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This is the author’s final version of the work, as accepted for publication following peer review but without the publisher’s layout or pagination.

The definitive version is available at http://dx.doi.org/10.1016/j.protis.2015.10.001

Paparini, A., Macgregor, J., Ryan, U.M. and Irwin, P.J. (2015) First molecular characterization of Theileria ornithorhynchi

Mackerras, 1959: yet another challenge to the systematics of the piroplasms. Protist, 166 (6). pp. 609-620.

http://researchrepository.murdoch.edu.au/29099/

http://dx.doi.org/10.1016/j.protis.2015.10.001

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Please cite this article in press as: Paparini A, et al. First Molecular Characterization of Theileriaornithorhynchi Mackerras, 1959: yet Another Challenge to the Systematics of the Piroplasms. Protist (2015),http://dx.doi.org/10.1016/j.protis.2015.10.001

ARTICLE IN PRESSPROTIS 25506 1–12

Protist, Vol. xx, xxx–xxx, xx 2015http://www.elsevier.de/protisPublished online date xxx

ORIGINAL PAPER1

First Molecular Characterization ofTheileria ornithorhynchi Mackerras, 1959:yet Another Challenge to the Systematicsof the Piroplasms

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Andrea Paparinia,1, James Macgregorb, Una M. Ryana, and Peter J. IrwinaQ16

aVector- and Water-Borne Pathogen Research Group, School of Veterinary & LifeSciences, Molecular and Biomedical Sciences, Murdoch University, Murdoch WA, 6150,Australia

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bCollege of Veterinary Medicine, School of Veterinary and Life Sciences, MurdochUniversity, 90 South Street, Murdoch, Western Australia, 6150

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Submitted March 13, 2015; Accepted October 3, 201512

Monitoring Editor: Frank Seeber13

Piroplasms, tick-transmitted Apicomplexa of the genera Theileria, Babesia and Cytauxzoon, are blood-borne parasites of clinical and veterinary importance. The order Piroplasmida shows a puzzlingsystematics characterized by multiple clades, soft polytomies and paraphyletic/polyphyletic genera.In the present study, screening of platypuses (Ornithorhynchus anatinus), was performed to infer theparasite molecular phylogeny. DNA was extracted from blood, ectoparasites and tick eggs and the 18SrRNA– hsp70–genes were used for the phylogenetic reconstructions. Microscopic analyses detectedpleomorphic intra-erythrocytic organisms and tetrads consistent with previous descriptions of Thei-leria ornithorhynchi Mackerras, 1959, but observation of possible schizonts could not be confirmed.DNA sequences obtained from blood and ticks allowed resolving the systematics of the first piroplasminfecting a monotreme host. Molecularly, T. ornithorhynchi formed a novel monophyletic group, basalto most known piroplasms’ clades. The ancestral position of this clade, isolated from an ancient lineageof mammalian host appears particularly fascinating. The present paper discusses the inadequacies ofthe current molecular systematics for the Piroplasmida and the consequences of incomplete sampling,morphology-based classification and ambiguous microscopic identifications. Likely when the currentsampling bias is rectified and more sequence data is made available, the phylogenetic position ofT. ornithorhynchi will be further contextualized without ambiguity.

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© 2015 Published by Elsevier GmbH.30

Key words: Platypus; 18S rDNA; molecular phylogeny; systematics; Bayesian analysis; Piroplasmida31

1Corresponding author; fax +61 8 9360 6628e-mail [email protected] (A. Paparini).

http://dx.doi.org/10.1016/j.protis.2015.10.0011434-4610/© 2015 Published by Elsevier GmbH.

dx.doi.org/10.1016/j.protis.2015.10.001


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ARTICLE IN PRESS2 A. Paparini et al.

Introduction32

Piroplasms, which include apicomplexan organ-33

isms of the genera Theileria, Babesia and34

Cytauxzoon, are blood-borne protozoan parasites35

transmitted by ticks. Members of these genera have36

similar phenotypes and can infect mammals and37

birds (Hunfeld et al. 2008). In addition, some piro-38

plasm species are of important human pathogens39

(Gorenflot et al. 1998; Homer et al. 2000; Kjemtrup40

and Conrad 2000; Senanayake et al. 2012).41

There are currently at least seven named42

Theileria spp. infecting Australian native mam-43

mals; these include: T. ornithorhynchi in the44

platypus (Ornithorhynchus anatinus) (Mackerras45

1959), T. tachyglossi in the short-beaked echidna46

(Tachyglossus aculeatus) (Priestley 1915) and47

T. perameles in the southern brown bandicoot48

(Isoodon obesulus), long-nosed bandicoot (Per-49

ameles nasuta) and long-nosed potoroo (Potorous50

tridactylus) (Clark 2004). More recently T. gilberti51

was described in the Gilbert’s Potoroo (Potorous52

gilbertii) (Lee et al. 2009), while T. penicillata53

was found in the woylie (or brush-tailed bettong,54

Bettongia penicillata), T. brachyuri in the quokka55

(Setonix brachyurus) and T. fuliginosa in the west-56

ern grey kangaroo (Macropus fuliginosus) (Clark57

and Spencer 2007). Importantly, these previous58

studies achieved identification and genus differen-59

tiation, based on morphological evidence and/or60

molecular analyses using only partial fragments of61

the 18S rRNA gene (18S rDNA).62

Most previous reports provide little evidence that63

these piroplasms are pathogenic to native Aus-64

tralian wildlife (Clark 2004; Paparini et al. 2012;65

Portas et al. 2014; Rong et al. 2012; Vaughan et al.66

2009). However, Babesia macropus was found to67

cause severe clinical signs in Eastern Grey Kanga-68

roos (Macropus giganteus) (Dawood et al. 2013)69

and T. ornithorhynchi was recently reported to70

cause fatal haemolytic anaemia in an orphaned71

juvenile female platypus (Kessell et al. 2014).72

The platypus is a prototherian mammal and73

the only living semi-aquatic monotreme (Gust74

and Griffiths 2009; Pasitschniak-Arts and Marinelli75

1998). Platypuses live only in Eastern and Southern76

Australia (including Tasmania) and are considered77

vulnerable due to their reliance on aquatic environ-78

ments. This animal shares the order Monotremata79

with the short-beaked echidna from Australia80

and New Guinea, and the long-beaked echidna81

(genus Zaglossus) from New Guinea (Groves 2005;82

Phillips et al. 2009). The unique phylogenetic posi-83

tion of the platypus (Warren et al. 2008) makes it a84

potentially interesting model for understanding the85

evolution of mammals and of the interactions with 86

their parasites. 87

An early study reported the presence of piro- 88

plasms in erythrocytes from two platypuses from 89

Queensland, Australia, based on microscopic 90

examination of blood films and proposed the name 91

T. ornithorhynchi (Mackerras 1959). The para- 92

site was further investigated by light microscopic 93

examination of blood smears from platypuses in 94

south-eastern Australia and protozoa were found 95

in the erythrocytes of 98% of the platypuses exam- 96

ined (Collins et al. 1986). This high prevalence of 97

infection was reported to be related to the high 98

frequency of ticks infecting the platypuses (Ixodes 99

ornithorhynchi Lucas, 1845), however the para- 100

sitaemia was low (∼1.0% of erythrocytes were 101

infected) and was apparently asymptomatic. In the 102

same study, a few inclusions were also observed in 103

leukocytes and identified as possible schizonts, but 104

this was not confirmed. The intra-erythrocytic forms 105

were examined by electron microscopy and were 106

observed to contain vacuoles, micronemes, rhop- 107

tries and double unit membrane cytosomes. Collins 108

et al. reported these parasites as T. ornithorhynchi 109

(Collins et al. 1986), which was later thought to be 110

the cause of fatal haemolytic anaemia in one sick 111

individual (Kessell et al. 2014). 112

Traditional piroplasm identification and classi- 113

fication has, to a large extent, been based on 114

morphology and serology (in domestic animals), 115

both of which are of limited utility in consis- 116

tently differentiating closely related apicomplexans. 117

Babesia spp. are typically distinguished from 118

Theileria spp., based on several life-cycle char- 119

acteristics, including distinctions in their biology 120

within the invertebrate host (e.g., trans-ovarial ver- 121

sus trans-stadial transmission, in Babesia and 122

T., respectively), the mode of transmission from 123

vector to vertebrate host, and the location of 124

replication in the vertebrate hosts (i.e., Babesia 125

multiplies only in red blood cells, while T. under- 126

goes extra-erythrocytic schizogony in lymphocytes 127

or macrophages, prior to the erythrocytic stage) 128

(Uilenberg 2006). 129

Despite these relatively clear biological distinc- 130

tions, piroplasm taxonomy is still confused, with 131

the molecular systematics characterized by various 132

multi-generic clades, unresolved taxa and para- 133

phyletic or polyphyletic genera. During the present 134

study, screening of wild platypuses was performed 135

to attempt to resolve the molecular classification 136

of protozoan parasites morphologically consistent 137

with T. ornithorhynchi Mackerras, 1959. It was 138

anticipated these analyses could provide insights 139

into the molecular phylogeny of the order and 140



ARTICLE IN PRESSMolecular Systematics of the Piroplasms 3

possibly improve the currently accepted systemat-141

ics.142

Results143

Ectoparasite Identification144

The ticks (adult and nymphs) recovered from the145

7 animals were identified as Ixodes ornithorhynchi146

Lucas, 1845 (Roberts 1970). For the sample pre-147

liminarily identified as a leech, the species identity148

could not be morphologically confirmed due to149

extensive tissue damage and poor conservation.150

Molecular identification of the leech was also151

attempted as part of another study (Paparini et al.152

2014a); however, despite the quality and length of153

the sequences obtained, molecular identification154

was only obtained to family level (Glossiphoniidae).155

Piroplasm Detection by PCR156

All blood samples analysed provided positive157

amplification with at least one primer set used.158

Positive amplicons were also obtained from four159

adult ticks (PT 38, PT 104.1, PT 104.2, PT 104.3),160

the two nymphs (PTN 50, PTN 104) and the leech161

(PTL 57). Neither the eggs nor the female tick that162

laid them (PT 104.4) provided positive PCR amplifi-163

cation for piroplasms (Table 1). However, compared164

to blood, ectoparasites and eggs had a significantly165

lower yield/quality of DNA, based on spectropho-166

tometric and electrophoresis assessments (data167

not shown). This may have led to false-negative168

results for these latter groups of samples.169

Of the various primer pairs used, the set170

BTF/BTR (Jefferies et al. 2007) provided short171

sequences but with the highest quality (as172

determined by clean and specific sequence chro-173

matograms). All positive samples identified by174

this assay were successively tested with primer175

pairs BT18SF/R (Paparini et al. 2012) and generic176

apicomplexan primers CRYPTOF/CRYPTOR177

(Eberhard et al. 1999). Despite positive amplifi-178

cation, the fraction of viable (i.e., not-mixed) and179

specific sequences obtained with these latter180

primer sets was limited, because the vertebrate181

host, and sometimes the invertebrate vector,182

co-amplified with the parasite 18S rDNA. For this183

reason, cloning of a selection of positive samples184

was required.185

Of the two primer sets used to amplify the hsp70186

locus (Soares et al. 2011), only HSP70 F1/R1187

worked. However, mixed chromatograms and/or188

non-specific products were often obtained and gel-189

purified amplicons from a random selection of190

samples (PB 9, PB 52, PB 94 and PT 38) were 191

cloned to confirm parasite identity. 192

Piroplasm Morphology 193

In all positive blood films (n = 26), piroplasms were 194

observed and appeared as dark, basophilic bodies 195

surrounded by pale cytoplasm with a fine limiting 196

membrane. The organisms identified morphologi- 197

cally as T. ornithorhynchi (Collins et al. 1986) were 198

highly pleomorphic and occurred mostly singly 199

within an erythrocyte, but occasional pairs and mul- 200

tiple parasites were noted such as tetrads (Fig. 1A). 201

A low parasitaemia of approximately 0.5-1% was 202

observed in each of the blood smears examined. In 203

addition, intra-cytoplasmic inclusions within leuko- 204

cytes were very occasionally observed (Fig. 1B). 205

Despite this, in agreement with previous studies, 206

compelling identification of these rare inclusions as 207

schizonts of T. ornithorhynchi could not be ascer- 208

tained with confidence. 209

Multiple Sequence Alignment and 210

Phylogenetic Analysis (18S rDNA) 211

The phylogenetic reconstruction was carried 212

out using stringent conditions, whereby only 213

sequences of the highest quality and neatly aligning 214

with the others were used. The final sequence list 215

used for the phylogenetic reconstruction was based 216

on quality, length, specificity and position within the 217

alignment. After discarding duplicates and shorter 218

products, three unique, nearly full length 18S 219

sequences from the present study were selected for 220

the phylogenetic reconstruction: PB 94 (1,611 bp), 221

PT38 (1,602 bp) and PTN104 (1,413 bp). Each 222

sequence was obtained from a single animal 223

blood (PB) or ectoparasite (PT and PTN), by join- 224

ing multiple (overlapping) bi-directional sequencing 225

products. Including the outgroup, the features of 226

the final Gblocks-curated (de-gapped) DNA align- 227

ment were: number of taxa = 36; length = 1,154 228

positions; pairwise identity = 95.9%; parsimony- 229

informative sites 133 (11.5%); conserved sites 935 230

(81.0%). 231

In the Bayesian tree, the platypus-derived 232

sequences consistently formed a monophyletic 233

clade, which never included sequences from other 234

hosts. The clade was basal to all other piro- 235

plasms, except those from the B. microti group 236

(archaeopiroplasmids) (Fig. 2). In all trees, the 237

clade received strong statistical support: poste- 238

rior probability (PP) or bootstrap values were 0.90, 239

0.93, 0.90, 0.89 and 1.00 for MP, ML, NJ, ME and 240

BI respectively. The position of the clade within the 241




Table 1. Summary of platypus and ectoparasite samples analysed and results..Q2

Sample no. Sample ID* Source species Molecular analysis Microscopy

Sampletype

Piroplasm18S rDNA

Sampletype

Results

1 PB 52 Platypus(Ornithorhynchusanatinus)

Wholeblood

Positive Bloodsmear

Positive

2 PB 70 Positive Positive3 PB 72 Positive Positive4 PB 73 Positive Positive5 PB 74 Positive no smear6 PB 75 Positive no smear7 PB 76 Positive Positive8 PB 77 Positive Positive9 PB 78 Positive Positive10 PB 79 Positive Positive11 PB 80 Positive Positive12 PB 81 Positive Positive13 PB 82 Positive Positive14 PB 83 Positive Positive15 PB 84 Positive Positive16 PB 85 Positive Positive17 PB 87 Positive Positive18 PB 88 Positive Positive19 PB 89 Positive Positive20 PB 9 Positive Positive21 PB 90 Positive Positive22 PB 91 Positive Positive23 PB 93 Positive Positive24 PB 94 Positive Positive25 PB 95 Positive Positive26 PB 96 Positive Positive27 PB 97 Positive Positive28 PB 104 Positive Positive

Positives (subtotal) 28 26

1 PT 11 IxodesornithorhynchiLucas,1845

Adultticks

Neg. Tissue n.a.

2 PT 20 Neg. n.a.3 PT 24.1 Neg. n.a.4 PT 24.2 Neg. n.a.5 PT 34 Neg. n.a.6 PT 38 Positive n.a.7 PT 104.1 Positive n.a.8 PT 104.2 Positive n.a.9 PT 104.3 Positive n.a.10 PT 104.4 Neg. n.a.




Table 1 (Continued)

Sample no. Sample ID* Source species Molecular analysis Microscopy

Sampletype

Piroplasm18S rDNA

Sampletype

Results

Positives (subtotal) 4

1 PTN 50 IxodesornithorhynchiLucas,1845

Ticknymphs

Positive Tissue n.a.

2 PTN 104 Positive n.a.


1 PTL 57 ? Leech Positive Tissue n.a.


1 PTE 104.1 IxodesornithorhynchiLucas,1845

Tickeggs

Neg. Tissue n.a.

2 PTE 104.2 Neg. n.a.

Positives (subtotal) 0*Note: Numbers refer to the ID of the animal host, and to the specific sample (e.g., PT 104.3 correspondsto: adult tick no. 3, from platypus no. 104). Abbreviations: Neg. = Viable amplification product/sequence notobtained; n.a. = Not applicable; ? = unknown.

Figure 1. Microscopic detection of protozoan parasites morphologically consistent with previously describedTheileria ornithorhynchi Mackerras, 1959. (A): photomicrograph of a blood film from a platypus (Ornithorhynchusanatinus) showing an intra-erythrocytic piroplasm (Theileria sp.). (B): Possible intra-cytoplasmic inclusionswithin leukocytes, indicative of an extra-erythrocytic life stage (image courtesy of M. Ansell).




Figure 2. Evolutionary relationships between the known groups of piroplasms and the novel platypus-derivedclade. Phylogenetic analysis was inferred using the Bayesian Inference method, based on nearly completesequences of the 18S rRNA gene. Posterior probabilities are indicated on the main branches. GenBankaccession numbers are given.

tree was consistent for the BI, MP and ML recon-242

structions. In the NJ and ME trees only the clade243

was basal to all other piroplasms, except those244

from the B. microti and B. duncani/conradae groups245

(data not shown).246

In the BI reconstruction, no taxon remained247

unresolved and all sequences fell within mul-248

tiple monophyletic clusters. Specifically, eight249

strongly supported (PP = 1.00) main clades were250

obtained: 1) B. microti group (archaeopiroplas-251

mids); 2) T. ornithorhynchi group (proposed, this252

study); 3) B. duncani/conradae group (“western253

clade” or prototheilerids); 4) B. poelea/B. uriae254

group (avian babesias) 5) Cytauxzoon spp. group;255

6) T. equi/B. equi group; 7) Babesia sensu strictu256

group (babesids or “true babesias”); and 8) Theile-257

ria spp. group (theilerids) (Fig. 2).258

Within the monophyletic T. ornithorhynchi group,259

there appeared to be a degree of genetic variation,260

with at least two distinct genotypes, separated by261

a maximum genetic distance of about 1.8%. Based262

on our trimmed alignment and set of sequences, B.263

uriae appeared the sequence most closely related264

to PT 38 and PTN 104, with a genetic distance of265

2.5% (B. uriae has been submitted to GenBank as 266

Babesia sp. MJY-2009a) (Yabsley et al. 2009). T. 267

sinensis from a bovine (Acc. No. KF559355) (Tian 268

et al. 2013) was 2.9% distant from PB 94. 269

Phylogenetic Analysis (hsp70) 270

After editing and trimming, two identical high-quality 271

hsp70 sequences from the present study were 272

included (PB 09 and PT38, from a blood and tick 273

sample, respectively). The final, de-gapped align- 274

ment included 23 partial sequences, 581 positions, 275

80.0% pairwise identity, 238 (41.0%) parsimony 276

informative sites, 286 (49.2%) conserved sites. In 277

the Bayesian tree, PB 09 and PT 38 clustered out- 278

side the Babesia sensu strictu group, and within a 279

paraphyletic clade of Theileria spp. (Fig. 3). 280

Based on genetic distance, the platypus-derived 281

genotype was marginally more similar to some iso- 282

lates of Babesia macropus (e.g., accession number 283

KM389895) than it was to the theilerias. How- 284

ever, the unrealistic genetic distances (44% - 48%), 285

biased by the scant reference set of taxa available 286

in GenBank, make this result likely meaningless. 287




Figure 3. Evolutionary relationships between known reference piroplasms and two identical platypus-derivedisolates. Phylogenetic analysis was inferred using the Bayesian Inference method, based on partial hsp70sequences. Posterior probabilities are indicated on the main branches. GenBank accession numbers are given.

Discussion288

A thorough review of earlier microscopic findings289

(Collins et al. 1986; Kessell et al. 2014; Mackerras290

1959) reveals that T. ornithorhynchi has been291

assigned to the genus Theileria based only on292

morphological observations of blood films from293

platypuses. Josephine Mackerras first described294

“minute, rounded, oval, or elongate parasites with295

(the) nucleus usually near one end” and that296

“single parasites predominated but some cells297

contained two, three or four forms” (Mackerras298

1959). The same author also noted that “these299

parasites” were also found by Duncan (in 1950)300

in Tasmania (Mackerras 1959), and subsequently301

named it T. ornithorhynchi (Mackerras 1959). More302

recently a case of fatal anaemia secondary to303

T. ornithorhynchi infection was reported (Kessell304

et al. 2014). Mackerras (Mackerras 1959) failed305

to find schizonts in blood films or tissues, while306

Kessell et al. (Kessell et al. 2014) provided lim-307

ited microscopic evidence of the parasite and308

inconclusive molecular data. Organisms presumed309

to be T. ornithorhynchi were also described by310

Collins et al. in Australian platypuses (Collins et al.311

1986). Interestingly, in agreement with the present312

investigation (Fig. 1B), this is the only study describ- 313

ing the sporadic presence of tetrads and putative 314

extra-erythrocytic (schizont) stages, which are the 315

distinguishing features of the genus Theileria. Apart 316

that the leukocytic inclusions might not necessar- 317

ily be part of the life cycle of the intraerythrocytic 318

forms (Collins et al. 1986), the compelling evidence 319

of schizonts (of T. ornithorhynchi) has never been 320

ascertained in any study. Thus, providing that all 321

organisms from the previous papers are the same, 322

it appears that the documentation of tetrads is the 323

only conclusive evidence that this parasite belongs 324

to the genus Theileria. Transovarial transmission 325

is a peculiarity of Babesia spp., used to distin- 326

guish them from members of the genus Theileria. 327

The absence of such a modality of transmission 328

would have further confirmed the similarity of T. 329

ornithorhynchi to the true theilerias. Unfortunately 330

despite the negative PCR results provided by the 331

tick eggs tested during the present study, the female 332

tick that laid the eggs (PT 104.4) was also negative 333

(or the extracted DNA was unviable) (Table 1). 334

The present paper represents the first molecu- 335

lar characterization of piroplasms morphologically 336

compatible with T. ornithorhynchi (Collins et al. 337

1986; Kessell et al. 2014; Mackerras 1959). 338




Clearly the true relationship between the parasites339

observed in the present study and those described340

earlier remains unknown. The paucity of known341

taxa available for comparison and the length of the342

product suggest caution in interpreting the results343

of the hsp70-based reconstruction. For instance,344

only distantly-related reference sequences from345

Babesia spp. and Theileria spp. could be retrieved346

from GenBank (99% query coverage, ≤ 80%347

sequence identity). However if the morphology-348

based assignment of T. ornithorhynchi to the genus349

Theileria is correct, the molecular position of this350

taxon, outside the clade of the theilerids is particu-351

larly fascinating.352

The current molecular systematics of the piro-353

plasms is an artificial classification representing354

the best approximation available up to now, to355

the true (unobservable) phylogeny of the order.356

It is based on available data which constantly357

expands and refine, allowing identification and clas-358

sification of novel isolates. The consequences of359

erroneous microscopic identifications can have pro-360

found repercussions on the molecular systematics,361

for instance when DNA sequences are obtained362

from misidentified isolates. On the other hand,363

molecular analyses and phylogenetic reconstruc-364

tions may also overestimate diversity and be prone365

to pitfalls (Berney et al. 2004).366

Interestingly, the genetic variation observed367

within the T. ornithorhynchi group (1.8% maxi-368

mum intra-group genetic distance) was significant,369

when compared to pairwise distances calculated370

between other valid named species. For exam-371

ple, the “true babesia” B. crassa (GenBank Acc.372

No. AY260176) was 1.7% distant from B. bigem-373

ina (EF458206); in the archaeopiroplasmids, B.374

duncani (HQ289870) was 0.4% distant from B.375

conradae (AF158702), while between T. ovis376

(AY260172) and T. buffeli (HQ840964) the dis-377

tance was only 0.1%. (Fig. 2). Clear-cut criteria378

have previously been adopted to define the genetic379

distances required for a piroplasm to be classi-380

fied as a distinct species (Schnittger et al. 2003).381

At the 18S rRNA locus, a genetic distance of382

distance ≥2.1% for Babesia and ≥0.7% for T.383

is required (Schnittger et al. 2003). Unfortunately384

these figures have relatively little use because385

the parameters, methods and the alignment used386

for calculating the pairwise distance values can387

all affect the values. Although a fraction of the388

observed intra-group genetic variation may be due389

heterozygosity and/or minor sequencing artefacts,390

this raises the question whether the platypus-391

derived sequences actually consisted of multiple392

genotypes or species.393

It is generally recognized that the 18S-based 394

molecular systematics of the order Piroplasmida 395

shows inadequacies, especially at deeper levels 396

(Schnittger et al. 2012). Among the many reasons 397

that can account for this impasse is that, so far, sam- 398

pling of isolates has been done in most cases with 399

a focus on medical (human babesiosis) or veteri- 400

nary (especially bovine, equine, ovine and canine 401

piroplasmosis) research questions. The piroplasm 402

sequences obtained during the present study are in 403

fact the first obtained from monotremes. Since the 404

topology of the 18S tree produced is remarkably 405

strong it sheds a new light on the deep phylogeny 406

of the order Piroplasmida, by revealing a novel 407

monophyletic clade of parasites isolated from an 408

ancient lineage of mammalian hosts. Nevertheless, 409

until more sequences are collected, the true evolu- 410

tionary relationships and particularly the ancestral 411

position of T. ornithorhynchi clade cannot be con- 412

firmed. 413

A recent analysis based on the cytochrome oxi- 414

dase I gene (Gou et al. 2013), suggests that the 415

divergence time for the piroplasms was during the 416

Paleocene, 56.48 million years ago (MYA) (95% 417

highest posterior density [HPD] 28.17–86.87). 418

Theileria spp. showed a node age of 23.38 MYA 419

(95% HPD 11.11–36.71) and 25.74 MYA (95% HPD 420

12.75–40.73) for Babesia. The phylogenetic recon- 421

struction presented in the study of Gou et al. (2013), 422

was based on species of “true theilerias” and “true 423

babesias”, but it did not include members of the B. 424

microti group. Thus, based on these estimations, 425

the appearance of the most ancestral piroplasms 426

(including B. microti and possibly T. ornithorhynchi) 427

may be placed sometime between 56.48 and 25.74 428

MYA (range 86.87-12.75 MYA). This time range 429

overlaps with the emergence of the hard-tick genus 430

Ixodes (vector of the archeopiroplasmids and T. 431

ornithorhynchi), which is believed to have formed 432

46.71 MYA (95% HPD 39.95–53.45), but it clearly 433

pre-dates the divergence of the platypus from the 434

echidna (19–48 MYA) (Phillips et al. 2009). This 435

suggests that the latter monotremal host may also 436

harbour piroplasm genotypes closely related to 437

T. ornithorhynchi (e.g., T. tachyglossi) and future 438

molecular studies of this species are warranted. 439

A number of piroplasm sequences from native 440

Australian marsupials are currently available in 441

GenBank (cf. (Dawood et al. 2013; Mans et al. 442

2015; Paparini et al. 2012)). These sequences have 443

not been included in either recent seminal recon- 444

structions (Lack et al. 2012; Schnittger et al. 2012) 445

or in the present analysis (due to their length). 446

Future studies dealing with the systematics of the 447

order should not overlook these hosts because 448




ancient sister piroplasmid lineages are likely to be449

encountered in Australian mammals, as previously450

suggested (Schnittger et al. 2012) and further sup-451

ported by the present study.452

Conclusions453

In conclusion, the present study represents the454

first comprehensive molecular characterization of455

protozoan parasites morphologically consistent456

with the previously described T. ornithorhynchi457

Mackerras, 1959, from the platypus.458

Molecular classification of T. ornithorhynchi was459

resolved by sequencing nearly-complete fragments460

of the 18S rDNA and the phylogeny obtained461

revealed a new monophyletic group, basal to most462

piroplasm clades but internal to the archeopiro-463

plasmids. The inclusion of the first piroplasm full464

length 18S sequence from a monotremal host in a465

novel monophyletic clade poses yet another chal-466

lenge to the molecular systematics of piroplasms.467

The position of a putative Theileria sp. outside the468

clades including the equine theilerias (i.e., T. equi)469

or the theilerids is fascinating, but also urges for a470

thorough revision of the molecular systematics of471

the order and a broader sampling effort. The gen-472

eral agreement that the B. microti group should be473

renamed and form a new genus (Cornillot et al.474

2012; Goethert and Telford 2003; Zahler et al. 2000)475

requires further consideration in light of the new476

phylogenetic reconstruction presented here.477

Methods478

Isolates/Animal sources: Samples were obtained from platy-479

puses captured in fyke nets, in the Inglis river catchment, in480

north-west Tasmania, Australia (40◦58’55.7“S 145◦43’35.2”E)481

as previously described (Whittington and Grant 1995). The482

study was approved by the Animal Ethics Committee of483

Murdoch University, Western Australia (Permit Number RW484

2422/11), Department of Primary Industries, Parks, Water and485

Environment (DPIPWE), Tasmania (Permit to take Wildlife for486

Scientific Purposes Numbers FA 11131 and FA 12165) and the487

Inland Fisheries Service, Tasmania (Exemption Permit Number488

2011-10). All animals were released after examination and the489

procedures described below.490

Blood samples and ectoparasites: Venous blood sam-491

ples (PB samples; n = 28) were collected from the bill sinus492

of anaesthetised platypuses captured between September493

and December 2012. The blood was collected into potassium494

EDTA-treated microtubes (Sarstedt, Germany), refrigerated at495

4◦C, until haematological analysis and subsequent dispatch to496

Murdoch University, where it was stored frozen at -20◦C, until497

processed as previously described (Paparini et al. 2014a)498

(Table 1). Whilst anaesthetised, the skin and hair coat of each499

platypus was inspected for ectoparasites and any that were500

observed were removed, stored in 70% ethanol and sent to 501

Murdoch University for identification and molecular analyses. 502

Ectoparasites were removed from 8 animals in total (platypuses 503

11, 20, 24, 34, 38, 50, 57 and 104) and included: adult ticks (PT 504

samples; n = 10), tick nymphs (PTN samples; n = 2) and one 505

leech (PTL 57) (Table 1). Moreover, one engorged adult tick 506

was kept alive in a humidified tube until it produced eggs. Initially 507

the eggs were incubated in humidified tubes at room tempera- 508

ture to encourage hatching, but when none had hatched after 509

3-4 weeks, these also were stored in 70% ethanol for molec- 510

ular analyses (PTE samples; n > 500 in two batches; from 511

an engorged tick, collected from one animal). The respective 512

female tick was preserved in 70% ethanol once egg-laying was 513

complete and then analysed. Ectoparasites were morpholog- 514

ically identified using a stereo microscope (Olympus SZ61 I, 515

Japan) and reference to a standard key (Roberts 1970), prior 516

to DNA extraction and molecular analysis. 517

Blood film analysis: A single drop of peripheral blood 518

was used to make thin blood smears, which were stained on 519

arrival at Murdoch University with a modified Wright’s stain 520

using an Ames Hema-Tek slide stainer (Bayer, Germany). 521

Stained blood films were systematically examined at x400 mag- 522

nification for the presence of trypomastigotes using a BX50 523

microscope (Olympus, Japan) with screen views generated 524

by a DP Controller v3.2.1.276 (Olympus, Japan). If parasites 525

were observed, the morphology was noted and measurements 526

were made at ×1000 magnification, using Image J software 527

(http://rsbweb.nih.gov/ij/). Additional assessment of peripheral 528

blood cell morphology and characteristics (e.g., anisocytosis, 529

polychromasia, estimate of platelet numbers and the presence 530

of intracellular inclusions; data not shown) was also conducted 531

at Murdoch University using the blood smears described previ- 532

ously. 533

DNA extraction: Ectoparasites were washed twice in ster- 534

ile, molecular grade water and sliced on sterile Petri dishes, 535

using sterile scalpel blades. Total genomic DNA was isolated 536

using the QIAamp DNeasy Animal Tissue Spin-Column Proto- 537

col (Qiagen, USA). Lysis was achieved by overnight digestion 538

with Proteinase K (Qiagen, USA), followed by bead beating 539

on a benchtop Vortex Genie 2 vortex (5 minutes, maximum 540

speed) (MO BIO, USA). DNA was eluted in 60 �L of TE buffer. 541

PTN samples (PTN 50 and PTN 104) consisted of two pools 542

of nymphs (5 nymphs/pool). Each pool consisted of ectopara- 543

sites collected from one animal only (i.e., from platypus 50 or 544

104, respectively) (Table 1). DNA from tick eggs was extracted 545

in the same way, except that no scalpel blade was used. For 546

whole blood/EDTA (200 �L), total genomic DNA was isolated, 547

according to the manufacturer’s instructions, using a Master- 548

Pure Purification Kit (Epicentre Biotechnologies, USA) and 549

resuspended in 50 �L of TE buffer. Mock extractions were car- 550

ried out from sterile molecular-grade water, to exclude DNA 551

contamination from reagents and consumables. All DNA prepa- 552

rations were checked for purity by agarose gel electrophoresis 553

and quantified by spectrophotometric absorbance using a Nan- 554

odrop ND-1000 (Thermo Scientific, USA). 555

Molecular analysis: DNA samples were screened for the 556

presence of piroplasmid-specific 18S ribosomal RNA gene 557

(18S rDNA), by nested PCR (850 bp), using BTF1/BTR1 558

and BTF2/BTR2 primers as previously described (Jefferies 559

et al. 2007). For confirmation, a second nested PCR 560

primer set (Paparini et al. 2014b) was used to obtain 561

longer 18S rDNA sequences (1,466 bp), from a subset 562

of positive samples; primer pairs BT18SF1/BT18SR1, fol- 563

lowed by BT18SF2/BT18SR2, were used as previously 564

described (Paparini et al. 2012). Additional generic api- 565

complexan 18S primers, used in ancillary assays, included 566


http://rsbweb.nih.gov/ij/



CRYPTOF/CRYPTOR (Eberhard et al. 1999). For hsp70 ampli-567

fications, primers HSP70 F1/R1 and HSP70 F2/R2 were used568

(Soares et al. 2011).569

All amplifications performed included negative and posi-570

tive controls consisting, respectively, of sterile molecular-grade571

water and genomic DNA preparations from piroplasm-infected572

animals identified during previous analyses. PCR products573

were checked by electrophoresis with SYBR Safe Gel Stain574

(Invitrogen, USA) and visualised with a dark reader trans-575

illuminator (Clare Chemical Research, USA). PCR amplicons576

corresponding to the expected length were excised from the577

gel and sequenced, bi-directionally, using an ABI Prism Termi-578

nator Cycle Sequencing kit (Applied Biosystems, USA), on an579

Applied Biosystem 3730 DNA Analyzer.580

Cloning: Gel-purified PCR products providing mixed or581

low-quality DNA sequencing chromatograms (18S rDNA and582

hsp70) were cloned in the pGEM-T Easy Vector System II583

(Promega, USA). For cloning, products from samples PB 9,584

PB 52, PB 94 and PT 38 were chosen. After transformation of585

competent cells, plasmid DNA was extracted using the QIAprep586

Spin Miniprep Kit (Qiagen, Germany) from a subset of transfor-587

mants and sequenced using the appropriate amplicon-specific588

primers, as described above.589

Phylogenetic analysis: For 18S, Sanger sequencing chro-590

matogram files were imported in Geneious Pro 8.1.6 (Kearse591

et al. 2012), and low quality regions were trimmed using a592

threshold value of 0.005: this operation trims all regions with593

more than a 0.5% chance of an error per base. Multiple594

Sanger sequencing chromatograms obtained by bi-directional595

sequencing from a single DNA source (i.e., from only one ani-596

mal or ectoparasite) were assembled. For this operation the597

following settings were adopted: custom sensitivity; minimum598

overlap length = 100; minimum overlap identity = 100%.599

Inclusion in the final subset used for the phylogenetic recon-600

struction was based on quality, length, specificity and position601

of the sequence within the alignment. Sequences selected for602

the phylogenetic reconstruction were submitted to GenBank603

(Acc. No.). The alignment was curated by Gblocks (Castresana604

2000), remotely (Dereeper et al. 2008) with the high-stringency605

set of options selected; the final alignment contained no gaps.606

Nucleotide substitution models were tested in MEGA6607

(Tamura et al. 2013). The Kimura 2-parameter + G (0.6) + I (0.6)608

model was chosen for the Bayesian Inference (BI) analysis609

which was carried out by MrBayes (Ronquist and Huelsenbeck610

2003), using the default options. Trees were visualized by611

FigTree v1.4.0 (http://tree.bio.ed.ac.uk/). Genetic distances612

were also computed in MEGA6 (Tamura et al. 2013), using the613

Kimura 2-parameter model. The rate variation among sites was614

modelled with a gamma distribution (shape parameter = 0.6). All615

positions containing gaps and missing data were eliminated.616

The evolutionary history was also inferred by the maximum617

likelihood (ML)–, neighbour-joining (NJ)–, maximum parsimony618

(MP)– and minimum evolution (ME)– phylogenies in MEGA6619

(Tamura et al. 2013) (bootstrap replicates = 500).620

For BI analysis based on the hsp70 locus the same pro-621

cedure was used. However alignment was not curated as it622

contained no gaps, and the Kimura 2-parameter + G (0.3) model623

was chosen, after testing in MEGA6 (Tamura et al. 2013).624

Acknowledgements625

The authors wish to thank Prof Michael J Wise626

(University of Western Australia) for constructive627

discussions, criticism and for independently testing628

our results using a variety of innovative bioinfor- 629

matics approaches. The authors also wish to thank 630

Dr Graeme Knowles (DPIPWE Tasmania) and 631

acknowledge specific assistance by Mel Ansell 632

(The Animal Health Laboratory), for conducting 633

the haematological and biochemical testing, and 634

Aileen Elliott and Louise Pallant (Murdoch Uni- 635

versity), for their assistance in the identification 636

of some of the ticks. Authors’ gratitude also goes 637

to Prof Matthew Bellgard (Murdoch University) for 638

useful suggestions; to Dr David Berryman, Ms 639

Frances Brigg, Dr John Fosu-Nyarko, Ms Elvina 640

Lee, Dr Steve Wylie (Murdoch University) and 641

A/Prof Christopher Peacock (University of Western 642

Australia) for technical support. 643

Financial and/or in-kind support for this project 644

was provided by: the Winifred Violet Scott Estate, 645

a Caring For Our Country Community Action 646

Grant, the Central North Field Naturalists, the 647

National Geographic Society, the Cradle Coast Nat- 648

ural Resource Management, Tasmanian Alkaloids, 649

the DPIPWE Tasmania, the Australian Geographic 650

Society, the Forestry Practices Authority, and the 651

Edward Alexander Weston and Iris Evelyn Fer- 652

nie Research Fund. The sponsors provided logistic 653

support and field-assistance during sampling, but 654

had no involvement in either the writing of the report 655

or the decision to submit the article for publication. 656

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