S19459 Normal /a n 34 145 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S33975 Engorged /a n 35 154 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S26504 Engorged /a n 36 158 Amblyomma australiense Female Mt. Isa QLD 2002 QLD S33702 Engorged /a n 37 159 Amblyomma australiense Nymph Mt. Isa QLD 2002 QLD S33702 Normal /a n 38 160 Amblyomma australiense Male Mt. Isa QLD 2002 QLD S33702 Normal /a

147 n 39 163 Bothriocroton concolor Female Brisbane QLD 1971 QLD S37499 Engorged /a n 40 177 Bothriocroton concolor Male Brisbane QLD 1971 QLD S37499 Normal /a n 41 182 Bothriocroton concolor Female Brisbane QLD 1959 QLD S21846 Engorged /a n 42 183 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 184 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 185 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 186 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 187 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 188 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 42 189 Bothriocroton concolor Nymph Brisbane QLD 1959 QLD S21846 Engorged /a n 43 193 Bothriocroton concolor Female Samford QLD 1938 QLD S21875 Normal /a n 44 194 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal /a n 44 195 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal /a n 44 196 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal /a n 44 197 Bothriocroton concolor Nymph Samford QLD 1938 QLD S21875 Normal /a n 45 212 Bothriocroton concolor Female Buccan QLD 2011 QLD S32257 Engorged /a n 46 214 Bothriocroton concolor Male Buccan QLD 2011 QLD S32257 Normal /a n 47 230 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Engorged /a n 47 231 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal /a n 47 232 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal /a n 47 233 Bothriocroton concolor Nymph Buccan QLD 2011 QLD S32257 Normal /a n 48 240 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a

148 n 48 241 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 242 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 243 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 244 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 245 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 246 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 247 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 48 248 Bothriocroton concolor Nymph Nerang QLD 1964 QLD S73036 Normal /a n 49 249 Bothriocroton concolor Male Nerang QLD 1964 QLD S73036 Normal /a n 50 252 Bothriocroton concolor Male Port Stewart QLD 1943 QLD S21858 Normal /a n 51 268 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged /a n 51 269 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged /a n 51 270 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Engorged /a n 51 271 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal /a n 51 272 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal /a n 51 273 Bothriocroton concolor Nymph Port Stewart QLD 1943 QLD S21858 Normal /a n 52 276 Bothriocroton concolor Female Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal /a n 53 277 Bothriocroton concolor Male Chinchilla, Gracehithgo QLD 1985 QLD S69926 Normal /a Engorged - From n 54 328 Bothriocroton concolor Female Maleny QLD 2000 QLD S45426 sick echidna /a Normal - From n 55 336 Bothriocroton concolor Male Maleny QLD 2000 QLD S45426 sick echidna /a n 56 337 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 338 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a

149 n 56 339 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 340 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 341 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 342 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 343 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 56 344 Bothriocroton concolor Nymph Ormiston QLD 1964 QLD S73166 Engorged /a n 57 345 Bothriocroton concolor Female Ormiston QLD 1964 QLD S73166 Engorged /a n 58 350 Bothriocroton concolor Male Ormiston QLD 1964 QLD S73166 Normal /a n 59 351 Bothriocroton concolor Female Brisbane QLD 1971 QLD S21871 Engorged /a n 60 374 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 375 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 376 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 377 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 378 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 379 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 380 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 381 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 382 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a n 60 383 Bothriocroton concolor Nymph Brisbane QLD 1971 QLD S21871 Engorged /a Lake Hart, Gairdner/Torrens Grossly n 61 422 Amblyomma triguttatum Female Basin SA 1978 SA J7608-26 Engorged /a Lake Hart, Gairdner/Torrens n 62 429 Amblyomma triguttatum Male Basin SA 1978 SA J7608-26 Engorged /a

150 Birdwood, Adelaide, Mt n 63 524 Bothriocroton concolor Male Lofty Ranges SA 1979 SA J7760-8 Normal /a Birdwood, Adelaide, Mt n 64 532 Bothriocroton concolor Nymph Lofty Ranges SA 1979 SA J7760-8 Normal /a South Australia (no n 65 534 Bothriocroton concolor Female locality data) SA 1973 SA J7782-3 Engorged /a South Australia (no n 66 535 Bothriocroton concolor Male locality data) SA 1973 SA J7782-3 Normal /a n 67 536 Bothriocroton concolor Female Victoria VIC 1974 SA J1798 Engorged /a n 68 542 Bothriocroton concolor Male Victoria VIC 1974 SA J1798 Normal /a Grossly n 69 543 Bothriocroton concolor Female Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a n 70 545 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a n 70 546 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a n 70 547 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a n 70 548 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a n 70 549 Bothriocroton concolor Nymph Victoria, Nowa Nowa VIC 1975 SA J1895 Engorged /a Millicent, Lower South Normal - from n 71 555 Bothriocroton concolor Male East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 72 559 Bothriocroton concolor Female East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 73 563 Bothriocroton concolor Nymph East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 73 564 Bothriocroton concolor Nymph East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 73 565 Bothriocroton concolor Nymph East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 73 566 Bothriocroton concolor Nymph East SA 1981 SA J7745-57 sick echidna /a Millicent, Lower South Normal - from n 73 567 Bothriocroton concolor Nymph East SA 1981 SA J7745-57 sick echidna /a n 74 661 Ixodes tasmani Female Fingal TAS 1997 ANIC 48003382 Engorged /a n 75 662 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Engorged /a n 76 663 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Engorged /a

151 n 77 664 Ixodes tasmani Female Antill Ponds TAS 1960 ANIC 48003473 Normal /a n 78 665 Ixodes tasmani Female Maydena TAS 1959 ANIC 48003458 Normal /a n 79 666 Ixodes tasmani Female Maydena TAS 1959 ANIC 48003458 Normal /a n 80 668 Bothriocroton hydrosauri Male Rockhampton QLD 1963 ANIC 48001159 Normal /a n 81 669 Bothriocroton hydrosauri Male Rockhampton QLD 1963 ANIC 48001159 Normal /a n 82 670 Amblyomma echidnae Male Townsville QLD 1979 ANIC 48000367 Normal /a n 83 676 Ixodes tasmani Nymph Scamander TAS 1996 ANIC 48003343 Normal /a n 83 677 Ixodes tasmani Nymph Scamander TAS 1996 ANIC 48003343 Normal /a n 84 678 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal /a n 85 679 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal /a n 86 680 Amblyomma australiense Male Darwin NT 1974 ANIC 48000312 Normal /a n 87 681 Amblyomma australiense Female Darwin NT 1974 ANIC 48000312 Engorged /a n 88 682 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal /a n 89 683 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal /a n 90 684 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal /a n 91 685 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal /a n 92 686 Bothriocroton hydrosauri Male Rockhampton QLD 1966 ANIC 48001160 Normal /a n 93 687 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged /a n 93 688 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged /a n 93 689 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Engorged /a n 93 690 Bothriocroton hydrosauri Nymph Rockhampton QLD 1966 ANIC 48001160 Normal /a n 94 691 Bothriocroton hydrosauri Female Rockhampton QLD 1966 ANIC 48001160 Engorged /a

152 Engorged - n 95 692 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 missing head /a Engorged - n 96 693 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 missing head /a Engorged - n 97 694 Amblyomma fimbriatum Female Justa Creek, China Wall NT 1967 ANIC 48000376 missing head /a n 98 704 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal /a n 99 705 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal /a n 100 706 Amblyomma fimbriatum Male Justa Creek, China Wall NT 1967 ANIC 48000376 Normal /a n 101 1 Bothriocroton concolor Female Heathcote NSW 1993 WA T141308 Engorged /a n 102 44 Bothriocroton concolor Female Warwick QLD 1963 ANIC 48001057 Engorged /a n 103 45 Bothriocroton concolor Female Warwick QLD 1963 ANIC 48001057 Engorged /a n 104 46 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged /a n 105 47 Bothriocroton concolor Female Tooloom NSW 1965 ANIC 4800106 Engorged /a n 106 59 Bothriocroton concolor Female Anakie QLD 1998 ANIC 48001065 Engorged /a n 107 60 Bothriocroton concolor Female Anakie QLD 1998 ANIC 48001065 Flattened /a n 108 191 Bothriocroton concolor Female Dayboro QLD 2006 QLD S73748 Engorged /a n 109 192 Bothriocroton concolor Female Dayboro QLD 2006 QLD S73748 Engorged /a Straun, Lower South n 110 440 Bothriocroton concolor Female East SA 1981 SA J7743-4 Normal /a Straun, Lower South n 111 441 Bothriocroton concolor Female East SA 1981 SA J7743-4 Normal /a Snug Cove, Kangaroo n 112 442 Bothriocroton concolor Female Island SA 1988 SA J7758 Engorged /a Bridgewater, Mt Lofty n 113 533 Bothriocroton concolor Female Ranges SA 1989 SA J7769 Engorged /a Nymphs Various degrees n 114 523 Bothriocroton concolor (x4)*** Healesville Sanctuary SA 1989 SA J7770 of engorgement /a

* Ticks removed from Queensland Museum archived echidna specimens with corresponding tissue samples ( see Appendix A, Table A.3)

153 ** Tick removed with a skin biopsy (see Appendix A, Table A.3)

*** Four nymphs removed from a vial containing archived echidna tissue with the same identification number (see Appendix A, Table A.3)

Extracted ticks within boarder contain the same host registration number

Nymphal pool - where more than one nymph was used per extraction to increase DNA yield

154 Table A.3. Metadata spreadsheet of archived echidna tissue biopsies Host Specimen Specimen Registration Sample ID # Genus Species Registration Code type Code Location State Year collected Description* 2 Tachyglossus aculeatus 2E skin 1 South of Mooloolah QLD 1992 Skin tissue at 2Ta tick bite site 3 Tachyglossus aculeatus 3E skin 1 South of Mooloolah QLD 1992 Skin tissue at 3Ta tick bite site 5 Tachyglossus aculeatus 5 liver 1 South of Mooloolah QLD 1992 Liver biopsy 6 Tachyglossus aculeatus 6 spleen 1 South of Mooloolah QLD 1992 Spleen biopsy 7 Tachyglossus aculeatus 7 kidney 1 South of Mooloolah QLD 1992 Right kidney biopsy 8 Tachyglossus aculeatus 8 muscle 1 South of Mooloolah QLD 1992 Rectus abdominis muscle biopsy 12 Tachyglossus aculeatus 12 skin 2 Forestdale QLD no date Skin tissue at 12Ta tick bite site 13 Tachyglossus aculeatus 13 skin 2 Forestdale QLD no date Skin tissue at 13Ta tick bite site 15 Tachyglossus aculeatus 15 liver 2 Forestdale QLD no date Liver biopsy 16 Tachyglossus aculeatus 16 spleen 2 Forestdale QLD no date Spleen biopsy 17 Tachyglossus aculeatus 17 kidney 2 Forestdale QLD no date Right kidney biopsy 18 Tachyglossus aculeatus 18 muscle 2 Forestdale QLD no date Rectus abdominis muscle biopsy 22 Tachyglossus aculeatus 22 skin** 3 Karana Downs QLD 2007 Skin with tick biopsy 23 Tachyglossus aculeatus 23 skin** 3 Karana Downs QLD 2007 Skin with tick biopsy 25 Tachyglossus aculeatus 25 liver 3 Karana Downs QLD 2007 Liver biopsy 26 Tachyglossus aculeatus 26 spleen 3 Karana Downs QLD 2007 Spleen biopsy 27 Tachyglossus aculeatus 27 kidney 3 Karana Downs QLD 2007 Right kidney biopsy 28 Tachyglossus aculeatus 28 muscle 3 Karana Downs QLD 2007 Rectus abdominis muscle biopsy 35 Tachyglossus aculeatus 35 liver 4 Queen Mary Falls QLD 2004 Liver biopsy 36 Tachyglossus aculeatus 36 spleen 4 Queen Mary Falls QLD 2004 Spleen biopsy 37 Tachyglossus aculeatus 37 kidney 4 Queen Mary Falls QLD 2004 Right kidney biopsy 38 Tachyglossus aculeatus 38 muscle 4 Queen Mary Falls QLD 2004 Rectus abdominis muscle biopsy 43 Tachyglossus aculeatus 43 skin** 5 Ipswich QLD 2011 Skin and tick biopsy (male and female) 45 Tachyglossus aculeatus 45 liver 5 Ipswich QLD 2011 Liver biopsy

155 46 Tachyglossus aculeatus 46 spleen 5 Ipswich QLD 2011 Spleen biopsy 47 Tachyglossus aculeatus 47 kidney 5 Ipswich QLD 2011 Right kidney biopsy 48 Tachyglossus aculeatus 48 muscle 5 Ipswich QLD 2011 Rectus abdominis muscle biopsy 52 Tachyglossus aculeatus 52 skin** 6 Highvale QLD 2006 Skin with tick biopsy 58 Tachyglossus aculeatus 58 muscle 6 Highvale QLD 2006 Rectus abdominis muscle biopsy 65 Tachyglossus aculeatus 65 liver 7 Malanda QLD 2003 Liver biopsy 66 Tachyglossus aculeatus 66 spleen 7 Malanda QLD 2003 Spleen biopsy 67 Tachyglossus aculeatus 67 kidney 7 Malanda QLD 2003 Right kidney biopsy

68 Tachyglossus aculeatus 68 muscle 7 Malanda QLD 2003 Rectus abdominis muscle biopsy

5 523 Tachyglossus aculeatus^ 23E skin J7770*** Healesville Sanctuary SA 1989 Skin tissue with numerous nymphs * For corresponding tick specimen refer to Appendix A, Table A.1

** Skin biopsy removed with an attached tick (see Appendix A, Table A.1)

*** This echidna will be referred to as host number 8 in the study

^ Sick echidna (recorded by the South Australia Museum for host registration code J7770)

156 Appendix B

Table B.1. Comparative character matrix for the genera Amblyomma, Bothriocroton, and Ixodes. Original work by author. Images sourced from Barker and Walker (2014). Character State Ambl. Both. Ixo. Reference

Dorsal View:

Body profile Narrow ☐ ☐  from above oval (unfed tick) For copyright purposes please refer to Figures 1a-m Broad  ☐ ☐ oval in Barker and Walker (2014)

Circular ☐  ☐

Eyes Present  ☐ ☐

Absent ☐  

Enamel Present   ☐ patterns on scutum or conscutum

Absent ☐ ☐ 

Festoons at Present   ☐ posterior body margin (unfed tick) Absent ☐ ☐ 

Pale rings Present   ☐ on legs

Absent ☐ ☐ 

Width of Wider ☐ ☐  mouthparts relative to Equal  ☐ ☐ basis capituli Narrower ☐  ☐

157

Ventral view:

Sclerotized Present ☐ ☐  plate aligns with anus Absent   ☐ (males only)

Anal groove Anterior ☐ ☐  position relative to anus Posterior   ☐

158 Appendix C

Table C.1. NGS mapping file

#Sample ID Index.1 Index.2 Forward. Primer Reverse. Primer Genus Species Specimen. type Life.stage. Instar Location State Year. collected Museum Archived. echidna. host. registration .# Other. description GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC South.of.Mool Skin.tissue.at.3Ta. tis-3 N716 S502 TAA TAAT Tachyglossus aculeatus skin Tissue oolah QLD 1992 QLD 1 tick.bite.site GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC South.of.Mool tis-6 N716 S503 TAA TAAT Tachyglossus aculeatus spleen Tissue oolah QLD 1992 QLD 1 Spleen.biopsy GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. Skin.tissue.at.13T tis-13 N716 S505 TAA TAAT Tachyglossus aculeatus skin Tissue Forestdale QLD date QLD 2 a.tick.bite.site GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. tis-16 N703 S502 TAA TAAT Tachyglossus aculeatus spleen Tissue Forestdale QLD date QLD 2 Spleen.biopsy GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Skin.with.tick.bio tis-23 N716 S507 TAA TAAT Tachyglossus aculeatus skin Tissue Karana.Downs QLD 2007 QLD 3 psy GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC tis-26 N716 S508 TAA TAAT Tachyglossus aculeatus spleen Tissue Karana.Downs QLD 2007 QLD 3 Spleen.biopsy GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Queen.Mary.F tis-36 N716 S510 TAA TAAT Tachyglossus aculeatus spleen Tissue alls QLD 2004 QLD 4 Spleen.biopsy GTGCCAGC GGACTACH Skin.and.tick.biop MGCCGCGG VGGGTWTC sy.male.and.femal tis-43 N716 S511 TAA TAAT Tachyglossus aculeatus skin Tissue Ipswich QLD 2011 QLD 5 e. GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC tis-46 N703 S503 TAA TAAT Tachyglossus aculeatus spleen Tissue Ipswich QLD 2011 QLD 5 Spleen.biopsy GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Skin.with.tick.bio tis-52 N718 S503 TAA TAAT Tachyglossus aculeatus skin Tissue Highvale QLD 2006 QLD 6 psy GTGCCAGC GGACTACH Healesville.Sa Skin.tissue.with.n tis-523 N703 S505 MGCCGCGG VGGGTWTC Tachyglossus aculeatus skin Tissue nctuary SA 1989 SA J7770 umerous.nymphs

159 TAA TAAT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC South.of.Mool e1 N718 S507 TAA TAAT Bothriocroton concolor tick Male oolah QLD 1992 QLD 1 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC South.of.Mool e2 N718 S508 TAA TAAT Bothriocroton concolor tick Nymph oolah QLD 1992 QLD 1 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC South.of.Mool e5 N718 S510 TAA TAAT Bothriocroton concolor tick Female oolah QLD 1992 QLD 1 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. e16 N718 S511 TAA TAAT Bothriocroton concolor tick Nymph Forestdale. QLD date QLD 2 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. e18 N719 S502 TAA TAAT Bothriocroton concolor tick Female Forestdale. QLD date QLD 2 Grossly.engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. e19 N703 S507 TAA TAAT Bothriocroton concolor tick Female Forestdale. QLD date QLD 2 Grossly.engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC no. e20 N719 S505 TAA TAAT Bothriocroton concolor tick Male Forestdale. QLD date QLD 2 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e23 N719 S506 TAA TAAT Bothriocroton concolor tick Female Karana.Downs QLD 2007 QLD 3 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e24 N719 S507 TAA TAAT Bothriocroton concolor tick Female Karana.Downs QLD 2007 QLD 3 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e25 N719 S508 TAA TAAT Bothriocroton concolor tick Female Karana.Downs QLD 2007 QLD 3 Slightly.engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e26 N719 S510 TAA TAAT Bothriocroton concolor tick Male Karana.Downs QLD 2007 QLD 3 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Queen.Mary.F e27 N719 S511 TAA TAAT Bothriocroton concolor tick Male alls QLD 2004 QLD 4 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Queen.Mary.F e29 N720 S502 TAA TAAT Bothriocroton concolor tick Nymph alls QLD 2004 QLD 4 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e32 N720 S503 TAA TAAT Bothriocroton concolor tick Female Ipswich QLD 2011 QLD 5 Engorged

160 GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e33 N720 S505 TAA TAAT Bothriocroton concolor tick Male Ipswich QLD 2011 QLD 5 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e35 N720 S506 TAA TAAT Bothriocroton concolor tick Nymph Ipswich QLD 2011 QLD 5 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e39 N720 S507 TAA TAAT Bothriocroton concolor tick Female Highvale QLD 2006 QLD 6 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e41 N720 S508 TAA TAAT Bothriocroton concolor tick Male Highvale QLD 2006 QLD 6 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e43 N720 S510 TAA TAAT Bothriocroton concolor tick Female Malanda QLD 2003 QLD 7 Slightly.engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC e45 N703 S508 TAA TAAT Bothriocroton concolor tick Nymph Malanda QLD 2003 QLD 7 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 1 N703 S510 TAA TAAT Bothriocroton concolor tick Female Tooloom NSW 1961 ANIC 48001053 Grossly.Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 2 N721 S503 TAA TAAT Bothriocroton concolor tick Nymph Tooloom NSW 1961 ANIC 48001053 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Flattened.slight.te 3 N721 S505 TAA TAAT Bothriocroton concolor tick Female Biddon NSW 1993 ANIC 48001072 ar GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 4 N721 S506 TAA TAAT Bothriocroton concolor tick Male Biddon NSW 1993 ANIC 48001072 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 5 N703 S511 TAA TAAT Bothriocroton concolor tick Male Batemans.Bay NSW 1957 ANIC 48001049 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 6 N721 S508 TAA TAAT Bothriocroton concolor tick Nymphs Batemans.Bay NSW 1957 ANIC 48001049 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 7 N721 S510 TAA TAAT Bothriocroton concolor tick Female Nanungo QLD 1968 ANIC 48000993 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 8 N721 S511 TAA TAAT Bothriocroton concolor tick Male Nanungo QLD 1968 ANIC 48000993 Normal GTGCCAGC GGACTACH 9 N704 S502 MGCCGCGG VGGGTWTC Amblyomma triguttatum tick Male Bowen QLD 1997 ANIC 48000639 Normal

161 TAA TAAT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Normal.slightly.fl 11 N704 S503 TAA TAAT Bothriocroton concolor tick Male Clermont QLD 1961 ANIC 48001055 attened GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 13 N722 S507 TAA TAAT Bothriocroton concolor tick Female Uralla NSW 1958 ANIC 48001045 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 14 N722 S508 TAA TAAT Bothriocroton concolor tick Male Uralla NSW 1958 ANIC 48001045 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 18 N704 S505 TAA TAAT Bothriocroton tachyglossi tick Female Yepoon QLD 1967 QLD 588998 Grossly.Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Cinnabar.Goo 19 N704 S506 TAA TAAT Bothriocroton tachyglossi tick Female meri QLD 1988 QLD 590000 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Monbulk.Mace 26 N723 S502 TAA TAAT Bothriocroton concolor tick Female lesfield VIC 1969 VIC 2016.17L Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 30 N723 S503 TAA TAAT Bothriocroton concolor tick Male Mt.Abbott QLD 2007 QLD S37861 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 31 N723 S505 TAA TAAT Bothriocroton concolor tick Female Mt.Abbott QLD 2007 QLD S37861 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 34 N723 S506 TAA TAAT Amblyomma australiense tick Female Mt.Isa QLD 2002 QLD S33975 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 36 N723 S507 TAA TAAT Amblyomma australiense tick Female Mt.Isa QLD 2002 QLD S33702 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 37 N723 S508 TAA TAAT Amblyomma australiense tick Nymph Mt.Isa QLD 2002 QLD S33702 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 38 N723 S510 TAA TAAT Amblyomma australiense tick Male Mt.Isa QLD 2002 QLD S33702 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 45 N723 S511 TAA TAAT Bothriocroton concolor tick Female Buccan QLD 2011 QLD S32257 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 46 N724 S502 TAA TAAT Bothriocroton concolor tick Male Buccan QLD 2011 QLD S32257 Normal

162 GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 47 N724 S503 TAA TAAT Bothriocroton concolor tick Nymphs Buccan QLD 2011 QLD S32257 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Chinchilla.Gra 52 N724 S505 TAA TAAT Bothriocroton concolor tick Female cehithgo QLD 1985 QLD S69926 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Engorged.From.si 54 N724 S506 TAA TAAT Bothriocroton concolor tick Female Maleny QLD 2000 QLD S45426 ck.echidna GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Normal.From.sick. 55 N724 S507 TAA TAAT Bothriocroton concolor tick Male Maleny QLD 2000 QLD S45426 echidna GTGCCAGC GGACTACH Lake.Hart.Gair MGCCGCGG VGGGTWTC dner.Torrens. 61 N724 S508 TAA TAAT Amblyomma triguttatum tick Female Basin SA 1978 SA J7608.26 Grossly.Engorged GTGCCAGC GGACTACH Lake.Hart.Gair MGCCGCGG VGGGTWTC dner.Torrens. 62 N724 S510 TAA TAAT Amblyomma triguttatum tick Male Basin SA 1978 SA J7608.26 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 67 N726 S503 TAA TAAT Bothriocroton concolor tick Female Victoria VIC 1974 SA J1798 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 68 N726 S505 TAA TAAT Bothriocroton concolor tick Male Victoria VIC 1974 SA J1798 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Millicent.Lowe Normal.from.sick. 71 N726 S506 TAA TAAT Bothriocroton concolor tick Male r.South.East SA 1981 SA J7745.57 echidna GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Millicent.Lowe Normal.from.sick. 72 N726 S507 TAA TAAT Bothriocroton concolor tick Female r.South.East SA 1981 SA J7745.57 echidna GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Millicent.Lowe Normal.from.sick. 73 N726 S508 TAA TAAT Bothriocroton concolor tick Nymphs r.South.East SA 1981 SA J7745.57 echidna GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 74 N726 S510 TAA TAAT Ixodes tasmani tick Female Fingal TAS 1997 ANIC 48003382 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 75 N726 S511 TAA TAAT Ixodes tasmani tick Female Antill.Ponds TAS 1960 ANIC 48003473 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 78 N727 S502 TAA TAAT Ixodes tasmani tick Female Maydena TAS 1959 ANIC 48003458 Normal GTGCCAGC GGACTACH 80 N727 S503 MGCCGCGG VGGGTWTC Bothriocroton hydrosauri tick Male Rockhampton QLD 1963 ANIC 48001159 Normal

163 TAA TAAT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 82 N727 S505 TAA TAAT Amblyomma echidnae tick Male Townsville QLD 1979 ANIC 48000367 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 83 N727 S506 TAA TAAT Ixodes tasmani tick Nymphs Scamander TAS 1996 ANIC 48003343 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 86 N727 S507 TAA TAAT Amblyomma australiense tick Male Darwin NT 1974 ANIC 48000312 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 87 N727 S508 TAA TAAT Amblyomma australiense tick Female Darwin NT 1974 ANIC 48000312 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 92 N727 S510 TAA TAAT Bothriocroton hydrosauri tick Male Rockhampton QLD 1966 ANIC 48001160 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 93 N727 S511 TAA TAAT Bothriocroton hydrosauri tick Nymphs Rockhampton QLD 1966 ANIC 48001160 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 94 N728 S502 TAA TAAT Bothriocroton hydrosauri tick Female Rockhampton QLD 1966 ANIC 48001160 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Justa.Creek.Ch Engorged.missing. 95 N728 S503 TAA TAAT Amblyomma fimbriatum tick Female ina.Wall NT 1967 ANIC 48000376 head GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Justa.Creek.Ch Engorged.missing. 96 N728 S505 TAA TAAT Amblyomma fimbriatum tick Female ina.Wall NT 1967 ANIC 48000376 head GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Justa.Creek.Ch 98 N728 S506 TAA TAAT Amblyomma fimbriatum tick Male ina.Wall NT 1967 ANIC 48000376 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Justa.Creek.Ch 99 N728 S507 TAA TAAT Amblyomma fimbriatum tick Male ina.Wall NT 1967 ANIC 48000376 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 101 N728 S508 TAA TAAT Bothriocroton concolor tick Female Heathcote NSW 1993 WA T141308 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 104 N728 S510 TAA TAAT Bothriocroton concolor tick Female Tooloom NSW 1965 ANIC 4800106 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 106 N728 S511 TAA TAAT Bothriocroton concolor tick Female Anakie QLD 1998 ANIC 48001065 Engorged

164 GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 108 N729 S502 TAA TAAT Bothriocroton concolor tick Female Dayboro QLD 2006 QLD S73748 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Straun.Lower. 110 N729 S503 TAA TAAT Bothriocroton concolor tick Female South.East SA 1981 SA J7743.4 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Snug.Cove.Kan 112 N729 S505 TAA TAAT Bothriocroton concolor tick Female garoo.Island SA 1988 SA J7758 Engorged GTGCCAGC GGACTACH Bridgewater. MGCCGCGG VGGGTWTC Mt.Lofty.Rang 113 N729 S506 TAA TAAT Bothriocroton concolor tick Female es SA 1989 SA J7769 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Healesville.Sa Various.degrees.o 114 N729 S507 TAA TAAT Bothriocroton concolor tick Nymphs nctuary SA 1989 SA J7770 f.engorgement GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC NT-1 N729 S508 TAA TAAT NTC NTC NTC NTC NTC NTC NTC NTC NTC NTC GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex2 N701 S503 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex3 N701 S505 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex4 N701 S506 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex5 N701 S507 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC EX-tis N701 S508 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex47 N701 S510 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC Ex48 N701 S511 TAA TAAT EXT EXT EXT EXT EXT EXT EXT EXT EXT EXT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 15 N702 S502 TAA TAAT Bothriocroton concolor tick Female Brisbane QLD 1941 ANIC 48001042 Normal GTGCCAGC GGACTACH 16 N702 S503 MGCCGCGG VGGGTWTC Bothriocroton concolor tick Male Brisbane QLD 1941 ANIC 48001042 Normal

165 TAA TAAT GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 20 N702 S505 TAA TAAT Bothriocroton concolor tick Male Gatton QLD 1954 ANIC 48001041 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 21 N702 S506 TAA TAAT Bothriocroton concolor tick Female Gatton QLD 1954 ANIC 48001041 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 32 N702 S507 TAA TAAT Bothriocroton concolor tick Female Mt.Ommaney QLD 2001 QLD S19459 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 33 N702 S508 TAA TAAT Bothriocroton concolor tick Male Mt.Ommaney QLD 2001 QLD S19459 Normal GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC 102 N702 S510 TAA TAAT Bothriocroton concolor tick Female Warwick QLD 1963 ANIC 48001057 Engorged GTGCCAGC GGACTACH MGCCGCGG VGGGTWTC NT2 N702 S511 TAA TAAT NTC NTC NTC NTC NTC NTC NTC NTC NTC NTC

166 Table C.2. Legend for taxonomic assignment of bacterial phyla in QIIME.

167 Table C.3. Legend for taxonomic assignment of bacterial genera in QIIME.

168

Figure C.1. Bacterial diversity in echidna tissue and tick species according to instar, generated in QIIME. 169 Appendix D

Figure D.1. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples of 1:10 dilution ratio. Amplification curves for the extraction blank and no-template control are labelled.

170

Figure D.2. The amplification plots (A and B) obtained for the 16S Mam qPCR assay for echidna tissue DNA samples at neat concentration. Amplification curves for the extraction blank and no-template control are labelled.

171

Figure D.3. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples at neat concentration. Amplification curves for the no-template control and positive control are labelled.

172

Figure D.4. The amplification plots (A) without positive spike DNA, and (B) with positive spike DNA, acquired for the flaB qPCR assay for echidna tissue DNA samples at 1:10 dilution ratio.

173