Wildlife reserves of Mozambique included in this study

Studies on tsetse flies in MZ were conducted at the Gorongosa National Park (GNP) (18°45′58″S 34°30′00″E) and Niassa National Reserve (NNR) (12°08′35″S, 37°40′08″E). GNP comprises an area of ~4000 km² (10,090 km² including the buffer zone) located in the centre of MZ in the Province of Sofala, at the southern end of the Great African Rift Valley (Fig. 1). Regular seasonal flooding contributes to the variety of ecosystems in GNP, with grassland plains, savannah, floodplains and rain forests, surrounded by mountains and limestone cliffs. In the past, this area supported uniquely large numbers of large mammals, but they were drastically reduced by the long lasting civil war at the end of the twentieth century. A wildlife restocking program has been introduced, and large herbivores such as Cape buffalo, antelopes and elephants have been relocated from other Mozambican and South African parks. At the time of tsetse capture, GNP harboured large herds of antelopes (waterbucks, impalas, bushbucks, reedbucks, nyalas, kudus), Cape buffalo, an increasing number of elephants and hippopotamus, and a notable abundance of warthogs. Livestock is virtually absent within GNP, but goats are present in bordering areas.

NNR is a large wilderness area covering over 42,000 km² located in the northern Provinces of Cabo Delgado and Niassa bordering Tanzanian Game Reserves (Fig. 1), allowing animal migration across the Trans-Frontier Conservation Area. NNR is mostly covered by miombo forest, open savannah and wetlands, with high wildlife diversity. There is a great concentration of elephants, antelopes (impala, kudu, sable, waterbuck, hartebeest, nyala, wildebeest, duiker and eland among others), Cape buffalo, hippopotamus, zebras, wild suids and great felines. GNP and NNR are home to a large diversity of small mammals, birds, amphibians and reptiles, including large populations of crocodiles. Game hunting, agricultural and livestock (mainly goats) occur in this area. NNR and GNP are separated by ~900 km.

Tsetse fly collection, identification and microscopic screening for trypanosomes

Tsetse flies were captured in highly infested areas of GNP (2007, 2009 and 2012) and NNR (2013 and 2014) (Additional file 1: Table S1 and Additional file 2: Table S2) in the morning and in the afternoon using slow-moving vehicles to attract the flies that were then manually collected. Flies were cleaned twice by immersion in sterilised water, dried on filter paper, dissected, and guts microscopically examined for the presence of trypanosomes. Samples of guts and mouthparts of flies selected by microscopy, and from flies that had not been previously microscopically surveyed, all preserved in ethanol, were examined in this study. Tsetse flies preserved in ethanol (99.5%) were identified by morphology and cytochrome c oxidase subunit 1 (cox1) barcoding [28].

Blood collection from wild and domestic ruminants in Africa and South America

Blood was collected from Cape buffalo in the GNP, from wildebeests and Cape buffalo from NNR, and from antelopes in a game reserve at the province of Sofala. Blood samples from livestock were from small herds of cattle and goats, collected in the provinces of Tete, Sofala (Centre) and Maputo (South) of MZ. Livestock was raised having rare contact with wild ruminants, except in areas bordering conservation areas and game reserves, where wild animals and livestock frequently interact (Fig. 1, Additional file 1: Table S1).

Molecular identification of trypanosomes in ungulate blood samples and tsetse flies

Aliquots (~1.0 ml) of blood were collected using EDTA-treated tubes, preserved in ethanol and employed for DNA preparation as previously described [8, 20, 21]. Trypanosoma vivax diagnosis in ungulates was conducted using a T. vivax-specific PCR (TviCATL-PCR) [20]. Tsetse flies (guts and mouthparts) were tested using the fluorescent fragment length barcoding (FFLB) and TviCATL-PCR methods [29, 30]. We previously standardised this method using reference species to define the fluorescence peak profiles for the agents of African Animal Trypanosomiasis using an ABI3500 Genetic Analyser (Garcia et al. in preparation). Tsetse DNA samples that were positive for T. vivax using FFLB were submitted to whole genome amplification (WGA) using the REPLI-g UltraFast Mini Kit (Qiagen, Hilden, Germany).

PCR amplification and sequencing of gGAPDH and ITS rDNA sequences

PCR amplification of gGAPDH sequences from DNA from tsetse flies and blood samples was conducted using a nested PCR method [31]. Each amplified DNA fragment was cloned, and a different number of clones was sequenced for each sample. This procedure was necessary, as infections with multiple trypanosome species and genotypes are common in tsetse flies. Sequences of ITS rDNA (ITS1 + 5.8S + ITS2) were amplified by PCR (~600 bp), cloned and sequenced (~5–7 clones from each sample) using primers and PCR conditions described previously [19]. PCR-amplification of ITS rDNA sequences from T. vivax and T. vivax-like obtained from tsetse flies had a lower sensitivity compared with FFLB and required cloning and sequencing of several clones from each sample. Sequences obtained were deposited in GenBank (Table 1, Additional file 1: Table S1, Additional file 2: Table S2).

Comparison of gGAPDH and ITS rDNA sequences by multidimensional scaling (MDS) analysis

To provide a visual representation of the level of similarity across sequences in the dataset, we carried out multidimensional scaling (MDS) analyses plotted in 2D and 3D using the R software platform (R Development Core Team, http://www.R-project.org). MDS was performed using the Bios2mds package [32].

Phylogenetic analyses of gGAPDH and ITS rDNA sequences

The whole gGAPDH sequences dataset was aligned and identical sequences from the same sample were removed from the final alignment, which also included sequences from GenBank of T. vivax and species of the subgenera Trypanozoon (T. brucei brucei, T. b. rhodesiense, T. B. gambiense and T. evansi), Pycnomonas (T. suis) and Nannomonas (T. congolense of the Savannah, Forest and Kilifi groups, T. simiae, T. simiae Tsavo and T. godfreyi). The final alignment was analysed using maximum parsimony (P) and the program PAUP*4.0b10, and maximum likelihood (ML) using RAxML with GTRGAMMA (500 maximum parsimony starting trees), model parameter estimated in RAxML over the duration of the tree search, and nodal support estimated with 500 bootstrap replicates.

Sequences of ITS rDNA (ITS1 + 5.8S + ITS2) obtained from DNA of tsetse and blood samples were aligned with sequences available in GenBank (Additional file 1: Table S1; Additional file 2: Table S2). The alignment was manually adjusted, and network split decomposition was inferred using the Neighbor-Net method with Kimura 2 parameters implemented as previously described [33, 34]. Internode support was estimated with 100 bootstrap replicates, using the parameters optimised for network inferences.

Tsetse fly identification and prevalence of T. vivax and T. vivax-like determined by fluorescent fragment length barcoding (FFLB)

Barcoded tsetse flies from GNP and NNR were identified as Glossina morsitans morsitans, which was the predominant species in both studied areas accounting for 85.5% of all flies captured, plus lower frequency (14.5%) of Glossina pallidipes. Similar prevalence of T. vivax and T. vivax-like was found for the two species of tsetse flies (Garcia et al. in preparation).

Prevalence rates of T. vivax and T. vivax-like in 151 fly mouthparts examined by FFLB ranged from 19.2% (29 positive flies of 151 examined) in GNP to 28.6% (93 positive flies of 325 examined) in NNR, and was not significantly different between the two-tsetse species (Garcia et al. in preparation). Altogether, results from GNP and NNR surveys unveiled FFLB profiles that were compatible with T. vivax in the mouthparts of 122 flies (25.6%) out of 476 flies examined, including 12 (2.5%) flies positive for T. vivax and T. vivax-like in both mouthparts and guts, and 8 (1.6%) flies showing only positive guts. Detection of T. vivax in fly guts suggests that parasites can be detected using sensitive diagnostic methods from a fresh blood meal or from contamination during fly dissection. However, as expected, FFLB of most gut samples did not reveal T. vivax or T. vivax-like (Garcia et al. in preparation). Isolates considered positive for T. vivax differed in some peaks of the whole FFLB profiles. However, Duttonella exclusive fluorescent peaks were always present.

Diagnosis of T. vivax and T. vivax-like in tsetse flies and ungulates by TviCATL-PCR

Analysis of MZ tsetse flies using TviCATL-PCR revealed infections in 43 out of 235 tsetse flies (18.2%) tested using this method, demonstrating that the sensitivity of this PCR method is inferior, but rather comparable to FFLB (25.6%, 122 out of 476 flies tested). In addition, TviCATL-PCR detected infection in 10 Cape buffalo (out of 98 animals tested) and one wildebeest (out of 15 animals). In Sunni antelopes (n = 5), reedbuck (n = 2) and warthogs (n = 7) the results were negative.

Comparison of Duttonella trypanosomes from East, West and South America by MDS and phylogenetic analyses of gGAPDH sequences

In previous studies, only five gGAPDH sequences of T. vivax were compared. These were distributed into the former groups: A and B, comprising Tanzania isolates; and group C, formed by isolates from SA (Brazil), WA (Nigeria) and EA (MZ) [25, 26]. In the present study, 65 out of 101 high-quality gGAPDH sequences (50 from GNP and 15 from NNR), obtained from 33 tsetse flies selected by FFLB and representative of the unveiled polymorphism, were compared by MDS. This analysis revealed a highly cohesive cluster, henceforward referred to as Tvv (the lineage including the reference T. vivax Y486), and a broader assemblage of sequences referred to as Tv-like (TvL) lineage, comprising a consistent cluster forward referred as TvL-Gorongosa, formed by groups A-D of isolates diverging from T. vivax Y486 by 4.8 to 5.7% (average of 5.2%) of gGAPDH sequence divergence. In addition, ~8.0% of sequences obtained from GNP and NNR tsetse flies remained unclustered, suggesting that they represent additional TvL lineages (Fig. 2a).

To obtain better resolved MDS clustering for the assessment of inter- and intra-cluster diversity, the dataset from each main cluster was analysed separately (Fig. 2b, c). Tvv unveiled four groups distributed in two major assemblages; one composed of TvvA clustered with TvvB, and the other composed of TvvC tightly related to TvvD (Fig. 2b). Average divergences of gGAPDH sequences were small: 0.2% for TvvA, 0.7% for TvvB, 0.5% for TvvC, and 0.8% for TvvD. TvvA (isolates from SA and WA livestock) diverged 0.7% from TvvB (WA and SA cattle isolates, and MZ cattle, buffalo and tsetse), 1.3% from TvvC (MZ tsetse), and 0.7% from TvvD (cattle, tsetse and nyala from MZ).

The highly divergent TvL sequences were arranged in four main clusters: TvLA (formerly referred to as “group A” [25] that clustered with the newly-identified group TvLB in addition to the cluster composed of TvLC and TvLD; altogether supporting TvL-Gorongosa lineage (Figs. 2 and 3). TvLA was closely related (1.5% of gGAPDH divergence) to TvLB, while TvLC tightly clustered with TvLD (2.0% of divergence). TvLA encompasses isolates from Tanzania and Kenya, and TvLB-D isolates are exclusively from MZ. TvLC and TvLD represented the predominant genotypes in tsetse flies from GNP and NNR (Figs. 2 and 3, Table 1).

The two major clusters were not arranged by order of wild or domestic hosts, tsetse species or date of collection. However, TvL-Gorongosa exclusively included isolates from EA, and Tvv comprised all samples from WA and SA, as well as some sequences from MZ and Ethiopia (Fig. 1, Table 1). Despite consistent results and high correspondence between MDS clusters and evolutionary lineages uncovered by gGAPDH sequences, with moderate support, the relationships among trypanosomes within the two lineages were not well-resolved, neither in P (Fig. 3) nor ML (data not shown) phylogenetic inferences using gGAPDH sequences. However, taken together, MDS and phylogenetic analyses allowed for the general delineation of 8 gGAPDH genetic groups as summarised in Table 1 (detailed information of each sample, host species and geographical origin are presented in Additional file 1: Table S1 and Additional file 2: Table S2).

In addition to the sequences that clustered into TvL-Gorongosa, 8 sequences from MZ tsetse flies remained isolated, although they were consistently included into the broad TvL assemblage. Similarly, a single sequence from Tanzanian tsetse, which was assigned to “group B” [25], largely diverged from all other sequences, however, its closest relatives appear to be members of Tvv.

Our analyses corroborated gGAPDH sequences as valuable markers for the identification of species, lineages and major intraspecific groups, but were unable to distinguish between the very closely related T. vivax genotypes circulating in SA, WA and some EA countries.

Genotyping and relatedness of Trypanosoma vivax and Trypanosoma vivax-like ITS rDNA sequences assessed by MDS

To further assess intra-lineage genetic diversity, we compared 263 ITS rDNA sequences determined herein with all available GenBank sequences (17) from MZ, Kenya, Tanzania and Ethiopia. The analysis included a large set of ITS sequences from WA (59 sequences from cattle) and SA (91 sequences from cattle, sheep, water buffalo and horses), most of them determined in the present study (Table 1, Additional file 1: Table S1, Additional file 2: Table S2). Analysis of ITS sequences that were representative of the whole genetic diversity, as well as of the geographical and host-species ranges revealed an extensive polymorphism in EA compared to WA/SA. Although the alignment of ITS sequences from EA isolates showed many ambiguities due to the extensive polymorphisms, blocks of ITS1 and ITS2 rDNA nucleotides characteristic for each group/genotype were detected (Additional file 3: Figure S1).

To investigate relatedness among the isolates, ITS sequences were submitted to MDS (Fig. 4). The results support a highly cohesive cluster of sequences representing Tvv, a broader arrangement of sequences supporting TvL-Gorongosa, and unclustered sequences representing other TvL lineages (Fig. 4a). In contrast to quite conserved gGAPDH sequences, large polymorphisms of ITS rDNA supported the separation between Tvv1 and Tvv2 genotypes. Tvv1clustered all WA isolates, while Tvv2 was restricted to SA, as unveiled by 3D MDS restricted to the whole set of SA/WA sequences (Fig. 4b) or comprising exclusively Tvv1 and Tvv2 (Fig. 4c) datasets. More relevant polymorphisms were observed within Tvv1 compared to Tvv2. Nevertheless, Tvv1 and Tvv2, which are to date restricted to livestock, share a greater degree of similarity between themselves compared with Tvv3 and Tvv4 of EA isolates from livestock, tsetse and wild buffalo. Two sequences from Ethiopian cattle clustered with Tvv1 and a single sequence was assigned to the new Tvv5 genotype (Fig. 4b). Tvv1 and Tvv2 are both genotypes of long-range dispersal linked to livestock: Tvv1 is the most geographically dispersed genotype occurring from the Gambia (WA) to Ethiopia (EA); Tvv2 is widespread in SA and was identified in Brazil, Venezuela, Colombia and French Guiana (Fig. 1).

Clustering of TvL ITS sequences analysed by 3D MDS support the highly heterogeneous lineage TvL-Gorongosa as defined by gGAPDH sequences (Fig. 4c, Table 1). Furthermore, ~8.0% of EA sequences from tsetse flies did not consistently nest within any cluster (Fig. 4a).

ITS rDNA network of T. vivax and T. vivax-like isolates

To investigate both the relationships and the possibility of recombination generating the remarkable polymorphic ITS sequences, we submitted the dataset to network split decomposition. All sequences from WA and SA clustered tightly together, whereas EA sequences largely dispersed in a complex and reticulated network (Fig. 5a).

The split between SA and WA sequences corroborated the MDS results and showed more relevant polymorphisms in Tvv1 compared to Tvv2 (Fig. 5b). Each isolate from WA exhibited a unique ITS sequence. Ethiopian isolates were assigned to Tvv1 and Tvv5, which are closely related to Tvv3 and Tvv4 from MZ, and to Tvv2 from SA (Fig. 5c).

Consistent with MDS analysis (Fig. 4), split decomposition network of ITS sequences uncovered greater diversity within TvL compared to Tvv. MDS clustering, network patterns, and polymorphisms on the aligned ITS sequences (Additional file 3: Figure S1) were all consistent with the lineage TvL-Gorongosa and its genotypes (Fig. 5c).

There was a high concordance between groups/genotypes defined by gGAPDH and ITS rDNA (Table 1). Genotypes TvL1 and TvL2 include isolates from tsetse, livestock and wild animals from MZ, whereas TvL3 clustered isolates from Kenyan cattle and Tanzanian buffalo. TvL1-TvL3 belongs to TvLA/B assemblage defined by gGAPDH, and TvL4-TvL7 are likely of TvLC/D grouping. Taking into account their close relationships and small degrees of sequence divergences, TvL1–7 are all considered genotypes of TvL-Gorongosa. This lineage comprises most ITS sequences obtained from tsetse and from wild ungulates from MZ and Tanzania, in addition to cattle isolates from Kenya and Ethiopia (Fig. 5c). Long branches in the network correspond to EA sequences assigned to TvL8–11 genotypes.

Results from our comprehensive analysis corroborate the high discriminatory power of ITS rDNA sequences allowing for the identification of known and novel genotypes. However, deeper nodes within Tvv and TvL remain uncertain. The ITS network displayed noticeable reticulation of TvL sequences and a moderate degree of reticulation among Tvv genotypes from EA, suggesting that EA populations may undergo genetic recombination in tsetse flies, whereas the small degree of reticulation among Tvv sequences from WA and SA suggest clonal populations.

Trypanosoma vivax and Trypanosoma vivax-like in tsetse flies and wild reservoirs from Mozambique

This is the first molecular survey of T. vivax in tsetse flies from MZ. Glossina m. morsitans was the predominant species, found in sympatry with a smaller population of G. pallidipes in GNP and NNR. These tsetse species are highly effective vectors of trypanosomes that are pathogenic to humans and livestock in the great Zambezi Valley and across EA [35–37]. Despite the absence of large-scale surveys on tsetse flies and animal trypanosomiasis in MZ, Glossina m. morsitans and G. pallidipes are known to be present from northern to central MZ. The investigations carried out in central provinces, including Sofala where GNP is situated, have shown that due to a high prevalence of Nagana, cattle production throughout large areas of Mozambique has to rely on preventive treatment against trypanosomiasis [38]. The present study aimed to characterise T. vivax isolates; a more substantial understanding of tsetse diversity and prevalence in MZ is beyond the scope of this work.

We investigated the presence of T. vivax in tsetse flies from GNP and NNR using the FFLB method [30, 39] previously employed for surveys of trypanosomes in tsetse flies from the wildlife reserves of Tarangire, Serengeti and Msubugwe in Tanzania [26, 29]. FFLB detected T. vivax and/or T. vivax-like in 25.6% of flies examined. The method of TviCATL-PCR [20] was herein confirmed as a simple, sensitive and specific diagnostic method, regardless of the parasite lineage/genotype or the existence of multiple trypanosome species. Despite being limited to a few samples, molecular surveys have identified T. vivax in a range of wild ungulates including antelopes, Cape buffalo, warthog and giraffe in Tanzania, Zambia and MZ [2, 24, 40, 41]. These findings corroborated morphological studies of blood trypanosomes reporting T. vivax and T. vivax-like in a range of wild ungulates in EA [42, 43].

Duttonella comprises T. vivax and T. vivax-like phylogenetic lineages constituted by a greater genotype repertoire in East Africa compared with West Africa and South America

There are growing amounts of molecular data uncovering a great repertoire of trypanosomes including new species and genotypes infecting tsetse flies in wildlife reserves. Novel trypanosomes from tsetse flies [7, 25, 26, 29, 30, 44–46] and African ungulates [2, 23, 24, 34] were reported in eastern and central-southern African wildlife conservation areas.

Here, inferences based on gGAPDH sequences strongly supported two major evolutionary lineages, Tvv and TvL, each one composed of four main groups. Relevant congruence among lineages/groups defined by gGAPDH and ITS rDNA sequences were demonstrated by MDS, phylogenetic trees and network approaches. Genotyping within each evolutionary lineage was better assessed using highly discriminatory ITS sequences. Tvv lineage includes the reference strain T. vivax Y486 of cattle from Nigeria [47]. This lineage comprises genotypes Tvv1, including all WA plus some EA sequences, Tvv2, exclusively from SA, and Tvv3-Tvv5, so far restricted to some EA samples. All analyses strongly supported a remarkable diversity within TvL formed by at least 11 genotypes represented, so far, by sequences from MZ, Tanzania, Kenya and Ethiopia [2, 27]. Most TvL genotypes were identified in tsetse flies, four were detected in wild ungulates, and only three genotypes were identified in livestock. Genotypes uncovered by our study almost certainly represented only a small part of the genetic diversity within the Duttonella subgenus.

Taking into account the remarkable diversity of Duttonella, and the intertwined network branching patterns of ITS rDNA sequences, it is tempting to speculate that in EA, new genotypes may have emerged by some recombination process during the parasite development in tsetse flies. Studies supporting clonal propagation of T. vivax in WA and SA populations have demonstrated the stability of predominant genotypes over time consistent with clonality [8, 22]. However, all data from WA and SA came from livestock production zones free of tsetse (SA) or with smaller tsetse fly density compared to GNP and NNR.

The subgenus Duttonella is a complex of species and genotypes

The gGAPDH phylogenetic inference strongly supports one monophyletic assemblage harbouring all isolates from tsetse flies and ungulates identified as T. vivax or T. vivax-like corresponding to the subgenus Duttonella. Phylogenetic inferences support Duttonella as the most basal clade, corroborating previous study results, indicating that it was the first lineage to diverge from the common ancestor of the phylogenetic clade T. brucei, which comprises the subgenera Duttonella, Nannomonas, Trypanozoon and Pycnomonas [19, 24, 48]. Representing the early branching African trypanosome, T. vivax has been explored by whole genomic and transcriptomic studies, showing significant differences from other African trypanosomes in the cell-surface developmentally regulated proteins and mitochondrial genomes, suggesting significant peculiarities in the parasite interaction with its ungulate hosts and vectors [49–51].

Phylogenetic positioning and sequence divergence separating T. vivax Y486 from TvL-Gorongosa justified its identification as another species of Duttonella. Our findings greatly expanded the species/genotype richness that has been discovered by phylogenetic analyses within the subgenus Duttonella. The possibility that genetic recombination has shaped the high genetic diversity observed in EA should be specifically addressed in further microsatellite, multilocus and whole genome studies. This is a very relevant question since the creation of genetic variants may produce undesirable parasite features related to pathogenicity, virulence and drug resistance, as well as being a source of novel genotypes for outbreaks [52].

Trypanosoma vivax and other species of Duttonella most likely arose in East Africa, from where genotypes adapted to livestock may have spread across Africa and South America

The greater diversity in EA conservation areas of trypanosome genotypes either highly similar or divergent compared with those from WA, and the basal position of TvL in phylogenetic trees, suggest that EA was the region of origin and diversification of Duttonella. Possibly, a range of genotypes emerged through recombination of genetically different parasites circulating between wild tsetse and several species of ungulates. Some genotypes may have adapted to livestock, then spread across EA and WA with the historical movement of infected livestock, accompanying human migration along sub-Saharan Africa. In Africa, T. vivax has a wide range of healthy domestic reservoirs, including native breeds of cattle, goats, sheep, donkeys, camels and even suids. Both migratory routes from WA to EA and vice versa may have played some role in the spreading of Tvv1 in cattle from The Gambia to Ethiopia. In MZ, we identified cattle infected with Tvv3 and Tvv4 genotypes that are highly closely related to Tvv1, but the existence of Tvv1 in this country, as well as in Tanzania and Kenya, must be assessed by examining more comprehensive cattle sampling.

In EA, cattle were found to be infected by Tvv and TvL genotypes as confirmed here in MZ and reported previously in Ethiopia, as well as in Zambia, Central-Southern Africa [7, 23, 25–27]. Despite the small sample, Ethiopian isolates clustered either into Tvv or TvL. The observation that some T. vivax genotypes from Ethiopian are closely related to WA genotypes [7] agreed with previous reports on isolates from MZ, which were different, but more related to those from WA than to those from EA (Tanzania and Kenya) [20, 24–26]. In addition to Ethiopia, Tvv genotypes closely related to those from WA/SA has been found in Zambia [23] and Uganda (Garcia et al. in preparation). Thus far, Tvv1 was the only one that has been found both in WA and EA, being highly prevalent in WA where the existence of other genotypes needs to be investigated. However, different from highly homogeneous SA populations of Tvv2 and despite tightly clustered together, substantial genetic diversity has been revealed among Tvv1 isolates, as demonstrated in this and in previous studies of isolates from The Gambia, Burkina Faso, Benin, Ghana, Nigeria and Cameroon [4, 8, 23, 24].

This study corroborated a tight relationship between Tvv1 (WA) and Tvv2 (SA) genotypes. However, to date, not a single ITS rDNA sequence from Africa was identical to those of Tvv2, which comprises all the 39 samples from Brazil, Venezuela, Colombia and French Guiana examined in our study. Our findings support the hypothesis that Tvv2 may be a bottlenecked genotype that recently diverged from closely related genotypes escaped out of WA. In a study based on microsatellite markers, we also demonstrated that T. vivax from WA and SA are highly similar, but not identical, and that diversity was far greater across WA than SA [8].

To date, all TvL isolates came from tsetse-infested areas in EA. Although an early study carried out in Ethiopia suggested that isolates from tsetse-free areas clustered with SA/WA genotypes, while those from tsetse-infested areas clustered with EA genotypes, analysis based on polymorphic ITS sequences did not support this hypothesis [7, 27]. In Ethiopia and other African countries, T. vivax is cyclically transmitted by tsetse flies and mechanically transmitted by other biting flies [6, 7, 27]. In SA, T. vivax is only mechanically transmitted by non-tsetse biting flies. Genomic studies comparing Venezuelan and WA T. vivax Y486 evidenced a drastic process of mitochondrial genome degradation in SA isolates, whereas the African T. vivax Y486 exhibited entirely functional mitochondria necessary for the development in tsetse flies [51].

Taxonomy and morphological, biological and immunopathological peculiarities of Duttonella trypanosomes in East Africa

Our study uncovered a complex of species/genotypes within the subgenus Duttonella. Molecular data provided herein corroborate differences between WA and EA T. vivax and among EA isolates regarding their morphology, infectivity to tsetse flies, pathogenicity and antigenic relationships. In the last taxonomical revision of Trypanosoma [42], in addition to T. (Duttonella) vivax vivax (= T. vivax) that is the type-species of the subgenus [42], T. (Duttonella) uniforme, was recognised as a valid species of Duttonella, characterised by small-sized blood forms. This species was reported in cattle, antelope, buffalo and giraffe, widespread in Uganda and Congo, and reported in Zululand, Tanzania and Ethiopia. T. (Duttonella) vivax ellipsiprymni was regarded as a subspecies due to the morphological intergradation of its blood forms with those of T. vivax [42, 53]. Trypanosoma uniforme and T. v. ellipsiprymni were thought to be transmitted by G. morsitans, G. palpalis and G. fuscipes in EA [42]. It is possible that these trypanosomes correspond to T. vivax-like reported in this and in previous studies [2, 7, 19, 23–26]. We previously described a T. vivax-like from a nyala antelope in MZ showing large blood trypomastigotes morphologically resembling those of T. v. ellipsiprymni [24]. This isolate was herein assigned to Tvv, and differ only by 0.7% of gGAPDH sequence divergence from T. vivax Y486.

Recently, T. (Pycnomonas) suis, which was initially phylogenetically positioned near T. brucei and provisionally named Trypanosoma sp. Musubugwe, was molecularly validated using DNA recovered from archived blood smears [46]. Isolates cryopreserved and archived collections of tsetse flies and ungulate blood smears of species of Duttonella can serve as DNA source for comparison with species candidates uncovered in this and other studies.

We identified different genotypes of Tvv and TvL in G. m. morsitans and G. pallidipes. Probably because G. morsitans was highly predominant in our study, a larger repertoire of trypanosomes was found in this species. Trypanosomes from Tanzanian G. pallidipes and G. swynnertoni [25, 26] slightly differed from those we identified in MZ tsetse flies. In accordance with long-range distribution, it was demonstrated that WA T. vivax developed in a range of Glossina species. In contrast, EA isolates of Duttonella likely prefer sympatric tsetse flies. Nevertheless, G. pallidipes was shown to be equally competent in experimentally transmitting T. vivax from Kenya and Nigeria [54, 55]. The presence of both TvL and Tvv in EA countries may be related to the coexistence of a range of tsetse species [56] feeding on many species of wild animals and large herds of livestock.

Corroborating the close relationships between T. vivax populations in WA, high antigenic cross-reactivity was reported for isolates from The Gambia and Nigeria, but not between these and distantly related Kenyan isolates. In addition, isolates from Kenya differed in resistance to drugs [57], antigenic cross-reactivity and pathogenicity. The severe hemorrhagic syndrome has, so far, been reported in Kenya and Uganda [10, 11]. Studies reporting a lack of significant differences in pathogenicity of T. vivax isolates from areas that were or were not infested by tsetse flies in Ethiopia did not include molecular characterization of the isolates [6]. Despite considerable immunopathological data on T. vivax infected cattle in Kenya, Ethiopia and Uganda, genetic diversity of a comprehensive sampling remains to be investigated.