Blood Parasites in Endangered Wildlife-Trypanosomes Discovered during a Survey of Haemoprotozoa from the Tasmanian Devil

The impact of emerging infectious diseases is increasingly recognised as a major threat to wildlife. Wild populations of the endangered Tasmanian devil, Sarcophilus harrisii, are experiencing devastating losses from a novel transmissible cancer, devil facial tumour disease (DFTD); however, despite the rapid decline of this species, there is currently no information on the presence of haemoprotozoan parasites. In the present study, 95 Tasmanian devil blood samples were collected from four populations in Tasmania, Australia, which underwent molecular screening to detect four major groups of haemoprotozoa: (i) trypanosomes, (ii) piroplasms, (iii) Hepatozoon, and (iv) haemosporidia. Sequence results revealed Trypanosoma infections in 32/95 individuals. Trypanosoma copemani was identified in 10 Tasmanian devils from three sites and a second Trypanosoma sp. was identified in 22 individuals that were grouped within the poorly described T. cyclops clade. A single blood sample was positive for Babesia sp., which most closely matched Babesia lohae. No other blood protozoan parasite DNA was detected. This study provides the first insight into haemoprotozoa from the Tasmanian devil and the first identification of Trypanosoma and Babesia in this carnivorous marsupial.


Introduction
Haemoprotozoan parasites are unicellular eukaryotic organisms with complex lifecycles that involve an invertebrate vector and often alternate in their tropism between the tissues and blood of their vertebrate hosts. Four major haemoprotozoan assemblages infect mammals [1]; (i) Trypanosomatids (Kinetoplastea), which are flagellated protists that are characterised by the presence of a unique The present study aimed to screen blood samples from Tasmanian devils for the presence of haemoprotozoan parasites. To the authors' knowledge, the present study provides the first survey of haemoprotozoa from Tasmanian devils.

Study Sites and Sampling
Animal use was approved by the University of Tasmania Animal Ethics Committee permit numbers A0015835 and A0016789) and the Department of Primary Industries, Parks, Water and Environment Animal Ethics Committee (permit numbers TFA19144 and TFA18028).
A total of 95 blood samples were collected from wild Tasmanian devils that were captured during the austral autumn (May 2018) from four sites across Tasmania: Black River, Takone, West Pencil Pine and Freycinet (Figure 1). The sampling effort at each site involved deployment of 40 PVC pipe traps during seven to 10 consecutive nights. Individual devils were identified via subcutaneously implanted microchips (AllFlex© ISO FDX-B).
Blood was collected (between 0.3-1 mL) from either the jugular vein (Takone) or marginal ear vein (Black River, Freycinet and West Pencil Pine). The puncture site was disinfected with sterile alcohol swabs for at least 15 s prior to collection and blood was stored in ethylenediaminetetraacetic acid (EDTA) vacutainers. The samples were kept refrigerated at 4 • C and then shipped to Murdoch University, Western Australia where they were stored at −20 • C until analysis.

DNA Extraction
The total genomic DNA was extracted from 200 µL of blood using a MasterPure DNA purification kit (Epicentre ® Biotechnologies, Madison, WI, USA) following the manufacturer's recommendations. Where 200 µL of blood was not available, sterile DNA free phosphate-buffered saline (PBS) was used to make samples up to 200 µL. Genomic DNA (gDNA) was eluted in 30 µL of TE buffer and stored at −20 • C . Extraction controls (EXBs), consisting of 200 µL sterile DNA free PBS, were randomly included in each extraction batch (N = 7).

PCR Assays
Molecular screening was carried out for haemoprotozoa on gDNA from blood samples (N = 95) and EXBs (N = 7). Reactions were carried out in 25 µL volumes containing 2 µL of gDNA or 1 µL of the primary product for nested and semi-nested PCR assays. Table 1 provides an overview of assays, including primer sequences. GCATTATCTGGATGTGATAATGGT [38] For the identification of trypanosomes (Trypanosoma and Leishmania spp.) samples were screened using a nested PCR assay targeting an~959 bp product of the second half of the 18S ribosomal RNA (18S rRNA) locus. This assay amplifies most species of Trypanosoma [32] and it was also validated in the present study to amplify Leishmania spp. using control gDNA of Leishmania infantum from a canine bone marrow sample and Leishmania macropodum culture isolate. External primers SLF/S762R and internal primers S825F/SLIR were used as an initial screen and then sequenced as per details below. Samples that were representative of the different genotypes identified then underwent an additional secondary assay with a second set of internal primers S823/S662R to yield a near full length 18S rDNA sequence. Reactions were carried out in 25 µL volumes containing 0.4 µM of each primer and 12.5 µL GoTaq®Green master mix (Promega, Madison, WI, USA). Thermal cycling conditions were as follows; initial cycle of 95 • C for 5 min, 50 • C for 2 min, 72 • C for 4 min, followed by 35 cycles of 95 • C for 30 s, 52 • C (primary) or 55 • C (secondary) for 30 s, 72 • C for 2 min. 20 s (primary) or 1 min. (secondary), and a final extension of 72 • C for 5 min. To obtain further phylogenetic information from the genotypes that were identified by 18S rRNA locus screening, samples underwent amplification of the glycosomal Glyceraldehyde Phosphate Dehydrogenase (gGAPDH) gene using a semi-nested PCR assay with external primers GAPDHF/GAPDHR and internal primers GAPDHF/Ga4 [32]. Reactions contained 1X buffer (KAPA Biosystems, Cape Town, South Africa), 2.0 mM (MgCl 2 ), 0.4 µM of each primer, 0.25 mM of each dNTP, and 0.5 U of Taq (KAPA Biosystems, Cape Town, South Africa). Thermal cycling conditions were as follows; initial cycle of 95 • C for 5 min, 50 • C for 2 min, 72 • C for 4 min, followed by 35 cycles of 95 • C for 30 s, 52 • C (primary) or 55 • C (secondary) for 30 s, 72 • C for 2 min. 20 s, and a final extension of 72 • C for 5 min.
Screening for piroplasms (Babesia and Theileria spp.) utilised a previously described nested PCR assay. Primers were used targeting an~800 bp product of the 18S rRNA locus with external primers BT1F/BT1R and internal primers BT2F/BT2R [35]. The reactions contained 1× buffer (KAPA Biosystems, Cape Town, South Africa), 2.0 mM (MgCl 2 ), 0.4 µM of each primer, 0.25 mM of each dNTP, and 0.5 U of Taq (KAPA Biosystems, Cape Town, South Africa). Thermal cycling conditions were as follows; initial cycle of 95 • C for 2 min, 58 • C for 1 min, 72 • C for 2 min, followed by 35 cycles of 95 • C for 30 s, 62 • C for 20 s, 72 • C for 45 s, and a final extension of 72 • C for 7 min.
A genus specific assay for Hepatozoon targeting the 18S rRNA locus was used with primers HepF300/Hep900 in order to amplify an~600 bp product [36]. Reactions contained 1× buffer (KAPA Biosystems, Cape Town, South Africa), 1.5 mM (MgCl 2 ), 0.4 µM of each primer, 0.25 mM of each dNTP, and 0.5 U of Taq (KAPA Biosystems, Cape Town, South Africa). Thermal cycling conditions were as follows; initial denaturation of 95 • C for 3 min, followed by 40 cycles of 95 • C for 30 s, 60 • C for 30 s, 72 • C for 1 min, and a final extension of 72 • C for 10 min.
Screening for haemosporidia (Plasmodium spp.) utilised a previously described nested PCR assay [37]. Primers targeting an~420 bp product of the cytochrome b (cytb) gene with external primers HAEMNF/HAEMNR2 and internal primers HAEMF/HAEMR2. Reactions contained 1× buffer (KAPA Biosystems, Cape Town, South Africa), 1. Suitable positive controls, extraction reagent blanks, and no-template PCR controls were included throughout the laboratory processes. Extractions, pre-PCR and post-PCR procedures were performed in laboratories physically separated from each other.

Gel Electrophoresis and Sanger Sequencing
Amplicons were electrophoresed on a 1% agarose gel stained with SYBR safe (Invitrogen, Waltman, MA, USA). Products of the correct size (see Table 1) were excised from the gel and purified while using the previously described methods [39]. Sanger sequencing was carried out in forward and reverse directions on all positive amplicons. Sequencing was performed at the Australian Genome Research Facility (Perth, Western Australia) on an Applied Biosystems 3730 using Big Dye Terminator chemistry version 3.1.

Phylogenetic Analysis
The sequences were imported and trimmed in Geneious 10.2.6 (https://www.geneious.com) and then subjected to BLAST analysis using BLASTN 2.10.0+ [40] against nucleotide collection (nt) database [41] to identify the most similar species and genotypes. Reference sequences were retrieved from GenBank [42] (details available in Supplementary Table S1) and aligned with sequences obtained in the present study using MUSCLE [43]. Alignments of the 18S rDNA and the gGAPDH sequences were then used for phylogenetic purposes. Phylogenies were inferred while using the maximum likelihood (ML) method. The optimal evolutionary model was selected while using ModelFinder [44] based on bayesian information criterion. Phylogenetic analysis was performed in IQ-TREE v1.6.11 [45] and bootstrap support was calculated using the ultrafast (UFBoot2) method with 10,000 replicates [46].
Maximum likelihood phylogeny based on the 18S rRNA locus was performed using a 1505 bp alignment based on the transition (AC = CG, AT = GT with equal base frequencies) (TIM3e) substitution model [47], with invariable sites (I = 0.408) and a discrete gamma distribution (four categories) (G4 = 0.438) [48]. To include a wider range of neighbouring reference sequences, a second phylogenetic analysis was performed on the V7-8 hypervariable region of the 18S rRNA locus; this region is useful to differentiate between closely related trypanosome sequences [49]. A phylogeny was produced using a 559 bp alignment that was based on the Kimura Two-Parameter (K2P) substitution model [50] with discrete gamma distribution (four categories) (G4 = 0.227) [48]. A 767 bp alignment of the gGAPDH gene was used for phylogenetic reconstruction based on the Tamura-Nei (TN) substitution model [51], with empirical base frequencies (F), invariable sites (I = 0.387) and discrete gamma distribution (four categories) (G4 = 0.893) [48]. Genetic sequence similarity was calculated while using the Kimura Two-Parameter method [51].

Statisical Analysis
The overall prevalence of Trypanosoma species was compared between males and females while using two-tailed Fisher Exact test.

Microscopy of Blood Smears
Thin blood smears were prepared from animals that were sampled at Takone site only (three per individual), within 4 h of collection, and then air dried and fixed in methanol. Blood smears were then stained with modified Wright-Giemsa (Hematek®Stain Pak) using a Hema-Tek Slide Stainer (Ames Company Division, Miles Laboratories Pty Ltd., Victoria, Australia) and a coverslip was mounted using DPX neutral mounting medium (LabChem, Victoria, Australia). The smears were inspected by light microscopy (Olympus BX51) for the presence of haemoprotozoa at ×400 magnification and under oil immersion (×1000).

Molecular Screening
Molecular screening identified 33.7% (32/95) of the blood samples were positive for Trypanosoma DNA. Sequence identity from BLAST results identified; T. copemani and T. cyclops-like in 10.5% (n = 10) and 23.2% (n = 22) of the individuals respectively (Table 2). Trypanosoma copemani was almost exclusively identified in males (90.0%, p = 0.0159), while T. cyclops-like genotypes were more commonly found in females (68.2%, p = 0.0432). All of the samples tested negative for presence of Leishmania, Hepatozoon, Plasmodium, and Theileria species. A comparison at the 18S V7-8 hypervariable region of the ten T. copemani sequences showed they were all >99% similar to each other and, as such, a representative sample (BRI115) was used for subsequent phylogeny. Analysis of long 18S alignment concluded that the BRI115 sample shared 99.1% similarity to T. copemani Charlton (GU966588) and 98.5% with T. copemani H26 (AJ009169) (Figure 2). At the gGAPDH locus BRI115 sample was identical to T. copemani Mika (GU966585), and 99.7% similar to T. copemani AAP (AJ620277) (Figure 3). There is no 18S rDNA sequence data for T. copemani Mika available. A comparison of T. copemani Charlton (GU966584) at the gGAPDH locus showed it was 99.9% similar to the BRI115 sample with one single nucleotide polymorphism (SNP) (data not shown). At the 18S V7-8 hypervariable region two genotypes of the T. cyclops clade were identified, referred to as Tasmanian devil genotype A (n = 13) and genotype B (n = 9). Representatives of each genotype were used for phylogenetic reconstruction that was based on a longer alignment of 18S rDNA sequences ( Figure 2). Analysis of near full length 18S rDNA alignment showed sequences were identical within each genotype (genotype A samples WPP585 and WPP601, and genotype B samples TKN211 and BRI111  (Figure 3), with just one SNP. The most similar sequence was Trypanosoma sp. ABF (AJ620278), which was 97.2% and 97.4% similar to WPP601 and WPP602, respectively. The next most similar sequence was T. cyclops (FJ649493), which was 95.7% and 95.8% similar to WPP601 and WPP602, respectively. Unfortunately, GAPDH sequences from genotype B were not successfully amplified. Phylogenetic reconstruction utilising the V7-8 hypervariable region of the 18S rRNA locus showed support for a monophyletic T. cyclops clade ( Figure 4). Sequences from the present study grouped with genotypes detected from Malaysia, Sri Lanka and Australia (Queensland and Victoria) with the clear distinction of two different genotypes of T. cyclops from Tasmanian devils (Supplementary Table S2).   Supplementary Table S1. A single sample (BRI115) obtained from Black River site was positive for a Babesia sp. and this individual was also positive for T. copemani infection. Analysis of an 800 bp fragment of the 18S rRNA locus revealed it was similar to Babesia sp. identified from Ixodes tasmani in Queensland (MG251436; 96.0% similarity) and Babesia lohae from Ixodes holocyclus in Queensland (MG593272; 95.5% similarity).

Microscopy
Microscopy of blood smears from Takone site did not yield any positive detection of haemoparasites.  Supplementary Table S1.

Discussion
The present study represents the first survey of haemoprotozoa from wild populations of the endangered Tasmanian devil. Our findings of Trypanosoma infections across all four sites suggests that this infection is widespread and potentially endemic within Tasmanian devil populations. This wide distribution of Trypanosoma across populations contrasts with the absence of Leishmania, Theileria, Hepatozoon, or Plasmodium species, and the low detection of Babesia, which was only detected in a single individual.
The extension of the host range of T. copemani is notable and it supports recent research identifying this parasite in a wide range of marsupial hosts from across Australia. To date, T. copemani has been identified in all Australian states and territories, except the Northern Territory and South Australia. The identification of this parasite in Tasmanian devils is potentially significant from a health perspective as previous reports have associated it with pathological changes in the woylie (Bettongia penicillata) [22,23] and in koalas (Phascolarctos cinereus) with co-morbidities [24]. With this report, there are now at least ten Australian vertebrate host records for T. copemani; therefore, this Trypanosoma species appears to be a marsupial generalist and capable of infecting a diverse range of mammals. Further studies for mapping the complete distribution and host range of T. copemani will help to provide insights into the co-evolution of this native trypanosome in its marsupial hosts.
The identification of genotypes from the Trypanosoma cyclops clade was unexpected. Trypanosoma cyclops was described from wild caught southern pig-tail macaques (Macaca nemestrina) in jungle areas of West Malaysia [52] and Hamilton et al. [53] identified novel genotypes of this clade from Australia, Papua New Guinea, and Sri Lanka. However, since its identification, there have been very few published reports of T. cyclops. Phylogeny within the T. cyclops clade was not well resolved, as demonstrated by the polytomy in Figure 4. The identification of two distinct genotypes in devils that do not form a monophyletic group and show different levels of relatedness to sequences overseas highlights the uncovered diversity within this clade. The closest named species to T. cyclops is Trypanosoma (Megatrypanum) theileri, which has a cosmopolitan distribution and predominantly infects cattle. The most important vector of T. theileri is thought to be tabanid flies (Tabanidae) [54,55]. While this trypanosome is generally considered non-pathogenic, chronic infection has been associated with the development of secondary diseases in cattle [56].
The gGAPDH gene has been shown to be a more suitable marker than the 18S rRNA locus for determining the phylogeny within the Trypanosoma genus [34]. A comparative analysis of T. copemani shows the intra-specific sequence similarity within the clade is 97.0-100% and inter-specific similarity to the nearest named species, T. gilletti, is 91.3-92.3%. Intra-specific sequence similarity within the T. cyclops clade was 94.8-99.9%, while the inter-specific sequence similarity to nearest named species, T. theileri, is 90.4%. Therefore, there is sufficient support that the sequences generated for devils can be attributed to T. cyclops. The differentiation of genotypes within this clade are best identified by analysis of the V7-8 hypervariable region of the 18S rRNA locus (Figure 4, Supplementary Table S2). This study further supports that T. cyclops is more closely related to sequences obtained from Australia, Papua New Guinea, and Sri Lanka than it is to T. theileri. [53]. There is no geographical or host distinction between the different genotypes, further supporting the taxonomy as a single species, as shown in the phylogeny within the T. cyclops clade (Figure 4). Research on functional traits, including growth dynamics in culture coupled with morphological analysis, is needed to better understand the taxonomy of this clade. The geographical and host extensions reported here will be useful for the future work that aimed at understanding the evolutionary history of the T. cyclops and T. theileri clades and the Trypanosoma genus more broadly [57].
The description of a T. cyclops-like species in this work expands the number of Trypanosoma species in Tasmanian mammals to four. The platypus-specific T. binneyi has been observed from blood samples and subsequently described phylogenetically [29,58,59] with suggestions that leeches are the likely vector [59]. Trypanosomes have also been identified in populations of the eastern barred bandicoot (Perameles gunnii) and southern brown bandicoot (Isoodon obesulus) [60]; however, due to well-described limitations of morphological identification [61], species-level identification has not been achieved. Additionally, T. copemani was identified for the first time in Tasmania in 2020 from wild populations of the eastern quoll (Dasyurus viverrinus) [62]. The current study now brings the number of known trypanosome hosts in Tasmania to five, with the addition of Tasmanian devils, and the confirmation of a new species from the T. cyclops clade. It is interesting to note that, while previous studies have only described one species of Trypanosoma in the sampled hosts from Tasmania, in the current study we have described at least two species circulating in devils. The higher diversity of Trypanosoma species found in devils might be a consequence of our larger sample size or spatial coverage in comparison to previous studies.
Despite the molecular identification of Trypanosoma infection in devils from Takone, no trypomastigotes were observed in corresponding blood smears. This finding is consistent with other studies that have found molecular tools are more sensitive than microscopy for trypanosome detection [63,64]. The absence of patent parasitaemia (i.e. no flagellates observed in fresh blood smears) could be the result of low-level infections; however, it may also indicate that devils are not a competent host of the Trypanosoma species [65].
The phylogenetic position of the T. cyclops clade within the stercorarian group means that its potential vector(s) could be a range of arthropod(s) with transmission occurring via contact with vector faeces. While it has been hypothesised that carnivory is important in the maintenance of trypanosomes, experimental studies do not support this [66] and, instead, suggested that increased infection is attributed to insectivores who consume infected arthropod vectors [67]. However, it is interesting that, although members of this clade have been identified from leeches and frogs [53], their phylogenetic position is distinct from other aquatic and leech-associated Trypanosoma species [34,59,68,69]. The range extensions of both T. copemani and T. cyclops clades that were recorded in the present study provide additional data to aid vector identification. The large geographical range of both these Trypanosoma species suggests that any vector(s) should be equally ubiquitous and capable of biting or coming into close contact with a wide range of vertebrate hosts.
In the present study, we validated that a previously published Trypanosoma PCR assay [32] is also capable of detecting/amplifying Leishmania DNA. Two controls of genetically diverse species of Leishmania, L. infantum and L. macropodum, [70] were used. In the context of this study, we determined that generic trypanosome primers are able to amplify L. macropodum, which was identified as a cause of cutaneous leishmaniasis in kangaroos [71]. The preferred diagnostic sample for the detection of Leishmania is generally bone marrow aspirate or tissue samples; however, previous studies have recorded detection via blood samples [72,73]. Obtaining bone marrow and internal tissue samples from free ranging wildlife is not practical or ethical, especially where populations are experiencing significant declines. While the present study reported the absence of Leishmania via PCR of blood samples, it is not possible to discount occult infection and future studies utilising additional sample types would be recommended. Although co-infections of Leishmania and Trypanosoma cannot be completely ruled out in the individuals tested here, the absence of Leishmania DNA is consistent with previous studies that have utilised the same assay on other Australian mammals [17,18,74]. Therefore, reports of Australia's only endemic Leishmania species remain confined to kangaroos and biting midges (Ceratopogonidae) in the Northern Territory [71,75,76].
An unexpected outcome was the low detection of Babesia species and lack of detection of Theileria and Hepatozoon species. Recent molecular studies in Australian wildlife have demonstrated a range of unique endemic piroplasms, with reports of up to 80-90% prevalence in some marsupial host populations [2,19,77]. Piroplasms and Hepatozoon spp. are transmitted by ticks and, therefore, dependent on the density and interactions between host and vector(s) to continue their lifecycle. Recent investigations have revealed the presence of numerous novel Babesia, Theileria, and Hepatozoon species in native Australian ticks [78][79][80]. With respect to the single identification of Babesia at the Black River site, the genetic relationship shows that it is most closely related to Babesia lohae sequences identified in Ixodes holocyclus and Ixodes tasmani ticks from the east coast of Queensland [78,79]. The low prevalence of Babesia might be attributed to the seasonality of tick infestations, as has been described in the relationship between I. tasmani ticks and brushtail possums hosts [81]. Future longitudinal sampling, including multiple seasons, are valuable to better understanding the phylogeny and epidemiology of this Babesia species circulating in devil populations.
In Tasmania, piroplasms and Hepatozoon species have been identified from three marsupial species and, more recently, from ticks (Ixodes spp.). A Hepatozoon has been identified in eastern barred bandicoots (Perameles gunnii) [82] by morphological examination of blood smears and the eastern quoll (Dasyurus viverrinus) through molecular tools [62]. Theileria species have been identified from eastern bettongs (Bettongia gaimardi) [83] and eastern quolls [62]. A molecular survey of ticks (I. tasmani) from Tasmanian devils demonstrated that 34.1% (15/44) of sample pools were positive for Hepatozoon sp. [84]. More recently, novel species of Theileria and Hepatozoon were identified from I. tasmani collected in Tasmania [78,79]. Given the high prevalence of I. tasmani parasitising sampled Tasmanian devils (Ruiz-Aravena pers. comms.), the low prevalence of piroplasms and Hepatozoon species is, therefore, unexpected. This raises important questions about the sylvatic lifecycle and reservoir hosts of these piroplasm and Hepatozoon species, suggesting potential and contrasting explanations: (i) Tasmanian devils are not competent reservoir hosts or (ii) they are natural hosts which are able to mount a sufficient immune response against the infection. In both cases, parasitaemia might be low or absent. Additional explanations could include (iii) infections by haemoprotozoa are acute and fatal to Tasmanian devils; however, the absence of Theileria and Hepatozoon in DFTD-free individuals makes this hypothesis less likely; or, (iv) the detection of haemoprotozoa in ticks is an opportunistic/accidental finding and they are not competent vectors of these organisms. Additional sampling from DFTD-free populations and capturing seasonal variation will be important for assessing the prevalence and susceptibility of Tasmanian devils to these haemoprotozoans.
The inclusion of morphological data obtained from blood smears and the application of additional molecular tools, such as next-generation sequencing to identify co-infections [85] and genome level information [86], are vital for further work. The collection of additional tissue samples may also prove to be useful in developing diagnostic tools, as well as better understanding the lifecycle of haemoprotozoa in the host. For example, bone marrow, skeletal muscle, tongue, brain, and liver samples have been tested positive for Trypanosoma DNA, while blood samples (including blood smears) were negative [87]. While there is a body of work demonstrating that Australian mammals are hosts to a range of endemic species of Trypanosoma, Leishmania, Babesia, Theileria, and Hepatozoon [17][18][19]31], the clinical impact of these haemoprotozoa is largely unknown. This is particularly important in the context of endangered species, such as the Tasmanian devil.
The results that are presented here highlight the need for additional studies on the haemoprotozoa and broader parasite community of Tasmanian devils. It is vital that the complete biology of this species be understood in order to provide meaningful information for management and conservation strategies. Conservation strategies, including translocation, captive breeding, and establishment of insurance populations, are management strategies that have been implemented in the case of Tasmanian devils. Parasite communities are an important consideration in species recovery, highlighting the need to further study host-parasite relationships [88]. For example, while the translocation of animals could facilitate introduction of parasites into new host populations, the same translocation could dilute adaptations of the receiving populations to the local parasites. This last case is particularly relevant when the local individuals in supplemented populations are outnumbered by the translocated individuals. Immediate future work is now needed in order to document the prevalence and diversity of haemoprotozoa in devils with a focus towards understanding the clinical impacts of infection and importantly impact of co-infection with DFTD.

Conclusions
Here, we have provided the first insights into haemoprotozoa infecting the endangered Tasmanian devil. Further research is urgently needed in order to document the full haemoprotozoan diversity in the Tasmanian devil, with a focus towards understanding the clinical significance of these infections and co-infections with DFTD. The identification of T. copemani and novel members of the T. cyclops clade provide further insights into the co-evolution of trypanosomes in marsupials, showing that Australian native fauna harbour genetically unique parasitic species. Finally, the lack of detection of major haemoprotozoa genera; Hepatozoon, Leishmania, Plasmodium, and Theileria in our study opens questions regarding the overall status of these parasites in the island ecosystem of Tasmania and whether their absence might reflect specific host-parasite interactions.
Supplementary Materials: The following are available online at http://www.mdpi.com/2076-0817/9/11/873/s1 . Supplementary Table S1. Trypanosoma sequences used in phylogenetic analysis in the present study. Supplementary Table S2. Pairwise genetic similarity matrix of the Trypanosoma cyclops clade at the 18S rRNA locus. Analysis conducted over at 559 bp alignment of the V7-8 hypervariable using the Kimura Two-Parameter (K2P) method.
Funding: This study was part-funded by the Australian Research Council (LP160100200), Bayer HealthCare (Germany) and Bayer Australia. S.L.E. was supported by an Australian Government Research Training Program (RTP) Scholarship. This project was also part supported by The Holsworth Wildlife Research Endowment & The Ecological Society of Australia (awarded to S.C. and S.L.E). Fieldwork and data collection were funded by the US National Institute of Health (R01-GM126563-01), the Save the Tasmanian Devil Appeal of the University of Tasmania Foundation (Eric Guiler Grant Scheme) and the Australian Research Council (DE170101116).

Acknowledgments:
We thank Sarah Munns for extraction of DNA and Chris Peacock (The University of Western Australia) for provision of Leishmania macropodum control. We acknowledge the generous support of the large number of volunteers who assisted with fieldwork and data collection.

Conflicts of Interest:
The authors declare no conflict of interest.

Abbreviations
The following abbreviations are used in this manuscript: 18S rDNA 18S ribosomal DNA DFTD Devil tumour facial disease gDNA genomic DNA gGAPDH glycosomal glyceraldehyde phosphate dehydrogenase gene SNP single nucleotide polymorphism