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rotist, Vol. 167, 425–439, November 2016ttp://www.elsevier.de/protisublished online date 2 August 2016

RIGINAL PAPER

orphological and Phylogeneticescription of Trypanosoma noyesi sp.ov.: An Australian Wildlife Trypanosomeithin the T. cruzi Clade

driana Boteroa, Crystal Cooperb,1, Craig K. Thompsona, Peta L. Clodeb,arrie Rosec, and R.C. Andrew Thompsona

School of Veterinary and Biomedical Sciences, Murdoch University, Murdoch,Western Australia 2009, AustraliaCentre for Microscopy, Characterisation and Analysis, The University of WesternAustralia, Crawley, Western Australia 6150, AustraliaAustralian Registry of Wildlife Health, Taronga Conservation Society Australia,Bradleys Head Rd, Mosman, New South Wales 2088, Australia

ubmitted December 8, 2015; Accepted July 23, 2016onitoring Editor: Dmitri Maslov

number of trypanosome isolates from Australian marsupials are within the clade containing theuman pathogen Trypanosoma cruzi. Trypanosomes within this clade are thought to have diverged

rom a common ancestral bat trypanosome. Here, we characterise Trypanosoma noyesi sp. nov. isolatedrom the critically endangered woylie (Bettongia pencillata) using phylogenetic inferences from threeene regions (18S rDNA, gGAPDH, and CytB) coupled with morphological and behavioural observations

n vitro. We also investigated potential vectors and the presence of T. noyesi in the grey-headed flyingox (Pteropus poliocephalus). Phylogenetic analysis revealed T. noyesi and similar genotypes groupedt the periphery of the T. cruzi clade. T. noyesi is morphologically distinct both from other species ofustralian trypanosomes and those within the T. cruzi clade. Although trypanosomes were not observed

n the digestive tract of ectoparasites and biting flies collected from T. noyesi infected marsupials,abanid and biting midges tested positive for T. noyesi DNA, indicating they are vector candidates.issues from flying foxes were negative for T. noyesi. This study provides novel information on the

orphology and genetic variability of an Australian trypanosome within the T. cruzi clade.

2016 The Author(s). Published by Elsevier GmbH. This is an open access article under the CC BY-C-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

ey words: Trypanosoma noyesi; Trypanosoma; Cytochrome B; 18S rDNA; gGAPDH; Bettongia pencillata.

Corresponding author;-mail crystal.cooper@research.uwa.edu.au (C. Cooper).

ttp://dx.doi.org/10.1016/j.protis.2016.07.002434-4610/© 2016 The Author(s). Published by Elsevier GmbH. This is an open access article under the CC BY-NC-ND

icense (http://creativecommons.org/licenses/by-nc-nd/4.0/).

426 A. Botero et al.

Introduction

Trypanosomes are protozoan blood parasites thatcan infect almost all vertebrates. They are respon-sible for a number of neglected tropical diseasesincluding African sleeping sickness and Chagasdisease in humans, and the economically importantNagana in cattle. Despite numerous reports of try-panosomes infecting Australian wildlife, only eightisolated from native mammals have been describedto the species level (reviewed in Thompson et al.2014). Descriptions of trypanosomes generally relyon their morphology in blood smears taken from thevertebrate host and, more recently, their character-isation using molecular techniques. Trypanosomesisolated from Australian mammals have revealeda number of very distantly related species, highlevels of genetic diversity within species (Boteroet al. 2013; Hamilton et al. 2004; Noyes et al.1999; Paparini et al. 2011; Stevens et al. 1999),a lack of species specificity (Averis et al. 2009;Botero et al. 2013), polymorphism in blood withinand between hosts (Austen et al. 2009; Thompsonet al. 2013), and also can occur in mixed infections(Botero et al. 2013; Paparini et al. 2011; Thompsonet al. 2013). These observations highlight the dif-ficulties of classifying trypanosomes isolated fromAustralian wildlife using traditional methods, whichhas resulted in reporting of a number of superficiallycharacterised, but unnamed, isolates (Thompsonet al. 2014).

A trypanosome of particular interest is T. sp.H25 (H25) isolated from an eastern grey kangaroo(Macropus giganteus) in New South Wales, whichis in the T. cruzi clade and is most closely related toT. wauwau from bats (Lima et al. 2015). This cladeincludes T. cruzi (the pathogenic agent of humanChagas disease), the non-human pathogenic T.rangeli, and the bat-restricted species T.dionisii, T.cruzi marinkellei, and T. vespertilionis, which wereall isolated in South America. However, T. ves-pertilionis and T.dionisii have been also found inEurope. The T. cruzi clade also contains the bat-restricted species T. livingstonei, T. erneyi, and T.wauwau (Lima et al. 2012, 2013, 2015), two iso-lates found in a civet and monkey respectivelyin Africa (Hamilton et al. 2009), and T. conorhiniisolated from a rat found throughout the tropics(Hamilton et al. 2012). In Australia, isolates highlysimilar to H25 have been reported from the com-mon brush tailed possum (Trichosurus vulpecula) -T. sp. AP2011 isolates 15/17 (AP2011), and thewoylie (Bettongia penicillata), boodie (Bettongialesueur), and banded-hare wallaby (Lagostrophusfasciatus) - T. sp. H25 G8 (H25 G8) in southwest

Australia (Botero et al. 2013; Paparini et al.2011).

The discovery that H25 is at the periphery of theT. cruzi clade raised questions about the diversifi-cation and dispersion of trypanosomes worldwideand led to the “southern super-continent hypoth-esis” to explain the evolution of T. cruzi cladetrypanosomes (Stevens and Gibson 1999; Stevenset al. 1999). The southern super-continent hypoth-esis, suggested that T. cruzi clade organismsemerged from trypanosomes present in marsupi-als more than 40 million years ago when SouthAmerica, Antarctica, and Australia were joined inthe super-continent, known as Gondwana (Stevenset al. 1999). However, the vast genetic diversityof trypanosomes isolated within Australia indicatedthat unlike their hosts, they had not been isolated inAustralia since the break-up of Gondwana. An alter-native hypothesis emerged, called the “bat seedinghypothesis”, which suggested trypanosomes withinthe T. cruzi clade were originally bat parasites thatsubsequently switched into terrestrial mammalianhosts facilitated by their mobility (Hamilton et al.2012).

An estimated 3000 Latin American immigrantschronically infected with T. cruzi were estimated toreside in Australia in 2006 (Gascon et al. 2010;Schmunis and Yadon 2006), representing a pos-sible biosecurity risk to native wildlife as well ashumans (Thompson and Thompson 2015). Consid-ering the close genetic proximity of T. cruzi and H25there are concerns that the unknown vector(s) forH25 could also transmit T. cruzi. T. cruzi is trans-mitted by bugs from the family Reduviidae andwhile the risk of reduiivid bugs (generally triatominebugs) becoming established as pests in Australia islow, one species Triatoma leopoldi, was reportedin Cape York Peninsula in Queensland (Monteith1974). It is unknown if any Australian invertebrates,can transmit T. cruzi. However, it was recentlyestablished that bedbugs can facilitate transmissionof T. cruzi in a mechanical capacity indicating thatother invertebrates may share this ability (Salazaret al. 2015). There are two main concerns associ-ated with this in Australia. Local marsupials couldbecome reservoirs for T. cruzi amplifying the num-ber of parasites in vectors and hosts as they do inSouth America (Travi et al. 1952). Alternatively, theintroduction of T. cruzi into these populations couldbe catastrophic as previous research has indicatedit can cause disease and mortality in Australianmarsupials (Backhouse and Bolliger 1951).

In this study, we characterise a novel try-panosome genotype closely related to Try-panosoma sp. H25, AP2011 isolates 15/17 and

Trypanosoma noyesi sp. nov. 427

H25 G8 from the critically endangered marsu-pial, the woylie. We describe the new specimens’morphology, identify potential vectors, and inves-tigate its presence in tissues from Australianbats. Combined with phylogenetic inferencesbased in the small ribosomal subunit (18SrDNA), glyceraldehyde-3-phosphate dehydroge-nase (gGAPDH) and cytochrome B (CytB) generegions, we classify this trypanosome as a newspecies.

Results

Isolation of Woylie Trypanosomes inCulture

A single blood sample from a woylie (WC6218)cultured in TRPMI 1640 (Roswell Park MemorialInstitute 1640 supplemented with tryptose) mediumand maintained at 28 ◦C resulted in the isolation oftrypanosomes. Subsequently, trypanosomes fromthis culture were able to grow in a number ofbiphasic cultures containing BHI (brain-heart infu-sion) blood medium as a solid phase, and eitherRPMI 1640 (Roswell Park Memorial Institute), LIT(liver infusion tryptose), or Grace’s media as a liq-uid phase. However, TRPMI 1640 best supportedits continuous cultivation in culture either with BHIor alone.

DNA Sequencing, PhylogeneticRelationships, and Genetic DistancesBased on 18S rDNA, gGAPDH, and CytBGenes

Comparison of sequences obtained from the 18SrDNA (1,441 bp) and gGAPDH (810 bp) genesusing generic primers confirmed the new try-panosome isolated from woylie blood was a novelgenotype closely related to the trypanosomesH25, AP2011 15/17, and H25 G8, from Australia.The new isolate differed at 1/3, 5/24, and 4/17(18S rDNA/gGAPDH) nucleotide sites from H25,AP2011 15/17, and H25 G8 respectively. Thewoylie isolate was more closely related to try-panosomes from outside Australia in the T. cruziclade than it was to other Australian trypanosomesisolated from marsupials such as T. copemani, T.gilletti, T. irwini, T. sp ABF and T. vegrandis as pre-viously indicated (Botero et al. 2013; Noyes et al.1999; Paparini et al. 2011) (Figs 1 and 2). Sim-ilar tree topologies were obtained from the 18SrDNA, gGAPDH (Figs 1 and 2), and CytB (Supple-mentary Material Fig. S1) phylogenetic analyses,

where the new isolate was within the well-resolvedT. cruzi clade with eight other Trypanosoma speciesincluding: T. cruzi, T. cruzi marinkellei, T. erneyi, T.dionisii, T. wauwau, T. rangeli, T. vespertilionis, andT. conorhini. The 18S rDNA phylogeny showed T.wauwau formed a sister clade to the one containingthe new isolate, H25, AP2011 15/17, and H25 G8(Fig. 1). However, the gGAPDH phylogeny showedT. wauwau was a sister clade to the one containingT. rangeli, T. conorhini, and T. vespertilionis, and notto the clade containing the new isolate (Fig. 2).

The genetic distance between the new iso-late and T. cruzi was larger when compared withT. rangeli and bat trypanosomes (SupplementaryMaterial Table S1). Phylogenies based on 18SrDNA and gGAPDH sequences showed that previ-ous isolates from Australian wildlife (H25, AP201115/17, and H25 G8) clustered tightly togetherwith the new isolate from woylie blood with shortgenetic distances (∼0.02), similar to those obtainedbetween different isolates of T. erneyi (∼0.01) andT. lewisi (∼0.01) (data not shown). The position-ing in the phylogenetic trees and the short geneticdistances between the new isolate and previousisolates from Australian wildlife (H25, H25 G8, andAP2011 15/17) strongly support their classificationas a new Trypanosoma species.

Species-specific PCRs

DNA extracted from the in vitro cultures of the newisolate tested positive to 18S rDNA clade C species-specific PCR confirming the genetic similarity withH25, AP2011 15/17, and H25 G8. WC6218 cul-ture was negative for T. copemani and T. vegrandisspecies specific PCR’s indicating the absence ofmixed infections with these Trypanosoma spp.,which are the only other species previously foundinfecting woylies. All 57 tissue samples from 31 batswere negative when screened using the clade Cspecies-specific PCR.

Morphology of the New Isolate in vitro

Trypanosomes were not seen in peripheral bloodsmears taken from the woylie WC6218, from whichthe new trypanosome was isolated in culture.Live-cell phase contrast microscopy of the newisolate in vitro demonstrated high levels of pleomor-phism (Fig. 3). The different morphological formsobserved in vitro ranged from epimastigotes, try-pomastigotes, and spheromastigotes to severalintermediate forms of epimastigotes and trypo-mastigotes (Fig. 4, Supplementary Material Fig.S2). Epimastigotes were the predominant form inculture (Fig. 3B). They divided intensely by binary

428 A. Botero et al.

Figure 1. Phylogenetic analysis of the relationships between Trypanosoma spp. and Trypanosoma noyesi sp. n.based on ribosomal subunit (18S rDNA) sequences. The phylogenetic tree was constructed using the Bayesianmethod. Bayesian posterior probabilities are shown at nodes. Trypanosoma noyesi sp. n. sequences isolatedin this study are highlighted. Scale bar indicates number of substitutions per site.

fission forming rosettes of different sizes (Fig. 3C).Epimastigotes were stout and wide at the anteriorthird of the cell tapering toward the posterior, andwere 21 ± 2 �m in length (n = 10). They appearedrigid and only seemed to move on close observa-tion. Epimastigotes attached to each other via around, extended flagellar sheath (EFS), visible as adark circle at the anterior end of the cell in the prox-imity of a short flagellum on smears stained withDiff-Quik (Fig. 3B). Diff-Quik stained specimensrevealed the nucleus and kinetoplast were closeto the anterior end of the trypanosome (Fig. 3C).

Most epimastigotes exhibited a very short flagellumextending past the end of the EFS (Figs 4E, 5A). Asecond epimastigote form was visible (15 ± 1 �min length, n = 10) with a long flagellum (20 ± 2 �min length, n = 10) that extended back towards theposterior end after exiting the flagellar pocket with-out the EFS (Figs 4F, 5B). This second form movedin a circular motion and was observed in rosettes.Epimastigotes generally possessed a smooth sur-face. However, on occasions the surface of thecells exhibited a budding of exo-vesicles to varyingdegrees (Fig. 5A). During epimastigote division, the

Trypanosoma noyesi sp. nov. 429

Figure 2. Phylogenetic analysis of the relationships between Trypanosoma spp. and Trypanosoma noyesi sp.n. based on glyceraldehyde 3-phosphate dehydrogenase (gGAPDH) sequences. The phylogenetic tree wasconstructed by the Bayesian method. Bayesian posterior probabilities are shown at nodes. Trypanosoma noyesisp. n. sequences isolated in this study are highlighted. Scale bar indicates number of substitutions per site.

anterior end of the epimastigote swelled as the kine-toplast and nucleus divided. The cell membraneremained intact until nucleus and kinetoplast divi-sion was complete and the organism had reachedfull length. Separation of the membrane beganat the posterior of the trypanosome and finishedwith the separation of the flagellar sheath (Fig. 5E,F). Division of epimastigotes in rosettes showedthe appearance of smaller spherical trypanosomes(Fig. 4C). Dividing rosettes attached to the surface

of cells (Fig. 5E) or to the glass coverslip (Fig. 5F),by the spreading of the flagellar sheath acrossthe substrate when grown with mammalian cellsat 37 ◦C. Multiple nuclei were observed in somedividing epimastigotes (Fig. 4I).

Two types of trypomastigotes were observed.At 37 ◦C a slender, long morphological form wasseen occasionally that was 25 ± 1 �m (n = 10)in length (Figure 6D), while a smaller, stumpyform was seen in cultures at 28 ◦C, these were

430 A. Botero et al.

Figure 3. A. Live-cell phase contrast microscopy image of Trypanosoma noyesi sp. n. in culture exhibitingvarious morphotypes. E1 = epimastigotes exhibiting extended flagellar sheath. E2 = epimastigote exhibitingemergent flagellum. S = spheromastigote. T = putative trypomastigote. Black and white dots representingglycosomes and acidocalcisomes respectively are observed in the cytoplasm (arrowhead). B. Morphology ofTrypanosoma noyesi sp. n. epimastigotes in the first passage from woylie blood in vitro. Staining with Diff-Quikand viewed by light microscopy. C. Morphology of Trypanosoma noyesi sp. n. rosettes in the first passage fromwoylie blood in vitro. Staining with Diff-Quik and viewed by light microscopy.

8 ± 1 �m- in length (n = 10) (Fig. 5A, D). An inter-mediate stage was observed exhibiting a nucleusand kinetoplast that was closer together and slightlywider (Fig. 4G) than the long slender trypomastig-ote (Fig. 4H). All trypomastigotes exhibited anelongated nucleus in the middle of the cell, anda compacted and rounded kinetoplast at the pos-terior end of the cell. Trypomastigote forms werefast moving and possessed a conspicuous undu-lating membrane that extended from the posteriorend of the cell and terminated in the flagellum.

Occasionally, stumpy trypomastigotes appeared toform nests (rosettes) in vitro, although scanningelectron microscopy (SEM) revealed these werenot dividing (Fig. 5D). Small spheromastigotes of2 ± 0.5 �m in length (n = 10) were observed by SEMbut they were too small to be resolved as try-panosomes in light microscopy.

Longitudinal and transverse TEM sections ofepimastigotes grown in vitro revealed a cellularstructure typical of epimastigotes of other Try-panosoma species (Fig. 6A, B). The nucleus and

Trypanosoma noyesi sp. nov. 431

Figure 4. Representation of morphotypes of Trypanosoma noyesi sp. n. in vitro. A. spheromastigote. B. stumpytrypomastigote. C. small dividing epimastigotes. D. common dividing epimastigotes. E. epimastigote. F. epi-mastigote with long flagellum. G. wide slender trypomastigote. H. slender trypomastigote. I. multiple fusion ofepimastigotes.

nucleolus were well defined. The kinetoplast was500 nm in length, 100 nm in width (Fig. 6C) and wasseen enclosed within the mitochondrial membrane.The posterior region of the basal body and the fila-ments that connect the kinetoplast to the basal bodywere observed attached to the kinetoplast (Fig. 6C).The kinetoplast was inside the mitochondrionmembrane making the mitochondria-kinetoplastcomplex, which was observed in a transverse sec-tion (Fig. 6B) extending along the dorsal surfaceof the trypanosome. The flagellar axoneme had a9 + 2 arrangement of microtubules (Fig. 6B). InSEM images the epimastigotes appeared to some-times lack a flagellum (Fig. 5E). However, it wasrevealed in TEM micrographs that the flagellum wasinside the sheath as the axoneme was visible insidethe EFS (Fig. 6B). Acidocalcisomes, reservosomesand glycosomes were also observed (Fig. 6A).Black and white dots indicating the presence ofglycosomes, reservosomes and acidocalcisomeswere visible in the cytoplasm of live cells (Fig. 3).

Trypanosomes Co-cultured with Cells

Stumpy and slender trypomastigotes of the new iso-late were unable to invade or develop within VEROcells in vitro. Both forms were seen swimming freelyin the medium and occasionally attached to cells bythe posterior end (Supplementary Material Fig S2).

Possible Vector

PCR results demonstrated that none of the 42fleas from the family Pulicidae and Stephanocir-cidae, 568 lice from the family Boopiidae, 1769mosquitoes from the family Culicidae, and 1456ticks from the family Ixodidae (pooled into 30, 140,86, and 295 samples respectively) were positivefor the new species. However, PCR and furthersequencing of the 18S rDNA gene showed thatof the 302 tabanid flies examined (family Taban-idae), 31 of the 102 pooled samples (30%) werepositive for the new species. Moreover, of the 578biting midges examined (family Ceratopogonidae),

432 A. Botero et al.

Figure 5. Scanning electron micrographs showing morphology of Trypanosoma noyesi sp. n. in vitro. A. epi-mastigotes grown at 28 ◦C exhibit an exposed flagellar sheath (left) and exo-vesicles (white arrow), and stumpytrypomastigote (right). B. epimastigote exhibiting long flagellum and short flagellum (left). C. slender trypo-mastigote grown at 37 ◦C. D. stumpy trypomastigotes forming a nest. E. rosette attached to a mammalian cellgrown at 37 ◦C. F. rosette attached to a glass coverslip grown at 37 ◦C, the spreading of the flagellar sheath isvisible (black arrowhead).

12 of the 21 pooled samples (57%) were positivefor the new isolate. Only nine 18S rDNA sequenceswere obtained from tabanid flies. Sequence align-ments of these nine isolates from tabanid flies andthe isolate from culture showed some polymor-phisms between them, and all grouped togetherin the 18S rDNA phylogeny (Fig. 1). CytB andgGAPDH sequences could not be obtained dueto difficulties with the low sensitivity of the CytBPCR, and due to unspecific binding of the gGAPDHprimers, which amplified DNA from other Try-panosomatids that infect insects (data not shown).Assuming that a positive pooled sample containedat least one positive insect, the minimum esti-mated proportion of tabanid flies and biting midges

positive for the new species was 10% and 2%respectively.

Taxonomy

Taxonomic positioning: Phylum EuglenozoaCavalier-Smith, 1981, class Kinetoplastea Honig-berg, 1963 emend. Vickerman, 1976, subclassMetakinetoplastina Vickerman, 2004, order Try-panosomatida (Kent, 1880) Hollande, 1952, familyTrypanosomatidae Doflein, 1951, and genus Try-panosoma Gruby, 1843.

Generic assignment: This new isolate fromthe woylie was placed in the genus Trypanosomabecause it exhibits cell morphotypes including

Trypanosoma noyesi sp. nov. 433

Figure 6. Transmission electron micrographs of Trypanosoma noyesi sp. n. A. longitudinal section showing thekinetoplast (white arrowhead), flagellum, acidocalcisomes (black arrowhead), glycosomes (white asterix), andreservosomes (black asterix). B. transverse section revealing the flagellum, axoneme, mitochondrion (arrow-head), and extension of the flagellar sheath (asterix). C. kinetoplast exhibiting kDNA enclosed in a membrane(arrowhead), which is attached to the basal body (asterix).

trypomastigotes and epimastigotes, and was foundto be phylogenetically affiliated with other try-panosome species. All phylogenies demonstratedthe new species was within the monophyletic groupof trypanosomes.

Species diagnosis: the new species describedin this study, can be unequivocally distinguishedfrom all other species of trypanosomes by itsunique gene sequences, and phylogenetic posi-tioning. The sequences from all genotypes withinthe species found in the woylie and tabanidsflies are deposited in GenBank under the fol-lowing accession numbers: T. noyesi WC6218(18S rDNA: KU354263, gGAPDH: KU354264,CytB: KU354265), T. noyesi TF27, TF212, TF857,TF1059, TF1160, TF1261, TF1362, TF2170 andTF3382 (18S rDNA: KX008312 - KX008320), T.noyesi H25 (18S rDNA: AJ009168, gGAPDH:AJ620276), T. noyesi AP2011 15/17 (18S rDNA:JN315381 – JN315381, gGAPDH: JN315395 –JN315396), and T. noyesi H25 G8 (18S rDNA:KC753537, gGAPDH: KC812988). T. noyesi epi-mastigotes are characterised by the attachment toeach other via a round, extended flagellar sheath,visible as a darkly staining circle at the anterior end

of the cell in the proximity of a very short flagellum.Two types of trypomastigotes are present, a slen-der form of about 25 ± 1 �m in length, and a stumpyform of about 8 ± 1 �m in length.

Trypanosoma noyesi sp. nov. Botero andCooper 2016

Type material: hapanotype Romanowsky-typestain ‘Diff-Quik’ smears of first passage in vitrofrom WC6218 blood and cryo-vials of WC6218deposited in liquid nitrogen cryobanks at MurdochUniversity.

Type host: brush-tailed bettong or woylie (Beton-gia penicillata)

Type location: Upper Warren region in WesternAustralia (34.2333◦ S, 116.1333◦ E)

Additional hosts: Eastern grey kangaroos(Macropus giganteus), brush-tailed possums(Trichosurus vulpecula), burrowing bettongs orbodies (Bettongia lesueur), and banded-harewallabies (Lagostrophus fasciatus) (Botero et al.2013; Noyes et al. 1999; Paparini et al. 2011).

Vector: Unknown.Etymology: Trypanosoma noyesi sp. nov. is

named in recognition of Dr. Harry Noyes who orig-inally isolated and described H25.

434 A. Botero et al.

Discussion

Evolutionary Relationships of T. noyesiwithin the T. cruzi Clade

Comparisons of phylogenetic trees inferred fromthe 18S rDNA, gGAPDH and CytB gene regions,demonstrate similar topologies and confirmed T.noyesi as a new species. The 18S rDNA phylogenyshowed T. noyesi isolates were positioned at theperiphery of the T. cruzi clade together with T.wauwau isolated from Pteronotus bats (Lima et al.2015). Previous studies using 18S rDNA have pos-itioned T. livingstonei, a trypanosome from Africanbats, in the most basal position of the T. cruziclade (Lima et al. 2013, 2015). The present studyfailed to demonstrate this, and showed T. living-stonei fell outside of the T. cruzi clade. Shorterfragments of 18S rDNA used in the present studycould have reduced the information required toshow this relationship. However, our gGAPDH phy-logeny positioned T. livingstonei within the T. cruziclade. T. noyesi from the woylie, H25 from the kan-garoo, and AP2011 15/17 from possums exhibitshort genetic distances between them indicatingthey are the same species exhibiting intraspecificdiversity, which has been observed in other Aus-tralian trypanosomes including T. copemani and T.vegrandis (Botero et al. 2013). The lack of hostspecificity in T. noyesi could be due to multiple hostswitching events over time, and could account forthe observed intraspecific diversity. However, asdetails of the entire host range of T. noyesi andother Australian trypanosomes remain unknown,the mechanism facilitating the host-switching isuncertain.

It has been demonstrated that most Try-panosoma species within the T. cruzi clade arepresent in bats, including T. cruzi (da Silva et al.2009; Lima et al. 2015). This study did not iden-tify the presence of T. noyesi in the Australian greyheaded flying fox. Within Australia four species oftrypanosomes have been reported infecting batsincluding, T. pteropi in flying foxes (Pteropus alecto)(Breinl 1913; Johnston 1916), and T. hipposideri inthe dusky round-leaf bat (Hipposideros albanen-sis) (Mackerras 1959). However, the descriptionof these species was based on their morphol-ogy in blood alone and nothing is known abouttheir life cycle, host-parasite interactions, or phy-logenetic relationships. T. vegrandis was identifiedinfecting the Gould’s wattled bat (Chalinolobusgouldii), lesser long-eared bat (Nyctophilus geof-froyi), little red flying fox (Pteropus scapulatus), andblack flying fox (Pteropus Alecto) using molecular

techniques (Austen et al. 2015). More recently, T.teixeirae has been described from the Australian lit-tle red flying fox (Pteropus scapulatus), which wasin the T. cruzi clade (Barbosa et al. 2016). This indi-cates more extensive sampling from the blood andtissues of other Australian bats are needed to deter-mine the possible origin and evolutionary history oftrypanosomes within the T. cruzi clade.

T. noyesi Morphology and Ultrastructurein vitro

The morphology of T. noyesi is different to othertrypanosomes within the T. cruzi clade demonstrat-ing the morphological diversity in this group, whichis well documented (Hoare 1972; Lima et al. 2013,2015). Notably, T. conorhini and T. rangeli were orig-inally placed in different subgenera based on theirmorphology and behaviour (Hoare 1972), and weremoved into the T. cruzi clade following molecularcharacterization (Hamilton et al. 2007). T. noyesiis also morphologically distinct from other Aus-tralian marsupial trypanosomes isolated in vitro,which includes T. copemani (Austen et al. 2009),T. sp. ABF (Hamilton et al. 2004), and T. thylacis(Mackerras 1959). T. noyesi has not been reportedin blood smears, despite several studies identify-ing the presence of T. noyesi DNA in the blood,which may indicate low parasitaemia (Botero et al.2013; Noyes et al. 1999; Paparini et al. 2011).However, considering that trypanosomes isolatedfrom Australian mammals exhibit polymorphism inblood within and between hosts (Austen et al. 2009;Thompson et al. 2013), and occur in mixed infec-tions (Botero et al. 2013; Paparini et al. 2011)describing species based on blood stream mor-phology is unreliable. Traditional taxonomy wasnot designed to cater for protozoan biology, whichhas resulted in a number of inconsistencies in thecurrent records of trypanosomes characterised inAustralia (Cooper et al. 2016; Thompson et al.2014; Votypka et al. 2015). Advances in high res-olution microscopy and in vitro cultivation haveimproved the ability to characterise trypanosomesby providing a stable environment to investigate try-panosome morphology and differentiation.

The dominant morphological form of T. noyesiin vitro is the epimastigote and not the promastigoteas described by Noyes et al. (1999). Promastig-otes are a common culture form of Leishmania spp.and monoxenous trypanosomatids. The epimastig-otes of T. noyesi appear similar to promastigotesin light microscopy images due to the rigidity oftheir structure. In promastigotes, the flagellum exitsthe trypanosome at the direct center of the anterior

Trypanosoma noyesi sp. nov. 435

end from the flagellar pocket. However, the flagel-lum is visible along the side of the trypanosomesin T. noyesi SEM micrographs. A small number of‘epimastigotes’ with a short emergent flagellum and‘nectomonads’ with long flagellum were also pre-viously reported (Noyes et al. 1999). However, thedata collected from optical and electron microscopyin this study support the proposition that these areall epimastigotes with varying lengths of flagella.The structure observed protruding at the end of theepimastigote of T. noyesi is referred to as an exten-sion of the flagellar sheath and links one cell toanother via junction complexes called hemidesmo-somes (Brooker 1970; Noyes et al. 1999). T. freitasi,found in opossums has been noted to have formsof unusual epimastigotes that produce an extendedflagellar sheath that occur in the lumen of the scentgland and not in an invertebrate host (Thomaz et al.1990). T. freitasi uses the extended flagellar sheathto attach to the outside of the cell membrane asa means of support whilst dividing (Thomaz et al.1990). In T. noyesi the sheaths were observedforming rosettes and attaching to cell membranesand glass coverslips indicating they are also usedto attach to cells. T. cruzi epimastigotes in vitro growin rosettes similar to T. noyesi, but they move veryfast lacking the adhesive sheaths and instead linkvia their flagella (Hoare 1972). T. noyesi epimastig-otes exhibited exo-vesicles budding on the surfaceof the cell in varying degrees. In Leishmania spp.,exo-vesicles occur after being exposed to changesin temperature during inoculations from the sand flyto the mammalian host, which induces a release ofproteins that changes the morphology of the par-asite and is speculated to deter macrophages inthe host (Hassani et al. 2011). Within the samesample T. noyesi exhibited both smooth and exo-vesicle producing surfaces, ruling out the likelihoodthat these are artefactual. The exo-vesicles couldbe an indicator of the health of the trypanosomesand further investigation is required in order tounderstand their presence and nature. The stumpytrypomastigotes present in vitro were described inT. sp H25 by Noyes et al. (1999), and in a numberof other trypanosomes referred to as T. lewisi-liketrypanosomes in the stercorarian subgenus Her-petosoma Doflein, 1901 (Hoare 1972) including T.wauwau (Lima et al. 2015). They occur in T. lewisiand T. zapi (two rodent trypanosomes) in vitro iso-lated from the flea rectum and maintained in vitroat 28 ◦C. The other trypomastigote observed for thefirst time was the slender trypomastigote, whichresembled other in vitro trypomastigotes, includ-ing those of T. cruzi, T. wauwau, and T. copemani(Botero et al. 2013; Hoare 1972; Lima et al. 2015).

In T. cruzi this morphological form is observedin the mammalian host and invades a number ofhost cells (Hoare 1972). T. noyesi was unable toinfect LLCMK1 cells (Rhesus monkey kidney cells)in vitro and mice in vivo in a previous study (Noyeset al. 1999). In the present study higher numbersof slender or stumpy trypmastigotes of T. noyesiwere co-cultured with African green monkey kidneyepithelial cells (VERO). T. noyesi was not observedinside cells, although many members of the T. cruziclade do not infect cells. Whether non-intracellulartrypanosomes in the T. cruzi clade have lost thisability or never developed it remains unknown.Considering that intracellular localisation has beenobserved in T. copemani isolated from the woylie(Botero et al. 2013), and suggestions that Aus-tralian trypanosomes may be pathogenic to theirhosts (Botero et al. 2013; McInnes et al. 2011), theimpact of trypanosomes on local wildlife remainsan area that warrants further study.

Possible Vector

The aim of this preliminary vector investigationwas to screen for trypanosome DNA from a largenumber of haematophagous arthropods in orderto identify putative vector candidates of T. noyesi.Samples of a relatively large number of insects andarachnids were pooled for DNA extraction, assum-ing that a positive pooled sample represented(at least) a single positive individual. Therefore,the minimum prevalence of positive arthropodshas been reported, with the actual prevalenceof insects that ingested T. noyesi possibly beinghigher. Tabanid flies and sand-flies are both vectorsof trypanosomes, including Tabanus spp. trans-mitting T. theileri, and Lutzomyia spp. transmittingT. leonidasdeanei (Hoare 1972). T. noyesi wasidentified from at least 10.3% of the tabanid fliessampled. A limitation of the methodology used herewas that a positive arthropod sample could notbe confirmed as a vector, which has an estab-lished trypanosome infection within its gut (Seblovaet al. 2014) as it could represent ingested orpartly-digested trypanosomes from a blood meal.It was outside the scope of this study to iden-tify the large number of insects to genus andspecies level. Therefore, it is not possible to com-ment further on the species that may be importantfor transmission of T. noyesi. There is a lack ofknowledge regarding the dispersion and vectorialcapacity of invertebrates in Australia (Thompsonand Thompson 2015). Although, considering thepresence of T. noyesi on both west and east

436 A. Botero et al.

Australia the vector is most likely a commonlyoccurring invertebrate.

Conclusions

This study describes an Australian trypanosome,T. noyesi, based on morphology and phylogeneticposition, providing insight into the relationshipsand diversity present within the T. cruzi clade. Inthe past a number of trypanosomes have beendescribed to species based on descriptions of theirmorphology in blood smears taken from the ver-tebrate host, or partial genetic data, especially inAustralia (Mackerras 1959; McInnes et al. 2011).Future studies are required to fill these gaps tounderstand the true nature of host-parasite inter-actions and evolutionary relationships in Australiantrypanosomes. In Australia where trypanosomesexhibit low parasitaemia, phenotypic differencesin bloodstream trypomastigotes, and no speciesspecificity, we have shown that in vitro culturesare essential when describing new species. Futurestudies are required to investigate the vectorial can-didates of T. noyesi as this is an area that requiresurgent attention due to the biosecurity risks involvedand the severe lack of knowledge on trypanosomevectors in Australia.

Methods

Isolation of woylie trypanosomes in vitro: Blood sampleswere collected from woylies previously known to be infected withT. noyesi by PCR in the Upper Warren region in Western Aus-tralia (34.2333◦ S, 116.1333◦ E) (Botero et al. 2013; Thompsonet al. 2013). Wildlife sampling was carried out under MurdochUniversity animal ethics approval permit numbers W2350-10,RW2659-14, and DEC animal ethics approval permit numberDECAEC/52/2009. Woylies were trapped in small cage traps(20 cm × 20 cm × 56 cm) baited with a mixture of rolled oats andpeanut butter. Traps were placed at set intervals (usually 200 m)along tracks in the study site. Blood was collected from thelateral caudal vein and blood smears were made using approx-imately 20 �l of blood. Cultures were established by inoculationof approximately 50 �l of peripheral blood into minicollect tubes(greiner bio-one) containing TRPMI 1640 supplemented with10% fetal calf serum and 1% Penicillin-Streptomycin. Tubeswere checked every week by light microscopy for motile try-panosomes. When trypanosomes were seen for the first time,trypanosomes were sub-cultured in 25 cm2 tissue culture flasksor NuncTM cell culture tubes (Thermo scientific) with biphasicmedium containing BHI medium, agar, gentamicin, and 10%defibrinated horse blood as a solid phase, and either TRPMI1640, LIT, or Grace’s media as a liquid phase. Cultures wereleft for one to two weeks and then the supernatant was removedand replaced with new liquid medium. Optimal growth require-ments in liquid medium alone were determined using RPMI1640, TRPMI 1640, LIT, and Grace’s media. Cultures were

maintained in liquid media by successive passages every threedays at 28 ◦C and were deposited in liquid nitrogen cryobanksat Murdoch University.

Cell infection: Monolayers of VERO cells (kidney epithelialcells) were trypsinised and seeded onto tissue culture-slides (16-wells) at a concentration of 1.5 × 104 cells/ml. After24 hours, the media was discarded to remove non-adherentcells and 100 �l of parasite suspension containing 1.5 × 105

parasites/ml was added to each well (1:10 cell/parasite ratio).Cultures from the stationary phase containing numerous trypo-mastigote forms were used to infect cells. Slides were incubatedat 37 ◦C and 5% CO2. At 24 hours post-infection, the super-natant was discarded and some slides were washed threetimes with 1 x phosphate buffer solution (PBS) to removenon-adherent parasites. Coverslips were removed and culture-slides were air-dried and stained with the commercial stain‘Diff-Quik’ for examination of intracellular parasites or attachedtrypanosomes by light microscopy. Experiments were repli-cated three times on separate occasions.

Optical, scanning and transmission electronmicroscopy: Blood smears taken from woylies and smears oflogarithmic and stationary phase from trypanosomes grownin culture were stained with Diff-Quick, and the morphologyof trypanosomes examined by light microscopy. Live-cellphase contrast microscopy of parasites was performed usinga Nikon A1 confocal laser microscope with an attached tokaihit incubation chamber. For scanning electron microscopy(SEM), culture forms were fixed in a 1:1 mixture of 5%glutaraldehyde in 1 x PBS: cell culture medium (pH 7.2) forone hour and then washed with 1 x PBS three times beforebeing stored in fresh 2.5% glutaraldehyde in 1 x PBS at 4 ◦C.Samples were mounted on poly-L-lysine coated coverslips,progressively dehydrated through a series of ethanol solutionsusing a PELCO Biowave microwave, and critical point dried aspreviously described (Edwards et al. 2011). Coverslips weremounted on stubs with adhesive carbon, coated with 2 nmplatinum (Pt) and 10 nm of carbon, and imaged at 3 kV usingthe in-lens secondary electron detector on a Zeiss 55VP fieldemission SEM. For transmission electron microscopy (TEM),trypanosomes were similarly fixed and stored. All subsequentprocessing was performed in a PELCO Biowave microwave,where samples were post-fixed in 1% OsO4 in PBS followed byprogressive dehydration in ethanol then acetone, before beinginfiltrated and embedded in Procure- Araldite epoxy resin.Sections 100–120 nm-thick were cut using a diamond knifeand mounted on copper grids. Digital images were collectedfrom unstained sections at 120 kV on a JEOL 2100 TEM fittedwith a Gatan ORIUS1000 camera.

Ectoparasites and haematophagous insect collectionand pooling: Ticks were removed from the skin of woyliesusing fine forceps; lice and fleas were collected from the furusing a fine toothcomb. Haematophagous insects were caughtusing both Marris style Malaise and Nzi traps; traps were baitedwith octenol lures, and contained a collection pot filled with70% ethanol. Free-living haematophagous insects analysedincluded tabanids (also called march-flies) (Tabanidae), bitingmidges (Ceratopogonidae) and mosquitoes (Culicidae). All ofthese haematophagous insects, along with the ectoparasiteswere removed from the woylies and stored in 70% ethanol forfurther identification and DNA extraction. Collected ectopar-asites and haematophagous insects were removed from the70% ethanol solution, allowed to air dry, and identified morpho-logically to at least family level. Each family of ectoparasitescollected from an individual woylie was pooled into separate1.5 ml microcentrifuge tubes, with a maximum of fifteen simi-lar ectoparasites per tube. Each family of dipterans was pooled

Trypanosoma noyesi sp. nov. 437

into a separate 1.5 ml microcentrifuge tube, with a maximum of30 insects per tube.

Collection of bat tissue samples: Tissue samples fromgrey-headed flying foxes (Pteropus poliocephalus) from NewSouth Wales were collected from sick-euthanised animals thatwere presented to Taronga Zoo for treatment, and from deadanimals sent for necropsy. Sick animals were euthanised dueto very poor body condition, severe injury, and poor progno-sis for return to the wild. A total of 57 tissue samples werecollected from 31 carcasses and at least two of the followingtissues were collected from each animal: kidney, liver, spleen,heart, and brain. All tissue samples were extensively washedwith 1 x PBS and stored in 100% ethanol for DNA isolation.

DNA extractions: Genomic DNA of trypanosomes isolatedin vitro and bat tissues were obtained using the QIAamp bloodand tissue DNA MiniKit (Qiagen, Hilden, Germany) accordingto the manufacturer’s instructions. DNA from the arthropodswas extracted using the Wizard® Genomic DNA PurificationKit as per the protocol for animal tissue extraction (Promega,Wisconsin USA), except that the arthropods were homogenisedin 600 �l of nucleic lysis solution, and then incubated at 65 ◦C for12 hours rather than the recommended 15–30 mins. ExtractedDNA from blood and arthropods was eluted in 60 �l of DNArehydration solution and stored at −20 ◦C. Negative controls(containing only reagents) were included.

Trypanosome species detection by PCR using genericTrypanosomatid primers and sequencing: Sequences wereobtained for the woylie trypanosome isolate (WC6218) from the18S rDNA, gGAPDH and CytB gene regions. Nine 18S rDNAsequences were also obtained from trypanosomes from tabanidflies. All sequences were deposited on the Genbank and acces-sion numbers are available in Supplementary Table 2. Anapproximately 1.4 kb fragment of the variable region of the 18SrDNA gene was amplified and sequenced using nested PCRswith generic trypanosomatid primers as described previously(McInnes et al. 2009). A second fragment of approximately810 bp of the gGAPDH gene was amplified and sequencedusing modified hemi-nested reactions previously described(Botero et al. 2013). Finally, a third fragment of approximately710 bp of the CytB gene was amplified using the primersL.cyt-S: 5′-GGTGTAGGTTTTAGTYTAGG -3′ and: L.cyt-R: 5′CTACAATAAACAAATCATAATATRCAATT-3′ (Kato et al., 2010).The amplification reactions were performed in a final volume of25 �l containing 0.2 units of Taq polymerase, 200 �M of eachdNTP, 200 nM of each primer, 1.5 mM MgCL2, and 1 �l of DNAtemplate. Amplification was performed in a PT100 thermocy-cler (MJ Research) and consisted of a denaturation step at94 ◦C for 5 min, followed by 35 cycles of 30 secs at 94 ◦C, 30 sat 52 ◦C, 50 s at 72 ◦C, and a final extension step at 72 ◦C for7 min. PCR products were run on a 1.5% agarose gel stainedwith SYBR safe (Invitrogen, USA), and visualised with a darkreader trans-illuminator (Clare Chemical Research, USA). PCRproducts were purified using Agencourt AMPure PCR Purifi-cation system following the manufacturer’s instructions andsequenced using an ABI PrismTM Terminator Cycle Sequenc-ing kit (Applied Bio-systems, California, USA) on an AppliedBio-system 3730 DNA Analyser at the Western Australian StateAgricultural Biotechnology Centre. Sequences were alignedand edited using Sequencher version 5.0.

DNA sequence alignments and phylogenetic inferences:18S rDNA, gGAPDH, and CytB sequences were aligned usingMUSCLE (Edgar 2004). Three different alignments were cre-ated for phylogenetic inference. First, a 1,441 bp sequence ofthe 18S rDNA gene was aligned with 57 Trypanosoma spp.sequences representing all known trypanosome clades, andfive other trypanosomatid sequences for use as outgroups.

Secondly, a 810 bp gGAPDH sequence was aligned with 59 Try-panosoma spp. sequences representing all major trypanosomeclades, and five other trypanosomatid sequences for use asoutgroups. Thirdly, a 710 bp CytB sequence was aligned with23 Trypanosoma spp. sequences representing all the T. cruziclade and using T. lewisi as an outgroup. All sequences wereobtained from GenBank (Supplementary Material Table S2).The new trypanosome sequences and T. lewisi CytB sequencewere deposited on GenBank. A Bayesian analysis was run inMr Bayes v. 3.1.2 (Ronquist and Huelsenbeck 2003). jModel-Test 2.1.1 was used to find the most appropriate nucleotidesubstitution model (Posada 2008). The model chosen was: theGTR+I+G for all alignments. The posterior probability distribu-tion was estimated using The Markov chain Monte Carlo, whichwas run for 10000000 generations, until the mean standarddeviation of split frequencies was lower than 0.01 and con-firmed by Mr Bayes potential scale reduction factor values closeto 1.00. Trees were sampled every 1000th generations andthe first 2500 trees (first 250000 generations), which usuallypresent very low likelihood values, were discarded as burn-in.

Trypanosome species confirmation by PCR using 18SrDNA species-specific primers: All positive samples werealso screened using Clade C species-specific PCR primers(Botero et al. 2013) that amplify H25, AP2011 15/17, and H25G8 DNA. To investigate the presence of mixed infections, sam-ples were also screened with Clade A species-specific primersthat amplify T. copemani DNA, and Clade B species-specificPCR primers that amplify T. vegrandis DNA (Botero et al. 2013).

Genetic distances: Estimates of evolutionary divergence inthe CytB, and 18S rDNA sequences were obtained between T.noyesi and the Trypanosoma spp. within the T. cruzi clade. Anal-yses were conducted using the Maximum Composite Likelihoodmodel in MEGA6 (Tamura et al. 2013). The analysis involvednucleotide sequences from seven and eight spp. All positionscontaining gaps and missing data were eliminated. There werea total of 412 positions for CytB and 1.148 for 18S rDNA in thefinal dataset. Evolutionary analyses were conducted in MEGA6.

Acknowledgements

Thanks are extended to Amy Northover and SarahKeatley for their invaluable help with the supplyof blood samples, and Department of Parks andWildlife. The authors acknowledge the facilities,and the scientific and technical assistance of theAustralian Microscopy and Microanalysis ResearchFacility at the Centre for Microscopy, Characterisa-tion & Analysis (CMCA), the University of WesternAustralia, a facility funded by the University, Stateand Commonwealth Governments. This researchwas supported with funding from the AustralianResearch Council and the Western Australian Gov-ernment’s state NRM program.

Appendix A. Supplementary Data

Supplementary data associated with this arti-cle can be found, in the online version, athttp://dx.doi.org/10.1016/j.protis.2016.07.002.

438 A. Botero et al.

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