Mem Inst Oswaldo Cruz, Rio de Janeiro, 113(1) January 2018
Original Article

Trypanosoma janseni n. sp. (Trypanosomatida: Trypanosomatidae) isolated from Didelphis aurita (Mammalia: Didelphidae) in the Atlantic Rainforest of Rio de Janeiro, Brazil: integrative taxonomy and phylogeography within the Trypanosoma cruzi clade

Camila Madeira Tavares Lopes1, Rubem Figueiredo Sadok Menna-Barreto2, Márcio Galvão Pavan3, Mirian Cláudia de Souza Pereira4, André Luiz R Roque1+

1Fundação Oswaldo Cruz-Fiocruz, Instituto Oswaldo Cruz, Laboratório de Biologia de Tripanosomatídeos, Rio de Janeiro, RJ, Brasil
2Fundação Oswaldo Cruz-Fiocruz, Instituto Oswaldo Cruz, Laboratório de Biologia Celular, Rio de Janeiro, RJ, Brasil
3Fundação Oswaldo Cruz-Fiocruz, Instituto Oswaldo Cruz, Laboratório de Mosquitos Transmissores de Hematozoários, Rio de Janeiro, RJ, Brasil
4Fundação Oswaldo Cruz-Fiocruz, Instituto Oswaldo Cruz, Laboratório de Ultraestrutura Celular, Rio de Janeiro, RJ, Brasil

Page: 45-55 DOI: 10.1590/0074-02760170297
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ABSTRACT

BACKGROUND Didelphis spp. are a South American marsupial species that are among the most ancient hosts for the Trypanosoma spp.

OBJECTIVES We characterise a new species (Trypanosoma janseni n. sp.) isolated from the spleen and liver tissues of Didelphis aurita in the Atlantic Rainforest of Rio de Janeiro, Brazil.

METHODS The parasites were isolated and a growth curve was performed in NNN and Schneideru2019s media containing 10% foetal bovine serum. Parasite morphology was evaluated via light microscopy on Giemsa-stained culture smears, as well as scanning and transmission electron microscopy. Molecular taxonomy was based on a partial region (737-bp) of the small subunit (18S) ribosomal RNA gene and 708 bp of the nuclear marker, glycosomal glyceraldehyde-3-phosphate dehydrogenase (gGAPDH) genes. Maximum likelihood and Bayesian inference methods were used to perform a species coalescent analysis and to generate individual and concatenated gene trees. Divergence times among species that belong to the T. cruzi clade were also inferred.

FINDINGS In vitro growth curves demonstrated a very short log phase, achieving a maximum growth rate at day 3 followed by a sharp decline. Only epimastigote forms were observed under light and scanning microscopy. Transmission electron microscopy analysis showed structures typical to Trypanosoma spp., except one structure that presented as single-membraned, usually grouped in stacks of three or four. Phylogeography analyses confirmed the distinct species status of T. janseni n. sp. within the T. cruzi clade. Trypanosoma janseni n. sp. clusters with T. wauwau in a well-supported clade, which is exclusive and monophyletic. The separation of the South American T. wauwau + T. janseni coincides with the separation of the Southern Super Continent.

CONCLUSIONS This clade is a sister group of the trypanosomes found in Australian marsupials and its discovery sheds light on the initial diversification process based on what we currently know about the T. cruzi clade.

Trypanosomes areobligate protozoan parasites that infect all vertebrate classes worldwide. Theirlife-cycle usually alternates between vertebrate hosts and a variety of sanguinivorousinvertebrate hosts that act as vectors. There are a variety of Trypanosomaspecies, but only some of them represent a severe public health and economicchallenge. These include Trypanosoma cruzi, which is responsible forChagas disease in South America and other parts of the world, and T. brucei,responsible for human and animal African trypanosomiasis.

Trypanosomaspecies from fish and anurans are grouped into the aquatic clade, while thespecies from the terrestrial clade is divided into two non-taxonomic groups,Salivaria and Stercoraria, according to the parasites' respective mode of transmission.Species from the Salivaria group are cyclically transmitted via salivary inoculationby tsetse flies (Glossina spp., Diptera) in Africa, while some are mechanicallytransmitted by other vectors on other continents, such as the Tabanus spp.and Stomoxys spp. The Salivaria group comprises five subgenera: Tejeraia,Trypanozoon, Nannomonas, Duttonella and Pycnomonas.Species from the Stercoraria group are transmitted via a contamination pathway,whereby parasites are eliminated in the faecal contents of different sanguinivorousinsects (triatomines and others). This group comprises three subgenera: Megatrypanum,Herpetosoma and Schizotrypanum (Lima et al. 2015 17 ).

It is proposedthat the genus Trypanosoma contains a monophyletic group of organismsthat evolved from a common African ancestor. Two main hypotheses were proposedfor the origin and dispersion of the species in the T. cruzi clade. Thefirst, the vicariant southern supercontinent hypothesis, is based on the continentaldrift as a source of separation between the ancestral Trypanosoma populations,which may have caused population divergence and speciation (Hamilton et al.2012). In this sense, the separation of Africa and the Americas would correspondwith the separation and later diversification of the species belonging to thetwo groups of Trypanosoma. The few species of the T. cruzi cladepresent in the Old World would have been unique to bats and therefore, dispersedas a result of their host migrations. However, Hamilton et al. (2009) demonstratedinfections in African terrestrial mammals by Trypanosoma species fromthe T. cruzi clade. This finding, associated with the fact that the vastmajority of the species within this clade is exclusively described in bats andbased on the phylogenetic inferences of the known Trypanosoma spp. inthis clade, presented "The Bat Seeding Hypothesis" that proposed bats as theancestral hosts of T. cruzi and T. rangeli (Hamilton et al. 2012 12 ).

Although not currentlyaccepted as an explanation for the origin of the Trypanosoma genus, thesouthern supercontinent hypothesis is accepted to explain the origin of theTrypanosoma infection in marsupials, some of the most ancient hosts ofthese parasites. Actually, the ancestors of the currently known marsupial speciesoriginated in the portion of the supercontinent that would eventually becomethe Americas and probably dispersed to the areas that now comprise Antarcticaand Oceania. It is expected that the parasites infecting these ancestral specieslikely dispersed with their hosts. Opossum species of the genus Didelphis(Didelphidae) would have originated after continental separation and are contemporaryrepresentatives of South America's native marsupial fauna (Mitchell et al. 2014 20 ).On this continent, and also in Central and North America, these opossums havea wide distribution, occupying distinct niches in natural and disturbed environments.Partially because of their synanthropic characteristics, they are some of themost studied hosts with regard to infections with species of Trypanosomaand Leishmania.

In a pioneeringstudy, Noyes et al. (1999) found a putatively new trypanosome species (H25)that infected the eastern grey kangaroo (Macropus giganteus) in Australia,which was recently described as T. noyesi (Botero et al. 2016 6 ). Sincethen, molecular tools have been widely employed to describe new Trypanosomaspecies, using blood as the main source of DNA. Some new species have been describedin recent years, especially in Australia, but the majority have been identifiedbased only on their molecular characteristics, without parasite isolation. Thisleaves some important biological aspects of these parasite species unknown.Only in a few instances new species of Trypanosoma have been isolatedand morphologically analysed (Noyes et al. 1999 22 , McInnes et al. 2011 19 , Lima etal. 2013, 2015).

In this study,we used an integrative approach to assess a new trypanosome species that wasobtained from the spleen and liver tissues of a Didelphis aurita capturedat Campus Fiocruz Mata Atlântica (CFMA), Rio de Janeiro, Brazil. Thisincluded the isolation of the parasite, as well as its characterisation andDNA sequencing. Through phylogenetic analyses, Trypanosoma janseni n.sp. provides new data on the evolutionary history of the origin of the T.cruzi clade.

 

MATERIALS ANDMETHODS

Parasites- Two populations of flagellated parasites (17665B and 17665F) were isolatedfrom the spleen and liver fragments of D. aurita (common opossum), duringa field expedition conducted at Campus Fiocruz of Mata Atlântica (Riode Janeiro, Brazil) in July 2012. This specimen was an adult male (with completedentition and 1.4 kg of body weight), captured using a Tomahawk® live-trap(Tomahawk Live Traps, Tomahawk, WI, USA) in a transition area between peridomiciliaryand forest areas (22º56'27.89''S; 43º24'23.17''W). Both parasite populationswere isolated using biphasic media composed of NNN (Neal, Novy, Nicolle) andSchneider's media with 10% foetal bovine serum (FBS), maintained in a BOD chamberat 28ºC.

Culture mediumselection and parasite growth curve - Five distinct types of culture mediawere supplemented with 10% FBS and tested to establish in vitro culturesof the parasites: DMEM (Dulbecco's Modified Eagle's Medium), EAGLE (minimumEagle Medium), TRPMI (Roswell Park Memorial Institute, commercial cell culturemedium supplemented with tryptose), RPMI 1640 (Roswell Park Memorial Institute,commercial cell culture medium), and LIT (Liver Infusion Tryptose) (Noyes etal. 1999). We also tested the feeder-layer technique (employed for distinct,non-cultivable trypanosome species), in which monolayers of Vero cells and RPMI1640 media were used with 10% FBS (v/v).

A growth assaywas performed in triplicate, in two distinct biological replicates, adaptingthe methodology described in Araújo et al. (2007) using Schneider's mediumcomplemented with 10% (v/v) of FBS and 2% (v/v) human male urine. The parasiteswere grown in test tubes containing 5 mL of the aforementioned culture mediainoculated with 1 × 106 parasites. The parasites were incubatedat 27ºC and counted daily using a Neubauer haemocytometer, until the dayafter the doubling time. Culture smears at the exponential phase of in vitrogrowth (day 2) were made in triplicate, Giemsa-stained, and observed undera Zeiss AxioObserver M1 microscope (Oberkochen, Germany).

Ultrastructureanalysis - For transmission electron microscopy (TEM) analyses, isolates17665B and 17665F were grown to exponential phase (5 × 107cells), washed thrice with PBS, and fixed in 2.5% glutaraldehyde (GA) in 0.1M sodium cacodylate buffer (pH 7.2) for 40 min at 25ºC. Post-fixation,cells were treated with 1% osmium tetroxide (OsO4) in 0.1 M sodiumcacodylate buffer (pH 7.2) containing 0.8% potassium ferricyanide and 2.5 mMcalcium chloride for 20 min at 25ºC. Samples were then dehydrated througha graded acetone series (50%, 70%, 90%, and twice at 100%), and embedded inPoly/Bed 812 resin. Ultrathin sections were contrasted with 5% uranyl acetate(20 min) and lead citrate (2 min) prior to examination with a JEOL JEM-1011transmission electron microscope (Tokyo, Japan). Alternatively, for scanningelectron microscopy (SEM) analysis, fixed parasites were adhered to poly-L-lysine-coatedcoverslips, post-fixed with 1% OsO4, and dehydrated through a gradedethanol series (50%, 70%, 90%, and twice at 100%).

The samples weredried using a critical point method with CO2, mounted on aluminiumstubs, coated with a 20 nm-thick gold layer, and examined with a JEOL JSM-6390LVscanning electron microscope (Tokyo, Japan). Ultrastructure analyses were performedat Plataforma de Microscopia Eletrônica, IOC, FIOCRUZ. Morphometric datawere taken from 50 SEM images obtained from each isolate using the ImageJ program v. 1.47. The measured parameters were the size of the body andflagellum and the total size of the parasite.

Nested polymerasechain reaction (PCR) targeting 18S SSU, PCR targeting glycosomal glyceraldehyde-3-phosphatedehydrogenase (gGAPDH), and DNA sequencing of both PCR products - NestedPCR targeting a portion of the variable region of the small subunit ribosomalgene (18S SSU) was performed in two rounds. For the first round, each 50 µLreaction contained 2 µL of DNA (50 ng/µL), 10 µL of 5x polymerisationbuffer with 100 mM dNTPs, 3 mM MgCl2, 1.4 U of Taq DNA polymerase(Promega, Madison, USA), and 16 pmol of the following external primers:TRY927F (5'-GAAACAAGAAACACGGGAG-3') and TRY927R (5'-CTACTGGGCAGCTTGGA-3'). Thermalcycling was conducted in an Esco Swift MaxPro Thermal Cycler SWT-MXP+SWY-MXP-BL-Cfor 30 cycles at 94ºC for 30 s, 55ºC for 60 s, and 72ºC for 90s. Products from the first amplification were diluted (1:10) in sterile deionisedwater. For the second round of PCR, 2 µL of this dilution was used asa template with the following internal primers: SSU561F (5'-TGGGATAACAAAGGAGCA-3')and SSU561R (5'-CTGAGACTGTAACCTCAAAGC-3'), using the same PCR reaction mixtureand cycle conditions described above (Smith et al. 2008 26 ). Amplified productswere separated by molecular weight on a 1.5% agarose gel run at 100 V for 1h in Tris-acetate EDTA buffer, stained with ethidium bromide, and visualisedby illumination with UV light. Samples producing a band of approximately 600bp were considered positive. For the gGAPDH target, we performed amplificationon an 800 bp portion of the gene as previously described (Borghesan et al. 2013 5 ).Reaction mixtures (50 µL) contained ∼100ng of DNA, 100 ng of each primer, 200 M of each dNTP, 1.5 mM MgCl2,and 2.5 U of Taq DNA polymerase. Reactions were run for 30 cycles of1 min at 94ºC, 2 min at 48ºC and 2 min at 72ºC, with an initialcycle of 3 min at 94ºC and a final cycle of 10 min at 72ºC in theaforementioned thermocycler.

Amplified productswere separated by molecular weight on a 1.5% agarose gel at 100 V for 1 h inTris-acetate EDTA buffer and stained with ethidium bromide. Bands were visualisedon an automated gel imaging system, Gel Logic 212 Pro, using the CarestreamMI SE image software (Carestream, Rochester, NY, USA). Samples with single,clear bands at approximately 600 bp and 800 bp for 18S SSU and gGAPDH, respectively,were considered positive.

Both moleculartargets were purified using the Wizard® SV Gel and PCR Clean-UpSystem (Promega, Madison, USA) following the manufacturer's instructions. Bothstrands were subjected to Sanger sequencing reactions (ABI PRISM®BigDye® Terminator v.3.1 Cycle Sequencing Kit, Applied Biosystems)and run on an ABI 3730 sequencer (Applied Biosystems®, California,USA) at the DNA Sequencing Platform PDTIS/ Fiocruz.

Phylogeneticand distance analyses of SSU rRNA and gGAPDH genes - For each gene, we visuallyinspected the forward and reverse DNA strands and generated consensus sequencesusing SeqMan Lasergene v.7.0 (DNASTAR Inc., Madison, Wisconsin, USA). Thesesequences were compared against the NCBI database (https://blast.ncbi.nlm.nih.gov)with the BLASTn algorithm, using values > 94% and > 96% as the identityand coverage cut-offs, respectively. We aligned our sequences with the sequencesretrieved from the GenBank database in MAFFT v.7.0, using the L-INS-i algorithm(Katoh & Standley 2013). The 18S SSU alignment was visually inspected andmanually edited to improve accuracy, especially in highly variable regions.Due to the high variability and low-confidence homology of a 165 bp fragmentfrom the total 737 bp 18S SSU alignment, this region was excluded from the analysis(Supplementary data, Fig.1).

We assessed thelevel of base substitution saturation in gGAPDH and 18S SSU sequences by calculatingthe entropy-based index implemented in DAMBE v.6. In brief, an index of substitutionsaturation (ISS) is calculated using the alignmentdata and compared to a critical index of substitution saturation (ISS.C),which determines a threshold for significant saturation of the data. If ISSis not significantly lower than ISS.C, sequences haveexperienced severe substitution saturation.

Possible recombinantregions in gGAPDH and 18S SSU sequences were identified with the RecombinationAnalysis Tool (Etherington et al. 2005 9 ) that uses a distance-based method ofrecombination detection, in which crossover points can be visually inspectedin the distance plots. We used different species that belong to the T. cruziclade as query for comparisons. All crossover points were statistically testedwith the Genetic Algorithm for Recombination detection (Pond et al. 2006 25 ) throughthe assessment of the goodness of fit with the Akaike information criterion(AIC), derived from a maximum likelihood model fit to each segment.

Akaike and Bayesianinformation criteria of the jModelTest v.2 were used to choose Tamura-Nei (Tamura& Nei 1993) with four gamma categories (TrN+Γ) with unequal nucleotide frequencies and TrN+Γ with equal frequencies of nucleotides as the best-fit evolutionary models forthe gGAPDH and 18S SSU datasets, respectively.

We tested whethera strict or a relaxed molecular clock best fit our data through a Bayesian randomlocal clock analysis (RLC) (Drummond & Suchard 2010) with BEAST v.1.8. Threeindependent runs were performed for 2 x 109 generations, samplingevery 20,000 generations. Convergence of parameters and proper mixing were confirmedby calculating the effective sample size (ESS) in TRACER v.1.6, excluding theinitial 10% (burn-in) of each run. All considered parameters had ESS > 500.

Phylogenetic treeswere reconstructed with both maximum likelihood (ML) and Bayesian methods. Weanalysed each gene separately and also gGAPDH + 18S concatenated with SeaView(Gouy et al. 2010 10 ). For ML reconstructions in PHYML v.3.0, we used both theNearest Neighbour Interchange and the Tree Bisection and Reconnection algorithmsto improve tree searching. Branch supports were assessed via 1000 bootstrapreplicates.

We used a BayesianMarkov chain Monte Carlo (MCMC) method for species coalescent analysis basedon multilocus data (18S SSU and gGAPDH). We inferred both 18S SSU and gGAPDHgene trees, concatenated sequences tree, and also a species tree with *BEAST(Heled & Drummond 2010), which is included in the BEAST v.1.8 package. Informationin the literature about the taxonomy of DNA sequences retrieved from GenBankwas used to assign prior information on each sequence to a species. In the caseof sequences without taxonomic identification at the species level, we maintainedeach sequence as a single "fictitious species" (Supplementarydata, Table). We used Herpetomonas muscarumsequences to root phylograms.

The Yule-coalescentmodel of speciation was imposed in tree reconstructions to assign individualsto species. This is the simplest birth-death model of speciation, which considerseach tree node as a speciation process. Therefore, it is the best choice forphylogenies with many species represented by a few sequences in a species treebased on one or few molecular markers (Ogilvie et al. 2016 23 ). Number of independentruns, parameter sampling, and inspection of parameter convergence were identicalto RLC analysis. Runs were combined using LogCombiner and a maximum clade credibility(MCC) tree based on 10,000 trees (burn-in = 2,000) was generated for both genefragments with a posterior probability limit of 0.6 using Tree Annotator (bothpart of the BEAST package). Statistical support for clades was assessed by theposterior probability (PP) method. All resulting species trees were visualisedin Figtree v.1.4 (http://tree.bio.ed.ac.uk/software/figtree/).

Species distancematrices were estimated for both 18S SSU and gGAPDH fragments using the Tamura-Neisubstitution model, available in MEGA 5. Distance variances were estimated with1,000 bootstrap pseudo-replicates.

The parametersthat we believed represented valid species included sequences that (i) clusteredinto a single and well-supported monophyletic clade (i.e., bootstrap ≥ 60% and posterior probability ≥ 0.8), and (ii) were genetically more distant from another species instead ofthe minimal genetic distance observed for two bona fide Trypanosomaspecies.

Estimation ofdivergence times - We tested single and multiple calibration points basedon previous phylogenetic data (Stevens & Rambaut 2001, Lewis et al. 2011 15 )to infer divergence times. Log marginal likelihood Bayes Factor (LBF) was usedto compare the marginal likelihood of each hypothesis, estimated with the pathsampling and stepping stone algorithms (Baele et al. 2013 4 ). We also sampledfrom all prior hypotheses (i.e., analysis with only priors and no sequencedata) to make sure that all priors were proper and together, they did not producean unexpected joint prior. Moreover, these results were used to estimate theinfluence of the prior and the sequence data on posterior results.

Ethics -The mammal capture was licensed by the Brazilian Institute of Environment andRenewable Natural Resources (IBAMA/CGFAU/LIC; license 3665-1). Tissue samplecollection and euthanasia were performed as regulated by the Federal Counselof Medical Veterinary under resolution number 1.000 approved on May 11th, 2012,and the procedures were approved by the FIOCRUZ Committees of Bioethics (LW81/12).

 

RESULTS

Parasite viability,growth behaviour, and morphology - The in vitro parasite growth assaygenerated a very short log phase and an absent stationary phase. As shown inFig. 1, parasite populations from the spleen and liver remainedunchanged in the first 24 h, followed by a rapid increase in parasite populationson day 2. Both parasites (isolates 17665B and 17665F) achieved a maximum growthrate at day 3, followed by a sharp decline. Based on these results, we establishedthat parasites should be passaged on the third day of culture. The growth curvewas performed only until day 4 because the parasite population started to decreaseand present more degenerative forms than viable parasites after that. In anattempt to find a nutrient medium which would promote better growth of the parasitesin vitro, we tested five different nutrient media (DMEM, EAGLE, TRPMI,RPMI 1640, and LIT) supplemented with 10% FBS, as well as a Vero cell monolayerculture. Neither the tested culture media nor the Vero cell culture succeededin the long-term maintenance of the parasites in culture. The maximum lengthof time in which motile parasites were observed was 48 h, although most of theparasites were already dead after the first 24 h. On the third day, no livingcell was observed in any of the tested media.

 

 

Another approachwas the morphological analysis of these parasites, which was performed on day2 of the growth assay (exponential phase). In Giemsa-stained culture smears,only epimastigotes could be observed (Fig. 2). The cellshad the characteristic kinetoplast localised in the anterior portion of theparasite, near the nucleus. The established culture was fairly homogeneous inform, but not in size, showing large variation in body size.

Ultrastructureand morphometric analysis - Morphometric analysis using SEM images of theisolated parasites revealed no significant differences (p > 0.05) betweenthe measured values of the two populations, 17665B and 17665F. For the two populations,the total length of the parasites was reported as [mean ± standard deviation(SD)] 23.3 ± 5.9 µm and 28.7 ± 6.2 µm, respectively,the length of the free portion of the flagellum was 9.5 ± 3.2 µmand 11.6 ± 3.0 µm, respectively, and the body length was 13.8 ±3.6 µm and 17.1 ± 3.7 µm, respectively. The high SD valuesdemonstrate the differences observed in parasite size among distinct cells inthe same population. The morphology of all the cells was compatible with theepimastigote form of the parasite, showing the emergence of a lateral flagellum(Fig. 3A-B).

Cells from bothcultures (spleen and liver) presented an elongated body with the nucleus inthe anterior portion and a single flagellum as revealed by TEM (Fig.3C). The kinetoplast shows similar stem morphology (Fig.3C-D) in a well-defined region of the mitochondria, near the flagellar pocket,compatible with the epimastigote form of the parasite. Mitochondria are branched,appearing throughout the cell body with double membranes and mitochondrial cristae(Fig. 3C-E). The structure of the Golgi is similar to thatfound in eukaryotic cells (Fig. 3E, inset), with a set ofstacked cisternae located close to the kinetoplast. The point of emergence ofthe flagellum was also observed. Moreover, an elongated, cellular, electron-densestructure was observed, with a simple membrane either present in small amountsor widely distributed throughout the parasite body, usually in sets of threeor four (Figs 3C-D).

Molecular characterisationand phylogenetic inferences - DNA extracted from both parasite populationswere PCR-amplified using 18S SSU and gGAPDH targets. The DNA sequences obtainedfrom the parasite populations derived from the spleen and liver were identicalfor the two targets, indicating that both isolates belong to the same species/genotype.BLASTn results using these sequences showed high similarity with T. wauwau(similarity > 95%; coverage > 99%), a species recently described inNeotropical bats (GenBank: KT030800, KT030801, KT030810, and KT030821). Thesequences also showed similarity to other Trypanosoma species detectedin marsupials of the families Potoroidae and Macropodidae from the West andSoutheast parts of Australia (GenBank: JN315381, JN315382, JN315395, JN315396,AJ009168, and AJ620276 for T. noyesi, and KC753537, KC812988, KU354263,and KU354264 for Trypanosoma sp. sequences).

In addition tothose genetically similar sequences, we added sequences for other Trypanosomatidaeinto the phylogenetic analysis to obtain a better picture of the evolutionaryposition of our isolates [Fig. 4, Supplementary data(Table)].

Nonsignificantsubstitution saturation was found in both 18S and gGAPDH sequences (ISS= 0.06, ISS.C = 0.70, p < 0.0001, for 18S sequences;and ISS = 0.17, ISS.C = 0.72,p < 0.0001, for gGAPDH sequences). No sign of recombination was found inthe 18S sequences, but distance plots demonstrated at least three crossoverpoints in the gGAPDH sequences of the Australian Trypanosoma sp. specimens(strains AB2013 G4, G5, and G7) and T. microti, which might indicatepossible recombination hotspots (Supplementary data,Fig. 2). The first crossover point at position160 of the gGAPDH alignment was statistically significant (p < 0.001).

Phylogenetic reconstructionscorroborated what was previously observed in recombination analysis. Indeed,preliminary analyses with ML and BI, assuming a strict clock, displayed incongruenttopologies for 18S and gGAPDH sequences. Trypanosoma (Schizotrypanum)and T. rangeli clades were monophyletic in a well-supported clade inthe 18S trees (BP = 71%; PP = 1.00; Fig. 4A),but paraphyletic in the gGAPDH trees. The exclusion of three Australian Trypanosomasp. gGAPDH sequences (GenBank: KC812985, KC812986, and KC812987) from the analysisproduced an identical topology to that observed in the 18S tree, i.e.,showing the monophyly of Schizotrypanum and T. rangeli clades.

RLC analysis indicatedthat gGAPDH sequences evolve almost twice as fast as 18S sequences (2.04 x 10-3and 1.09 x 10-3 substitutions/(site million years), respectively).Moreover, both datasets exhibited changes in mutation rates across samples (meanrate changes of 2.67 and 2.55 for gGAPDH and 18S sequences, respectively). Therefore,the implementation of a 'relaxed' clock was more suitable to our 18S and gGAPDHdatasets. Indeed, when a relaxed clock was assumed, gGAPDH and 18S topologieswere perfectly congruent (Fig. 4B).

It is noteworthythat irrespective of the methods used, all phylogenetic analyses showed thatthe parasite species described herein belongs to the T. cruzi clade,grouping with T. wauwau in a well-supported clade (BP = 100%; PP = 1.00).Both species are exclusive and reciprocally monophyletic. This clade (T.janseni n. sp. + T. wauwau) is a sister group to the Australian trypanosomes,T. noyesi and Trypanosoma sp. (BP = 100%; PP = 1.00).

A multispeciescoalescent model in *BEAST with a 'relaxed' clock and ML tree with concatenateddata was consistent with gene tree reconstructions assuming a 'relaxed clock'and thus, provided a good fit to the 18S SSU and gGAPDH data (Fig.4C-D). Our samples were clearly separated from T. wauwau (BP = 100%;PP = 1.00). Moreover, genetic distances between T. janseni and T.wauwau, based on the 18S SSU and gGAPDH sequences [(mean distance ±variance) 1.7 ± 0.5% for 18S SSU and 9.7 ± 1.2% for gGAPDH], weresimilar to those observed between T. erneyi and T. dionisii (2.4± 0.7% and 9.2 ± 1.2%, respectively), T. erneyi and T.cruzi marinkellei (1.8 ± 0.6% and 9.5 ± 1.3%, respectively),and T. conorhini + T. rangeli and T. vespertilionis (2.4± 0.6% and 7.0 ± 3.5%, respectively). Altogether, these resultsprovide strong evidence that T. janseni represents a bona fidespecies within the T. cruzi clade.

Estimated datesfor the evolutionary history of the T. cruzi clade - MCMC chains did notconverge to a stationary distribution in important parameters of the tree (suchas posterior, likelihood, and clock rate) when we used the single calibrationpoint based on the split between T. cruzi and T. cruzi marinkellei,sampling more chains, optimising parameters from previous runs, or increasingthe number of chains. Therefore, this model was excluded from the analysis.

Marginal likelihoodvalues (and consequently, model probabilities calculated with LBF) were quitesimilar for the trees estimated with a single calibration point based on thesplit between Trypanosoma (Schizotrypanum) and T. rangeliclades. With respect to two calibration points, the same was observed basedon the split between Schizotrypanum and T. rangeli clades andbetween T. cruzi and T. cruzi marinkellei (Table).

According to theoutgroup comparison, the T. cruzi clade was estimated to have taken placeat ~81 Mya (CI: 123-52 Mya; Fig. 4D). The separationof the South American T. wauwau + T. janseni from the AustralianT. noyesi clade (T. noyesi + AB2016a + AB2013 G8) was dated at~ 26 Mya (CI: 40-15 Mya).

Class Kinetoplastea Konigberg, 1963

Order Trypanosomatidae (Kent 1880) Holande, 1952

Family Trypanosomatidae Doflein, 1901

Genus Trypanosoma Gruby, 1843

Trypanosoma janseni n. sp. (Figs 2-3)

 

 

 

 

Description- Only epimastigotes were observed in culture. The evolutive form was confirmedby the position of the flagellum output. T. janseni n. sp. cells showeda small and elongated appearance, with a slightly twisted body and a singleflagellum, a nucleus in the posterior portion, and a kinetoplast showing similarstem morphology localised in the anterior portion of the parasite, near thenucleus. Mitochondria appear branched throughout the cell body with double membranesand mitochondrial cristae. The structure of the Golgi is similar to that foundin eukaryotic cells, with a set of stacked cisternae located near the kinetoplastand the output of the flagellum. We also observed various endosomal vesiclessimilar to acidocalcisomes positioned near the flagellar pocket. Additionally,there was an elongated, cellular, electron-dense structure, with a simple membranepresent either in small quantities or widely distributed throughout the protozoanbody, usually in sets of three or four.

Type host- Didelphis aurita (Wied-Neuwied, 1826)

Site in thehost - Spleen and liver

Type locality- Atlantic Rainforest biome, Rio de Janeiro/RJ, Brazil (22º56'27.89''S;43º24'23.17''W).

Type data anddepository - Hapantotype; cultures of parasite populations derived fromthe spleen and liver are deposited in the Coleção de Trypanosomade Mamíferos Silvestres, Domésticos e Vetores, COLTRYP/FIOCRUZ(www.coltryp.fiocruz.br)under the accession numbers COLTRYP 715 and 716. The newly generated sequenceswere deposited in the GenBank database under the accession numbers KY243025and KY243026 (18S rRNA gene) and KY549444 and KY549445 (gGAPDH gene).

Vector -Unknown

ZooBank registration- In accordance with section 8.5 of the International Code of Zoological Nomenclature(ICZN), details of the new species have been submitted to ZooBank with the lifescience identifier (LSID) zoobank.org:pub: 61AA3E83-7ECB-408B-8955-499EEBF4EAC4.

Etymology- The name T. janseni n. sp. was given in honour of Dr Ana Maria Jansen,a distinguished parasitologist from Brazil, who worked for several years withopossums as experimental models. Having an animal facility to study this mammalianhost not only provided her with material for several manuscripts, but also 15years of legal issues until she proved that everything was properly, legally,and ethically conducted. With this new species name, we recognise the meritof a researcher that persistently and painstakingly endeavoured to investigateall possible factors involved in the very complex trypanosome life-cycle.

 

DISCUSSION

Small mammals havebeen described as important reservoirs of different trypanosomatids such asT. cruzi, T. rangeli, and Leishmania spp. In recent years,several studies have investigated the importance of these hosts in maintainingthe sylvatic cycle of the Leishmania spp. The role of the Didelphisspp. as Leishmania reservoirs was already proposed. In studies withLeishmania spp., the gold standard for isolating parasites included thecultivation of hematopoietic tissues (such as the spleen, liver, and bone marrow)using Schneider's media with 10% FBS. On the other hand, T. cruzi isusually isolated from blood samples of the infected hosts. Aiming to diagnoseand isolate both trypanosomatids, we routinely cultivate blood, liver, skin,and spleen samples from small mammals that we capture during field work. Here,we isolated parasite populations from the blood, spleen, and liver samples froma D. aurita captured in the Atlantic Rainforest of Rio de Janeiro, Brazil.This mammalian host displayed mixed infection, as it was infected with T.cruzi in the blood (as demonstrated by haemoculture; data not shown) andwith the new species described here, in the spleen and liver. Its vectoris unknown, but it is probably associated to marsupials because dixenous haemoflagellatesusually adapt their transmission to ecological conditions and environment oftheir hosts. Alternatively, T. janseni may be a monoxenous trypanosomatidin the process of pre-adaptation to a mammal host (Lukešet al. 2014).

It was very surprisingthat a new Trypanosoma species could be described in one of the mostintensively studied reservoirs of T. cruzi and Leishmania spp.in the last century. One hypothesis is that the co-infection with T. cruzimay have altered the host immune responses, resulting in a population increaseof T. janseni n. sp., allowing its isolation in axenic culture. Anotherpossible reason for the detection of T. janseni n. sp. is the fact thatwe were searching for Leishmania parasites, which led us to use Schneider'smedium complemented with 10% FBS as culture media. It is also possible thatif the same tissues of this host were also infected with the Leishmaniasp., the in vitro growth of the latter probably would have supplantedthe population of T. janseni, hampering its detection. Furthermore, moststudies aiming to identify Trypanosoma parasites investigate only bloodsamples, as observed in the Australian studies of trypanosomes (McInnes et al.2011). Multiple variables can act as a biological filter by selecting the parasiticpopulations that could be isolated, both inside a host or under in vitroconditions. Our success in the isolation of this new species of Trypanosomawas the result of a combination of factors, and we are thus not able to notethe most important one.

Another importantpoint is that we know nothing yet about the biology of this new parasite innature. We are also unsure if the epimastigote form observed in the culturewas the unique form. However, a trypomastigote form is surely expected and webelieve that T. janseni n. sp. also possesses this morphological formunder particular conditions in the host and/or axenic culture that we were unableto mimic under laboratory conditions. We tried different culture media, butdid not observe a transition from epimastigote to other morphological forms.In addition, we cannot infer anything about the parasite's course of infectionin a host co-infected by another Trypanosoma, as was the case with T.cruzi. Even if T. janseni n. sp. has the ability to colonise othertissues, such as blood, the presence of T. cruzi could be influencingthis colonisation. However, having been isolated in highly vascularised tissues,parasitic populations present in the spleen and liver could be present in themicrovessels of those tissues. Spleen microvessels, for example, are a commonsite for T. musculi infections (Albright et al. 1999 1 ). In a preliminaryexperiment, we infected cultured peritoneal macrophages, but infection was notestablished (data not shown).

SEM images confirmedthat the morphological characteristics are similar to axenic epimastigote forms.The morphometric analysis also showed an intense polymorphism, with widely rangingbody length and width between cells in the same isolate. The cells showed asmall and elongated appearance, with a slightly twisted body, similar to T.livingstonei (Lima et al. 2013 16 ). All Trypanosoma parasite stagespresent a single flagellum that emerges from the anterior region, kinetoplast,branched mitochondria, flagellar pocket, basal bodies, subpellicular microtubules,paraflagellar rod, and acidocalcisomes (de Souza 2008 7 ). The ultrastructure analysisshowed most of these structures, along with an unidentified cell structure withno parallel in other trypanosomes, which presented as single membrane structuresusually grouped in stacks of three or four. To the best of our knowledge, thestructure we observed herein (although displaying differences in size and shape)resembles the glycosomes of the Phytomonas sp. isolated from Euphorbiacharacias (Attias & de Souza 1995). However, further studies must becarried out to characterise this structure.

Another componentof the integrated approach employed here was the molecular taxonomy based ona partial region (737-bp) of the small subunit (18S) ribosomal RNA gene and708 bp of the nuclear marker glycosomal glyceraldehyde-3-phosphate dehydrogenase(gGAPDH) genes. These targets are the most suitable choice to establish thetaxonomic portion of a given trypanosomatid flagellate (Lukešet al. 2014). Phylogenetic analyses of both markers with a 'relaxed' clock andconcatenated data showed similar topology, with small discrepancies. In general,gene, concatenated trees, and species trees showed the following: (i) the populationsof the spleen and liver parasites belong to the same species; (ii) this speciesclusters with T. wauwau, a species recently described in Neotropicalbats (Lima et al. 2015 17 ), in a well-supported clade where they are exclusiveand monophyletic; and (iii) the genetic divergence between T. wauwauand T. janseni is similar to those observed for interspecific comparisonsamong trypanosomes. Cumulatively, these findings clearly demonstrate that theseisolates represent a new Trypanosoma species (Votýpka et al. 2015 29 ).

Phylogenetic MLand Bayesian trees based on concatenated data showed T. livingstoneicloser to the rest of the T. cruzi clade, followed by T. lewisiand T. microti, and Australian trypanosomes (AB2013 G1, G2, G3, G4, G5,and G7, and Trypanosoma sp. wombat) as a basal clade, such as previouslyobserved by Lima et al. (2015). Gene and species trees with a 'relaxed clock'clustered T. lewisi + T. microti and Australian trypanosomes togetherin a clade, and T. livingstonei sequences were positioned in a basalclade. Our results did not fully elucidate if this discrepancy was due to therecombination in gGAPDH sequences of these specimens, or if the rapid evolutionof these lineages possibly distorted the topology of the tree, by long-branchattraction (Noyes & Rambaut 1998).

Most clonal speciesundergo recombination (Omilian et al. 2006 24 ), but it seems to be infrequent inthe trypanosomatid taxa (Weir et al. 2016 30 ). We observed one crossover pointat position 160 of the gGAPDH sequences in the Australian Trypanosoma sp.specimens (strains AB2013 G1, G2, G4, G5, and G7) and T. microti, whichpromotes significant topological incongruence (p < 0.001). However, excludingthis and neighbour sites, Trypanosoma (Schizotrypanum) and T.rangeli groups remained paraphyletic, T. lewisi + T. microtiand Australian trypanosomes were clustered in a monophyletic clade, and T.livingstonei sequences were maintained in a basal clade. Only the exclusionof the Australian Trypanosoma spp. gGAPDH sequences mentionedabove or the application of a 'relaxed' clock grouped T. (Schizotrypanum)and T. rangeli as a monophyletic clade. Resolving the correct phylogeneticposition of these Australian samples, T. livingstonei, T. microti,and T. lewisi will require additional sampling, inclusion of more outgroups,and analysis of more nuclear markers.

Mutation rate estimationsunder the 'relaxed' clock assumption are very close to previous estimationsof 18S and other nuclear trypanosomatid sequences (Stevens & Rambaut 2001,Lewis et al. 2011 15 , Weir et al. 2016 30 ). Assuming a 'relaxed' clock model, it waspossible to observe that, in general, gGAPDH evolves twice as fast as 18S.

Fossils and geologicaldata suggest that marsupials have originated in South America and dispersed,together with their parasites, several times between the Americas and Australiaduring the period when both formed a southern supercontinent along with Antarctica(McInnes et al. 2011 19 ). Some authors also believe that marsupials play an importantrole in the diversification of trypanosomatids because they are natural hostsfor dixenous parasites. Some monoxenous species could also pre-adapt to thedixenous life cycle as they can survive and multiply in the marsupial anal scentglands, which protects them from the host immune system and provides an ideallower body temperature environment (Lukešet al. 2014). Our phylogenetic analysis dated the separation of the AustralianT. noyesi and South American T. janseni n. sp. and T. wauwauat ~26 Mya (CI: 40-15 Mya). The upper limit calculated here coincides with theopening of the South Tasman Rise and Drake Passage, which led to the completeseparation of Australia, Antarctica, and South America around 45-40 Mya. Thisresult would possibly explain why some Australian trypanosomes present highsimilarity with T. janseni n. sp., a trypanosome isolated from a SouthAmerican marsupial.

The presence ofT. janseni n. sp. in the same clade as the recently described bat trypanosome,T. wauwau, re-opens the question of the origin of the Trypanosomaspecies from indigenous Australian mammals. Instead of having evolved froma bat trypanosome, as proposed by Lima et al. (2015), those Australian marsupialTrypanosoma species might have originated from South American trypanosomes,as suggested by the similarity between T. janseni and some of the Australiantrypanosomes. In this proposed scenario, South American marsupials infectedwith the ancestral lineage of T. janseni and T. wauwau would haveentered Australia, and the parasite would have dispersed among Australian marsupials.Later, diversification gave rise to T. janseni and the closely relatedTrypanosoma species in Australian marsupials, which further gave riseto T. noyesi. T. janseni would have been maintained in marsupialspecies from the southern supercontinent, and a spill-over event to bats froma common ancestral lineage would have formed T. wauwau and the otherrelatives of the bat Trypanosoma species. In this scenario, T. janseni,along with T. wauwau, are the "missing-links" that shed light on theinitial diversification process based on what we currently known about the T.cruzi clade.

The history ofthe evolution of trypanosomes has been changing, in tandem with the emergenceof molecular tools with more discriminatory power. The total Trypanosomauniverse from the T. cruzi clade is immense, subsampled, and therefore,still unknown. The more effort that is put into the search for trypanosomatids,more pieces of this puzzle will be revealed. Considering the information obtained,the hypothesis proposed here appears to be a more parsimonious explanation forsome of the most ancestral species within the T. cruzi clade.

In conclusion- The description of a new parasite species must include, at least, morphologicaland molecular data to follow the "best practices" of the International Codeof Zoological Nomenclature. In this work, we integrate classical taxonomy andphylogenetic tools to characterise a new species of parasite, T. janseni.This species was isolated from the spleen and liver tissues of the opossum D.aurita in the Atlantic Coastal Rainforest of the Rio de Janeiro municipalityin Brazil. The parasite populations were isolated and maintained in Schneider'smedium complemented with 10% (v/v) FBS and 2% (v/v) human male urine. It wasdetermined that the parasite should be passaged on the third day of in vitroculture. Only epimastigote forms were observed, and the ultrastructure analysisshowed most of the structures described in the other species within the Trypanosomagenus. Additionally, an unknown feature, which presented as single membranestructures usually grouped in stacks of three or four and very similar to theglycosomes of the Phytomonas sp. isolated from Euphorbia characias,might be a diagnostic characteristic for T. janseni. Phylogenetic analysesof the partial region of the lower ribosomal subunit 18S gene and gGAPDH confirmedT. janseni as a new species within the T. cruzi clade and showedthat this species clusters with T. wauwau in a well-supported clade,with a close relationship to trypanosomes isolated from Australian marsupials.

 

ACKNOWLEDGEMENTS

To Marcos Antôniodos Santos Lima and Carlos Alberto Ardé for providing technical supportand to the PDTIS/Fiocruz sequencing platform for sequencing our samples. Wealso thank our friends and co-workers at LABTRIP, LBPMSR, and CFMA.

 

AUTHORS' CONTRIBUTION

CMTL and ALRR conceivedand designed the experiments; RFSMB and MCSP designed, performed, and analysedthe ultrastructure analysis; CMTL performed and analysed the biological andmolecular characterization; MGP conceived and performed the phylogeography analysis;CMTL, MGP, and ALRR wrote the manuscript. All the authors read and approvedthe final manuscript. The authors declare that they have no competing interests.

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Financial support: FIOCRUZ, CNPq, FAPERJ.
ALRR and RFSMB were u201cJovem Cientista do Nosso Estadou201d provided by FAPERJ; a post-doctoral grant was provided by CAPES to MGP.
+ Corresponding author: roque@ioc.fiocruz.br
Received 25 July 2017
Accepted 18 September 2017

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