Genetic recombination between pathogens derived from humans and livestock has the potential to create novel pathogen strains, highlighted by the influenza pandemic H1N1/09, which was derived from a re-assortment of swine, avian and human influenza A viruses. Here we investigated whether genetic recombination between subspecies of the protozoan parasite, Trypanosoma brucei, from humans and animals can generate new strains of human pathogen, T. b. rhodesiense (Tbr) responsible for sleeping sickness (Human African Trypanosomiasis, HAT) in East Africa. The trait of human infectivity in Tbr is conferred by a single gene, SRA, which is potentially transferable to the animal pathogen Tbb by sexual reproduction. We tracked the inheritance of SRA in crosses of Tbr and Tbb set up by co-transmitting genetically-engineered fluorescent parental trypanosome lines through tsetse flies. SRA was readily transferred into new genetic backgrounds by sexual reproduction between Tbr and Tbb, thus creating new strains of the human pathogen, Tbr. There was no evidence of diminished growth or transmissibility of hybrid trypanosomes carrying SRA. Although expression of SRA is critical to survival of Tbr in the human host, we show that the gene exists as a single copy in a representative collection of Tbr strains. SRA was found on one homologue of chromosome IV in the majority of Tbr isolates examined, but some Ugandan Tbr had SRA on both homologues. The mobility of SRA by genetic recombination readily explains the observed genetic variability of Tbr in East Africa. We conclude that new strains of the human pathogen Tbr are being generated continuously by recombination with the much larger pool of animal-infective trypanosomes. Such novel recombinants present a risk for future outbreaks of HAT.
Genetic recombination allows transfer of harmful traits between different strains of the same pathogen and enables the emergence of genetically novel pathogen strains that the host population has not previously encountered. This can be particularly important when a pathogen acquires a virulence trait that allows it to spread beyond its normal host population. Here we show that this happens among the single-celled parasites—trypanosomes—that cause human African trypanosomiasis (HAT) or sleeping sickness carried by the tsetse fly. Genetic recombination readily occurs between the human and animal parasites when they are co-transmitted by the tsetse fly, creating new pathogen genotypes or strains. There is a single gene that confers human infectivity and each of the genotypes that inherits this gene is potentially capable of infecting humans. In this way new strains of the human pathogen can be generated by recombination between the human-infective and animal-infective trypanosomes. Such novel recombinants present a risk for future outbreaks of HAT.
Citation: Gibson W, Peacock L, Ferris V, Fischer K, Livingstone J, Thomas J, et al. (2015) Genetic Recombination between Human and Animal Parasites Creates Novel Strains of Human Pathogen. PLoS Negl Trop Dis 9(3): e0003665. doi:10.1371/journal.pntd.0003665
Editor: Paul Andrew Bates, Lancaster University, UNITED KINGDOM
Received: December 1, 2014; Accepted: March 2, 2015; Published: March 27, 2015
Copyright: © 2015 Gibson et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This work was funded by The Wellcome Trust www.wellcome.ac.uk through project grants to WG and MB (079375, 088099). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Genetic recombination can generate new pathogen strains to which host populations have no prior immunity. This can have disastrous consequences; for example, the human population is at risk of an influenza pandemic caused by recombination between viruses derived from humans and domestic livestock. Microbial genetic recombination facilitates the transfer of genes for virulence and drug resistance into new genetic backgrounds, potentially creating pathogen strains with novel phenotypes as well as accelerating the spread of drug resistance. Among eukaryote pathogens, the impact of sexual reproduction is hard to predict, because of the wholesale mixing of genes from different strains.
Trypanosoma brucei is the protist parasite responsible for the vector-borne disease human African trypanosomiasis (HAT) or sleeping sickness. In East Africa the disease is a zoonosis caused by T. b. rhodesiense (Tbr) which is morphologically indistinguishable from the non-human infective subspecies, T. b. brucei (Tbb). Both subspecies may occur in the same range of wild or domestic mammalian hosts and there has been a long-standing controversy about their identification . This was resolved by the discovery that human infectivity in Tbr was governed by expression of a single gene (Serum Resistance Associated, SRA)  and the presence of the SRA gene now serves as a convenient marker for Tbr [3–5].
Clearly, transfer of this single gene could potentially generate new strains of human-infective trypanosomes, and this has been demonstrated experimentally by transfection of the SRA gene into Tbb, resulting in a trypanosome with a human-infective phenotype . Population genetics analyses have failed to find consistent genotypic differences between Tbr and Tbb, other than presence/absence of SRA, and the idea that Tbb and Tbr are freely interchangeable by transfer of SRA has become central to the interpretation of population genetics data for Tbr and Tbb ; evidence of genetic admixture between Tbr and Tbb from recent genome comparisons of the two subspecies also supports this interpretation [7,8]. Genetic exchange in T. brucei occurs in the insect vector, the tsetse fly (genus Glossina)  and recent results show that it has the hallmarks of conventional eukaryote sexual reproduction: meiosis and production of haploid gametes [10,11]. All subspecies of T. brucei, including Tbr, have been shown to express meiosis-specific genes . Genetic crosses between Tbr and Tbb have been carried out in the laboratory [12–14], but analysis of the progeny was carried out before the significance of SRA was recognised and presence/absence of the gene was not determined. Potential human infectivity of hybrid progeny was tested by analysing resistance to lysis by human serum ; however, this is not such a reliable test for human infectivity as presence of the SRA gene.
SRA appears to be a single copy gene that resides in one of the telomeric expression sites (ES) for variant surface glycoprotein (VSG) genes, such that, when this ES is transcribed, SRA is also expressed . The ES containing SRA is unusually short in that it contains only three ES-associated genes (ESAGs 5, 6 and 7), with SRA located between ESAG 5 and the telomeric VSG gene . Both the SRA gene and its immediate genomic environment are conserved in different Tbr strains . The chromosome carrying SRA has not been identified, though from its size (1.6 Mb ), it appears to be one of the smaller diploid chromosomes described in T. brucei . It is also uncertain whether all Tbr strains carry only a single SRA allele or have multiple ES with SRA. It is technically difficult to sequence T. brucei ES because of their telomeric location , and the few studies to date show within-strain similarity of ES in structure and gene content [19–21], making it difficult to distinguish between different ES in the same trypanosome strain.
From an evolutionary perspective, it seems unlikely that Tbr would have only a single SRA gene, as that would make it dependent on only a single ES for infection in the human host; antigenic variation would be restricted to replacement of the VSG in this ES, and switching to expression of another ES, which lacked SRA, would be lethal for the parasite. Dependence on this one ES in the human host would lock the trypanosome into expression of the single transferrin receptor encoded by the ESAG 6 and ESAG 7genes co-transcribed with SRA . Moreover, according to the hypothesis that allelic variation in ESAG 6 and ESAG 7is adaptive for uptake of different mammalian transferrins [23,24], the receptor encoded by alleles in the SRA ES should be specific for human transferrin. A further problem confronts the trypanosome on transmission from tsetse to human, because metacyclics, the infective forms inoculated with the fly’s saliva, express a restricted set of VSGs residing in specialized ES lacking ESAGs  and presumably also SRA. Without protection of the SRA protein to inactivate the trypanolytic effect of human serum, how is it possible for Tbr metacyclics to survive the transition from fly to human?
Here we provide the definitive experimental proof that SRA is readily transferred between Tbr and Tbb during sexual reproduction, creating new genotypes of the human pathogen Tbr, because the SRA gene is now in a new genetic background consisting of an equal mixture of the parental Tbr and Tbb genomes. We show that SRA is present as a single copy on one homologue of chromosome IV in the majority of Tbr strains analysed and explore the implications for the epidemiology of HAT in East Africa.
Materials and Methods
Animal experiments were approved by the University of Bristol Ethical Review Group (Home Office licence PIL 30/1248) and carried out under the UK government Animals (Scientific Procedures) Act 1986.
Trypanosomes and cell culture
The following tsetse-transmissible strains of Trypanosoma brucei rhodesiense (Tbr) and T. b. brucei (Tbb) were used: Tbr 058 (MHOM/ZM/74/58 [26,27]); Tbr LUMP 1198 (MHOM/UG/76/LUMP 1198 [26,27]); Tbr TOR11(MHOM/UG/88/TOR11 ); Tbb J10 (MCRO/ZM/73/J10 CLONE 1 [26,27]); Tbb 1738 (MOVS/KE/70/EATRO 1738 [27,29]); Tbb 427 (MOVS/UG/60/427 VAR3 ). These strains represent a range of Tbr and Tbb genotypes from East Africa; isolate details are in S1 Table. Tbr 058 and all three Tbb strains have proved mating-competent in previous crosses. The Tbr and Tbb clones carried cytoplasmically-expressed genes for enhanced green fluorescent protein (GFP)  or monomeric red fluorescent protein (RFP) [32,33], respectively.
Procyclic form (PF) trypanosomes were grown in Cunningham’s medium (CM)  supplemented with 10% v/v heat-inactivated foetal calf serum, 5 μg/ml hemin and 10 μg/ml gentamycin at 27°C. PF were transfected by electroporation as previously described  and clones were obtained by limiting dilution of PF in CM in 96 well plates incubated at 27°C in 5% CO2.
Nine pairwise crosses were carried out, each involving one Tbr GFP clone and one Tbb RFP clone (crosses 1–9, Table 1), such that hybrids carrying both fluorescent markers appear yellow . Groups of 15–25 tsetse flies (Glossina morsitans morsitans or G. pallidipes) were infected on their first feed essentially as described previously [35,36]. The infective bloodmeal consisted of approximately 8 x 106 bloodstream form (BSF) trypanosomes ml-1 in sterile horse blood (TCS Biosciences, UK), or approximately 107 PF trypanosomes ml-1 of washed horse red blood cells resuspended in Hank’s Balanced Salt Solution, supplemented with 10mM L-glutathione . Infected flies were maintained on sterile horse blood until dissection approximately 5 weeks following the infective feed. Salivary glands (SG) were dissected in a drop of phosphate buffered saline and examined for the presence of fluorescent trypanosomes using a DMRB microscope (Leica) equipped with a Retiga Exi camera (QImaging) and Volocity software (PerkinElmer). SG containing an approximately equal mixture of trypanosome clones as judged by fluorescence were taken forward for isolation of hybrids (Fig. 1).
Experimental crosses were carried out by co-transmitting green fluorescent Tbr and red fluorescent Tbb through tsetse. Salivary glands with a mixed infection of red and green fluorescent trypanosomes were taken forward for analysis. BSF, bloodstream forms; PF, procyclic forms; HSR, human serum resistant.
Isolation and analysis of progeny clones
Metacyclics from infected SG were inoculated into mice (SCID or immunosuppressed MF1) and infected blood was harvested from the first peak of parasitaemia. Aliquots of approximately 107 BSF cells in whole blood were (a) transformed directly to PF by incubation in CM at 27°C, or (b) incubated in HMI-9 medium  with heat-inactivated human serum (WG serum donor) for 24 hours at 37°C to select human serum resistant (HSR) parasites, followed by inoculation into a mouse (SCID) and subsequent transformation of BSF from the first peak of parasitaemia into PF as in (a) above (Fig. 1). Clones were obtained from populations (a) unselected and (b) HSR by limiting dilution as above (Table 1), and grown in CM for purification of DNA using a spin column DNA purification kit. Microsatellite analysis was performed as described [33,36] for between four and six loci per clone, depending on the allelic differences between the parental clones used for the cross. The presence of the SRA gene was detected by PCR using primers SRA E (5’-TACTGTTGTTGTACCGCCGC) and SRA J (5’-GTACCTTGGCGCGCTCGCGCTG) followed by gel electrophoresis .
Samples for pulsed field gel (PFG) electrophoresis were prepared by lysing and deproteinising trypanosomes in situ in agarose blocks . PFG electrophoresis, blotting and hybridization were carried out essentially as described  using PCR-amplified DNA fragments as specific probes for genes encoding SRA and DNA topoisomerase (TOPO; chromosome IV).
Kinetoplast DNA maxicircle type was determined for selected clones as previously described .
Chromosomal location of SRA
Two approaches were used to identify the chromosomal location of SRA: (a) Quantitative PCR (qPCR) of SRA and chromosome-specific genes for chromosomes I-V (S2 Table). DNA was extracted from individual chromosome bands of Tbr 058 and LUMP 1198 after PFG chromosomal separation; gel bands were cut out and purified using GeneJet Gel Extraction Kit (Fermentas) according to manufacturer’s instructions for large chromosomes (>10kb DNA). All qPCRs were executed with 300nM primer concentrations (S2 Table) using a SYBR Green/ROX qPCR Master Mix (Fermentas) according to manufacturer’s instructions with 5 ng of template DNA per reaction; melting curve analysis was carried out to verify amplification of a single PCR product. Resulting data were analysed using MX Pro software (Agilent Technologies). (b) Sequential hybridisation of PFG blots with probes for various genes [β-tubulin (TUB), chromosome I; trypanothione synthetase (TS), chromosome II; paraflagellar rod protein (PFR1), chromosome III; DNA topoisomerase (TOPO), chromosome IV; lysosomal membrane protein (P67), chromosome V] was used to establish co-localisation with SRA. PFG samples were prepared from various Tbr isolates (S1 Table) and analysed as described above.
Copy number of SRA
SRA copy number relative to the housekeeping gene encoding triose phosphate isomerase (TIM) was determined in a range of Tbr samples (S1 Table); TIM is present in two copies on homologous chromosomes . QPCR was used to analyse the copy number of both genes and deduce the ratio of SRA to TIM, using SYBR-Green for detection and quantification of amplified DNA. QPCR conditions for amplification were optimized using a ten-fold dilution series of a plasmid construct containing one copy of each gene; after optimization, the nucleotide primers (S1 Fig) were used at 300nM SRA and 500nM TIM final concentration. All qPCR reactions were performed in triplicate and a positive control (with reference DNA) and a negative control (without DNA) were included in each set of reactions; qPCR reactions were run using a SYBR Green/ROX qPCR Master Mix (Fermentas) according to manufacturer’s instructions with 5 ng of template DNA per reaction; melting curve analysis was carried out to verify amplification of a single PCR product. Resulting data were analysed using MX Pro software (Agilent Technologies).
Inheritance of human infectivity
We set out to test whether genetic recombination between Tbr and Tbb enabled transfer of SRA into new genetic backgrounds and created potentially human infective hybrid genotypes. To detect hybrids we carried out pairwise crosses of three green fluorescent clones of Tbr with three red fluorescent strains of Tbb (crosses 1–9, Table 1), such that hybrids would appear yellow (Fig. 2) . Each of the three Tbr strains successfully mated with at least one of the Tbb strains, as judged by the production of hybrid clones; no hybrid progeny were recovered from crosses 5, 8 and 9 (Table 1). Clones were isolated either before (population a, unselected) or after incubation with human serum (population b, selected) (Fig. 1). The majority of clones (252 of 305, 83%) had the SRA gene whether derived from the selected or unselected populations (Table 1), demonstrating that SRA+ trypanosomes were not outcompeted by SRA- trypanosomes during development in the fly or growth as BSF in the mouse. A few SRA- clones survived incubation with human serum (12 of 119, 10%), but the majority of human serum resistant clones had SRA (107 of 119, 90%). Each clone was genotyped by microsatellite and molecular karyotype analysis, and also, where informative, kinetoplast maxicircle DNA type. Some genotypes were represented by more than one clone and found in both the human serum selected and unselected populations. Of the hybrid genotypes recovered, over half carried the SRA gene (Table 1), confirming that this gene can be transferred into different genetic backgrounds by sexual reproduction. We also confirmed presence of the SRA gene in hybrid clones from two previous crosses of Tbr 058 (crosses 10 and 11, Table 1).
A. Part of salivary gland containing T. b. rhodesiense 058 GFP and T. b. brucei 1738 RFP. Arrows: yellow fluorescent trypanosomes. Scale bar 100 μm. B. Part of salivary gland containing T. b. rhodesiense LUMP 1198 GFP and T. b. brucei J10 RFP; despite the presence of yellow fluorescent trypanosomes (arrows), no hybrid trypanosomes were recovered from this cross. Scale bar 50 μm.
Chromosomal location of SRA
In previous analysis of another Tbr strain, ETat 1, there appeared to be only one copy of the SRA gene, residing in an unusual truncated VSG expression site (ES) that contained only three ES associated genes (ESAGs) . In other Tbr isolates the local genomic environment of SRA was conserved , but there could be more than one copy of this ES and hence more than one copy of SRA. To investigate the chromosomal location of SRA, we purified DNA from individual chromosomal bands of Tbr 058 and LUMP 1198 and tested for the presence of various chromosome-specific genes (S2 Table) by qPCR. The Ct values for each gene tested are shown in Tables S3 and S4 and the results are shown graphically in Fig. 3. The lowest Ct value for Tbr 058 was for chromosomal band A5 corresponding to chromosomes IV and V, while that for Tbr LUMP 1198 was for chromosomal band B5 corresponding to chromosomes I–IV (Tables S3, S4 and Fig. 3). The combined results are consistent with the localisation of SRA to chromosome IV.
A. Chromosomal bands of T. b. rhodesiense 058 and LUMP 1198 separated by PFG; boxes outline the bands cut out and purified. B. Graphical summary of quantitative PCR results for individual chromosomal bands; the filled black boxes indicate the chromosome band with SRA. Ct values are given in Tables S3 and S4.
To confirm this result, we separated chromosome-sized DNA molecules of different Tbr strains by pulsed field gel electrophoresis (PFG) and hybridised with SRA (Fig. 4). Although the molecular karyotypes of the Tbr strains differed markedly in number and size of chromosomal bands, in each strain SRA located to one or two chromosomes of about 2 Mb in size (Fig. 4B); the fainter hybridisation signals result from weak hybridisation with SRA-related VSG genes and hence can be disregarded. Sequential hybridisation of identical blots with chromosome-specific probes revealed that SRA co-localized with the gene for DNA topoisomerase on chromosome IV (Fig. 4C). The location of SRA on one or both copies of chromosome IV was confirmed for most of the other Tbr isolates tested (Fig. 4D), with the exception of KETRI 2355 for which another (unidentified) chromosomal band hybridised with SRA (Fig. 4D). For LUMP 1198, SRA hybridized with the compression zone (cz), a region of the gel where DNAs from several large chromosomes co-migrate, as well as chromosome IV (Fig. 4D). However, our subsequent analysis of SRA copy number and inheritance in crosses of LUMP 1198 demonstrated the presence of only a single SRA gene (see below), so we assume that the cz signal derived from SRA–related VSG genes rather than SRA itself.
A. Ethidium bromide stained gel comparing the molecular karyotypes of clones from four strains of T. b. rhodesiense (Tbr). Size marker: chromosomal DNA from Hansenula wingei; mc = minichromosomes of 50–100 kb in size. B, C. Autoradiographs of blots of this PFG gel following hybridization with the probes indicated. Blots were washed to 0.1 × SSC at 65°C. D. Diagram of SRA and TOPO gene co-localisation. The first four samples are those shown in panels A-C, while the other six samples were run on other gels. TOR1 and TOR 4 gave identical results to TOR11, revealing that both chromosome IV homologues carry SRA in these three Tbr strains. LUMP 1198 and KETRI 2355 were the only Tbr examined that had a copy of SRA on a chromosome other than chromosome IV (black band); for KETRI 2355, SRA and TOPO did not co-localise.
Although the SRA gene and its immediate genomic environment have diverged in Tbr strains from northern and southern regions of East Africa [16,27], here SRA was located on chromosome IV in representative northern (TMRS 117) and southern (Gambella II, 058, EATRO 181) Tbr strains sequenced in the previous studies.
Copy number of SRA
The karyotype results suggest that Tbr strains generally have a single copy of SRA, or at most two copies. To verify this result, we estimated SRA copy number by quantitative PCR (qPCR) analysis, using copy number of the gene for triose phosphate isomerase (TIM) as the standard; in the diploid genome of T. brucei there are two copies of TIM . The relative rates of amplification of SRA and TIM [ratio d(SRAnorm-TIM)], were calculated for genomic DNA from sixteen different Tbr strains using 3 replicates for each strain (Fig. 5). Most strains, including LUMP 1198 and KETRI 2355, had a ratio of approximately 1:2 SRA:TIM, except for Tbr TOR11, which had a ratio of approximately 1:1. This agrees with the karyotype analysis above, where most Tbr strains had a single chromosomal band hybridizing with SRA, except TOR11, which had two.
Each bar shows the mean of 3 replicate experiments; bar is standard deviation. Dotted line at 1.0 indicates 1:2 ratio of SRA to TIM, while 1:1 ratio is at 0. Ct values for SRA were normalized (SRAn) using the standard curve obtained for the test plasmid containing one copy of each gene (S1 Fig), and then subtracted from TIM Ct values. The test plasmid was used as the 1:1 control.
The single, non-allelic copy of SRA in Tbr 058 and LUMP 1198 should segregate into 50% of hybrid progeny clones, assuming the rules of Mendelian inheritance are obeyed. Fig. 6 shows karyotype results for clones isolated from crosses of LUMP 1198 x 1738; the two chromosome IV homologues of Tbr LUMP 1198 co-migrate, but only one (red A) carries the SRA gene (Fig. 6B, lane 1). Three identical hybrid clones from cross 1198/1738-1 (lanes 2–4) lack SRA and are therefore assumed to have chromosome IV homologue A; these clones have also inherited the smaller chromosome IV homologue of 1738, B. Hybrid clones from cross 1198/1738-2 (lanes 6–13) demonstrate inheritance of parental chromosome IV homologues in all possible combinations (Fig. 6B, C).
A. Ethidium bromide stained gel comparing the molecular karyotypes of parental clones T. b. rhodesiense LUMP 1198 and T. b. brucei 1738 with those of 13 progeny clones; clones 1–5 and 6–13 are from two different tsetse flies, i.e. from two separate crosses. Size marker: chromosomal DNA from Hansenula wingei; mc = minichromosomes of 50–100 kb in size. B, C. Autoradiographs of blots of this PFG gel following hybridization with the probes indicated. Blots were washed to 0.1 × SSC at 65°C. Identification of chromosome IV homologues is shown below panel C: LUMP 1198 has homologues of the same size AA, while homologues of 1738 are designated B (lower) and C (upper); the red letter indicates the homologue carrying SRA. For hybrid clone 6, microsatellite analysis showed 3 alleles for a chromosome IV locus, indicating inheritance of both 1738 homologues.
In contrast, as SRA is located on both chromosome IV homologues in Tbr TOR11, diploid hybrid progeny from crosses with Tbb are expected to inherit a single copy of SRA. Results for crosses of TOR11 x J10 are shown in Fig. 7, where it can be seen that clones 3, 4 and 6 all have a single chromosome IV homologue carrying SRA from TOR11. However, all the other seven hybrid clones have both chromosome IV homologues with SRA from TOR11. Hybridisation intensities of individual chromosome bands suggest that these clones are trisomic for chromosome IV, with only one homologue from J10; this is obvious for clones 7 and 8, for which the chromosomal bands are well-separated (Fig. 7C); these clones also had three microsatellite alleles for the chromosome IV locus examined, confirming this result. Polyploid hybrids also occurred in the crosses involving Tbr 058 or LUMP 1198, explaining why a far greater proportion of hybrid clones than expected inherited SRA in these crosses (82%, 36 of 44 hybrid clones had SRA).
A. Ethidium bromide stained gel comparing the molecular karyotypes of parental clones T. b. rhodesiense TOR11 and T. b. brucei J10 with those of 10 progeny clones; clones 1–6, 7–9 and 10 are from three different tsetse flies, i.e. from three separate crosses. Size marker: chromosomal DNA from Hansenula wingei; mc = minichromosomes of 50–100 kb in size. B, C. Autoradiographs of blots of this PFG gel following hybridization with the probes indicated. Blots were washed to 0.1 × SSC at 65°C. Identification of chromosome IV homologues is shown below panel C: J10 homologues are designated A (lower) and B (upper), while TOR11 homologues are designated C (lower) and D (upper) and both carry SRA, denoted by red letters.
Our experimental crosses of Tbr and Tbb demonstrate unequivocally that the SRA virulence gene can be transferred by genetic exchange, thus creating new genotypes of potentially human infective parasites. The genetic heterogeneity of field isolates of Tbr from different regions of East Africa, together with their similarity to some Tbb isolates, first suggested that there might be hybridization between these two subspecies [42–44], and later studies have provided extensive evidence of genetic admixture [6,8].
Our crosses involved Tbr of the northern (LUMP 1198, TOR11) and southern (058) types , judged to differ in severity of HAT , and Tbb of different genotypic groups. Tbb J10 and 1738 belong to the kiboko/kakumbi group, distinguished from other East and West African Tbb such as Lister 427 by unusual isoenzymes, kinetoplast DNA maxicircle polymorphisms and microsatellite profiles [6,26,29]. Kiboko/kakumbi group isolates have never been found in human patients and originate from areas of East Africa that have a rich, large mammal fauna [46,47]. The tight association of kinetoplast and nuclear DNA polymorphisms suggested that the kiboko/kakumbi group circulates in separate wild animal-tsetse transmission cycles, without frequent sexual reproduction with other Tbb/Tbr strains. Contrary to this, we have shown that kiboko/kakumbi strains readily mate with different Tbr, as do other Tbb strains from both East and West Africa. Thus, there do not appear to be any intrinsic genetic barriers that prevent mating of Tbr and Tbb.
The accumulated data on location and copy number of SRA support the hypothesis that most Tbr strains have a single copy of SRA located in a VSG ES at the end of chromosome IV ([2,16] and this paper). As a consequence, SRA is only expressed when this ES is active, which means that the parasite is effectively restricted to use of this single ES in the human host. As noted above, a switch to another ES without SRA would be lethal for the trypanosome in a human host. This seems peculiar in a trypanosome that depends on antigenic variation for survival in the mammalian host and has multiple ES, especially considering that the SRA ES is truncated and lacks most ESAG’s . How can we explain this? One possibility is that there are fitness costs associated with expression of SRA in other non-human mammalian hosts, though there is currently no evidence for this. Tbr is a zoonotic pathogen that arguably depends on a large population of non-human hosts for longterm persistence in endemic areas. Hence, the ability to easily switch off a single copy of SRA by swapping to VSG expression from another ES might be advantageous. Although it has been suggested that there are fitness costs associated with resistance to human serum in Tbr in tsetse , this seems unlikely; bloodstream form ES are silenced during trypanosome development in the insect vector, with activation of another set of specialized ES lacking ESAG’s in the infective metacyclics in the salivary glands ; therefore SRA is probably not expressed in the fly.
Our results suggest a more plausible hypothesis based on the dynamic between Tbb and Tbr. SRA is a truncated VSG gene [50,51] and is assumed to have evolved once, since the sequence and local genomic environment of SRA is conserved among different Tbr strains [16,27]. We do not know when this event occurred, but SRA would only have become advantageous when it allowed extension of T. brucei’s host range to include hosts with the trypanolytic factor, Apolipoprotein L1 (APOL1), in their serum ; this probably dates the evolution of SRA, and hence Tbr, to somewhere in the last 10 million years or so, when the ape lineages with APOL1 diverged [53,54]. Although Tbr might subsequently have been subject to selective pressure for gene or ES duplication, depending on how significant the size of the host population with APOL1, any increase in copy number of SRA would have been rapidly diluted by mating with Tbb. Currently, there are likely to be more Tbb than Tbr strains circulating in East Africa, considering the relative numbers of infected human and non-human hosts and the restricted distribution of Tbr. Hence the probability of mating between Tbr strains will be far lower than between Tbb and Tbr, except possibly in the midst of an epidemic. This may explain the duplication of SRA in TOR11 and other isolates TOR1 and TOR4 from the same HAT outbreak (S1 Table). We can assume that these isolates represent one Tbr strain that arose either by hybridization between Tbr strains or as a mutated strain with duplication of the SRA ES.
Since Tbr typically has only a single copy of SRA in a bloodstream form ES, metacyclics presumably do not express SRA when inoculated into the human host and will therefore not be protected from lysis by APOL1. Indeed, we were unable to demonstrate expression of SRA by RT PCR of RNA prepared from tsetse salivary glands infected with Tbr 058. In vitro experiments comparing the resistance of Tbr and T. b. gambiense (Tbg1) to lysis by human serum showed that few Tbr metacyclics, but the majority of Tbg1 metacyclics, grew in medium containing human serum  and these authors hypothesized that survival of Tbr metacyclics in the human host depends on them being deposited in the skin tissue rather than bloodstream during tsetse bite, so that they are not directly exposed to the trypanolytic factor in the blood . In support of this hypothesis, the absence of APOL1 in human tissue fluid needs to be verified.
In conclusion, new human infective strains of the human pathogen Tbr can be generated by recombination of Tbr with the much larger pool of animal-infective trypanosomes, Tbb. Such novel recombinants present a risk for future outbreaks of HAT.
S1 Fig. Comparative efficiency of quantitative PCR (qPCR) for SRA and TIM.
S1 Table. Trypanosoma brucei rhodesiense isolates; all are from human hosts and SRA positive.
S2 Table. Primers used for quantitative PCR (qPCR).
S3 Table. Ct values for qPCR of individual chromosomal bands of T. b. rhodesiense 058.
S4 Table. Ct values for qPCR of individual chromosomal bands of T. b. rhodesiense LUMP 1198.
We are very grateful to the IAEA lab in Vienna for provision of tsetse and to Dr Chris Helps for advice on qPCR.
Conceived and designed the experiments: WG LP MB. Performed the experiments: WG LP VF KF JL JT MB. Analyzed the data: WG LP KF JL JT MB. Wrote the paper: WG KF.
- 1. Gibson W. Will the real Trypanosoma brucei rhodesiense please step forward? Trends Parasitol. 2002; 18: 486–490. pmid:12473364 doi: 10.1016/s1471-4922(02)02390-5
- 2. Xong VH, Vanhamme L, Chamekh M, Chimfwembe CE, Van den Abbeele J, Pays A, et al. A VSG expression site-associated gene confers resistance to human serum in Trypanosoma rhodesiense. Cell. 1998; 95: 839–846. pmid:9865701 doi: 10.1016/s0092-8674(00)81706-7
- 3. Radwanska M, Chamekh M, Vanhamme L, Claes F, Magez S, Magnus E, et al. The serum resistance-associated gene as a diagnostic tool for the detection of Trypanosoma brucei rhodesiense. Am J Trop Med Hyg. 2002; 67: 684–690. pmid:12518862
- 4. Njiru ZK, Mikosza ASJ, Armstrong T, Enyaru JC, Ndung'u JM, Thompson ARC. Loop-Mediated Isothermal Amplification (LAMP) Method for Rapid Detection of Trypanosoma brucei rhodesiense. PLoS NTD. 2008; 2: e147. doi: 10.1371/journal.pntd.0000147
- 5. Gibson W. Species-specific probes for the identification of the African tsetse-transmitted trypanosomes. Parasitology. 2009; 136: 1501–1507. doi: 10.1017/S0031182009006179. pmid:19490726
- 6. Balmer O, Beadell JS, Gibson W, Caccone A. Phylogeography and taxonomy of Trypanosoma brucei. PLoS NTD. 2011; 5: e961. doi: 10.1371/journal.pntd.0000961
- 7. Sistrom M, Evans B, Bjornson R, Gibson W, Balmer O, Mäser P, et al. Comparative genomics reveals multiple genetic backgrounds of human pathogenicity in the Trypanosoma brucei complex. Genome Biol Evol. 2014; 6: 2811–2819. doi: 10.1093/gbe/evu222. pmid:25287146
- 8. Goodhead I, Capewell P, Bailey JW, Beament T, Chance M, Kay S, et al. Whole-genome sequencing of Trypanosoma brucei reveals introgression between subspecies that is associated with virulence. MBio. 2013; 4: e00197–00213. doi: 10.1128/mBio.00197-13. pmid:23963174
- 9. Jenni L, Marti S, Schweizer J, Betschart B, Lepage RWF, Wells JM, et al. Hybrid formation between African trypanosomes during cyclical transmission. Nature. 1986; 322: 173–175. pmid:3724860 doi: 10.1038/322173a0
- 10. Peacock L, Ferris V, Sharma R, Sunter J, Bailey M, Carrington M, et al. Identification of the meiotic life cycle stage of Trypanosoma brucei in the tsetse fly. Proc Natl Acad Sci U S A. 2011; 108: 3671–3676. doi: 10.1073/pnas.1019423108. pmid:21321215
- 11. Peacock L, Bailey M, Carrington M, Gibson W. Meiosis and haploid gametes in the pathogen Trypanosoma brucei. Curr Biol. 2014; 24: 1–6. doi: 10.1016/j.cub.2013.11.017. pmid:24332542
- 12. Gibson WC. Analysis of a genetic cross between Trypanosoma brucei rhodesiense and T. b. brucei. Parasitology. 1989; 99: 391–402. pmid:2575239 doi: 10.1017/s0031182000059114
- 13. Gibson W, Garside L, Bailey M. Trisomy and chromosome size changes in hybrid trypanosomes from a genetic cross between Trypanosoma brucei rhodesiense and T. b. brucei. Mol Biochem Parasitol. 1992; 52: 189–200. doi: 10.1016/0166-6851(92)90069-v
- 14. Gibson W, Winters K, Mizen G, Kearns J, Bailey M. Intraclonal mating in Trypanosoma brucei is associated with out-crossing. Microbiology. 1997; 143: 909–920. pmid:9084175 doi: 10.1099/00221287-143-3-909
- 15. Gibson WC, Mizen VH. Heritability of the trait for human infectivity in genetic crosses of Trypanosoma brucei ssp. Trans Roy Soc Trop Med Hyg. 1997; 91: 236–237. pmid:9196780 doi: 10.1016/s0035-9203(97)90236-4
- 16. Gibson W, Ferris V. Conservation of the genomic location of the human serum resistance associated gene in Trypanosoma brucei rhodesiense. Mol Biochem Parasitol. 2003; 130: 159–162. pmid:12946855 doi: 10.1016/s0166-6851(03)00168-3
- 17. Turner CMR, Melville SE, Tait A. A proposal for karyotype nomenclature in Trypanosoma brucei. Parasitol Today. 1997; 13: 5–6. pmid:15275158 doi: 10.1016/s0169-4758(96)20056-0
- 18. Becker M, Aitcheson N, Byles E, Wickstead B, Louis E, Rudenko G. Isolation of the repertoire of VSG expression site containing telomeres of Trypanosoma brucei 427 using transformation-associated recombination in yeast. Genome Res. 2004; 14: 2319–2329. pmid:15520294 doi: 10.1101/gr.2955304
- 19. McCulloch R, Horn D. What has DNA sequencing revealed about the VSG expression sites of African trypanosomes? Trends Parasitol. 2009; 25: 359–363. doi: 10.1016/j.pt.2009.05.007. pmid:19632154
- 20. Hertz-Fowler C, Figueiredo LM, Quail MA, Becker M, Jackson A, Bason N, et al. Telomeric expression sites are highly conserved in Trypanosoma brucei. PLoS One. 2008; 3: e3527. doi: 10.1371/journal.pone.0003527. pmid:18953401
- 21. Young R, Taylor JE, Kurioka A, Becker M, Louis EJ, Rudenko G. Isolation and analysis of the genetic diversity of repertoires of VSG expression site containing telomeres from Trypanosoma brucei gambiense, T. b. brucei and T. equiperdum. BMC Genomics. 2008; 9: 385. doi: 10.1186/1471-2164-9-385. pmid:18700033
- 22. Steverding D, Stierhof YD, Chaudhri M, Ligtenberg M, Schell D, Becksickinger AG, et al. ESAG 6 and ESAG 7 products of Trypanosoma brucei form a transferrin-binding protein complex. Eu J Cell Biol. 1994; 64: 78–87.
- 23. Bitter W, Gerrits H, Kieft R, Borst P. The role of transferrin-receptor variation in the host range of Trypanosoma brucei. Nature. 1998; 391: 499–502. pmid:9461219 doi: 10.1038/35166
- 24. Gerrits H, Mussmann R, Bitter W, Kieft R, Borst P. The physiological significance of transferrin receptor variations in Trypanosoma brucei. Mol Biochem Parasitol. 2002; 119: 237–247. pmid:11814575 doi: 10.1016/s0166-6851(01)00417-0
- 25. Graham SV, Terry S, Barry JD. A structural and transcription pattern for variant surface glycoprotein gene expression sites used in metacyclic stage Trypanosoma brucei. Mol Biochem Parasitol. 1999; 103: 141–154. pmid:10551359 doi: 10.1016/s0166-6851(99)00128-0
- 26. Gibson WC, Marshall TFdC, Godfrey DG. Numerical analysis of enzyme polymorphism: a new approach to the epidemiology and taxonomy of trypanosomes of the subgenus Trypanozoon. Adv Parasitol. 1980; 18: 175–246. pmid:7001872 doi: 10.1016/s0065-308x(08)60400-5
- 27. Gibson W, Backhouse T, Griffiths A. The human serum resistance associated gene is ubiquitous and conserved in Trypanosoma brucei rhodesiense throughout East Africa. Infect Genet Evol. 2002; 1: 207–214. pmid:12798017 doi: 10.1016/s1567-1348(02)00028-x
- 28. Stevens JR, Lanham SM, Allingham R, Gashumba JK. A simplified method for identifying subspecies and strain groups in Trypanozoon by isoenzymes. Ann Trop Med Parasit. 1992; 86: 9–28. pmid:1616401
- 29. Gibson WC, Fase-Fowler F, Borst P. Further analysis of intraspecific variation in Trypanosoma brucei using restriction site polymorphisms in the maxi-circle of kinetoplast DNA. Mol Biochem Parasitol. 1985; 15: 21–36. pmid:2985985 doi: 10.1016/0166-6851(85)90026-x
- 30. Peacock L, Ferris V, Bailey M, Gibson W. Fly transmission and mating of Trypanosoma brucei brucei strain 427. Mol Biochem Parasitol. 2008; 160: 100–106. doi: 10.1016/j.molbiopara.2008.04.009. pmid:18524395
- 31. Bingle LEH, Eastlake JL, Bailey M, Gibson WC. A novel GFP approach for the analysis of genetic exchange in trypanosomes allowing the in situ detection of mating events. Microbiology. 2001; 147: 3231–3240. pmid:11739755
- 32. Campbell RE, Tour O, Palmer AE, Steinbach PA, Baird GS, Zacharias DA, et al. A monomeric red fluorescent protein. Proc Natl Acad Sci U S A. 2002; 99: 7877–7882. pmid:12060735 doi: 10.1073/pnas.082243699
- 33. Gibson W, Peacock L, Ferris V, Williams K, Bailey M. The use of yellow fluorescent hybrids to indicate mating in Trypanosoma brucei. Parasit Vector. 2008; 1: 4. doi: 10.1186/1756-3305-1-4
- 34. Cunningham I. New culture medium for maintenance of tsetse tissues and growth of trypanosomatids. J Protozoo. 1977; 24: 325–329. pmid:881656 doi: 10.1111/j.1550-7408.1977.tb00987.x
- 35. Peacock L, Ferris V, Bailey M, Gibson W. Mating compatibility in the parasitic protist Trypanosoma brucei. Parasit Vector. 2014; 7: 78. doi: 10.1186/1756-3305-7-78. pmid:24559099
- 36. Peacock L, Ferris V, Bailey M, Gibson W. Intraclonal mating occurs during tsetse transmission of Trypanosoma brucei. Parasit Vector. 2009; 2: 43. doi: 10.1186/1756-3305-2-43. pmid:19772562
- 37. Macleod ET, Maudlin I, Darby AC, Welburn SC. Antioxidants promote establishment of trypanosome infections in tsetse. Parasitology. 2007; 134: 827–831. pmid:17306056 doi: 10.1017/s0031182007002247
- 38. Hirumi H, Hirumi K. Continuous cultivation of Trypanosoma brucei bloodstream forms in a medium containing a low concentration of serum protein without feeder cell layers. J Parasitol. 1989; 75: 985–989. pmid:2614608 doi: 10.2307/3282883
- 39. MacLeod A, Tweedie A, McLellan S, Taylor S, Cooper A, Sweeney L, et al. Allelic segregation and independent assortment in Trypanosoma brucei crosses: Proof that the genetic system is Mendelian and involves meiosis. Mol Biochem Parasitol. 2005; 143: 12–19. pmid:15941603 doi: 10.1016/j.molbiopara.2005.08.013
- 40. Van der Ploeg LHT, Schwartz DC, Cantor CR, Borst P. Antigenic variation in Trypanosoma brucei analysed by electrophoretic separation of chromosome-sized DNA molecules. Cell. 1984; 37: 77–84. pmid:6202420 doi: 10.1016/0092-8674(84)90302-7
- 41. Gibson WC, Osinga KA, Michels PAM, Borst P. Trypanosomes of subgenus Trypanozoon are diploid for housekeeping genes. Mol Biochem Parasitol. 1985; 16: 231–242. pmid:3840571 doi: 10.1016/0166-6851(85)90066-0
- 42. Tait A, Barry JD, Wink R, Sanderson A, Crowe JS. Enzyme variation in Trypanosoma brucei spp. II. Evidence for T. b. rhodesiense being a set of variants of T. b. brucei. Parasitology. 1985; 90: 89–100. pmid:3856830 doi: 10.1017/s0031182000049040
- 43. Gibson WC, Wellde BT. Characterisation of Trypanozoon stocks from the South Nyanza sleeping sickness focus in Western Kenya. Trans Roy Soc Trop Med Hyg. 1985; 79: 671–676. pmid:4095748 doi: 10.1016/0035-9203(85)90187-7
- 44. Komba EK, Kibona SN, Ambwene AK, Stevens JR, Gibson WC. Genetic diversity among Trypanosoma brucei rhodesiense isolates from Tanzania. Parasitology. 1997; 115: 571–579. pmid:9488868 doi: 10.1017/s0031182097001856
- 45. MacLean L, Chisi JE, Odiit M, Gibson WC, Ferris V, Picozzi K, et al. Severity of Human African Trypanosomiasis in East Africa is associated with geographic location, parasite genotype and host-inflammatory cytokine response profile. Infection and Immunity. 2004; 72: 7040–7044. pmid:15557627 doi: 10.1128/iai.72.12.7040-7044.2004
- 46. Godfrey DG, Baker RD, Rickman LR, Mehlitz D. The distribution, relationships and identification of enzymic variants within the subgenus Trypanozoon. Adv Parasitol. 1990; 29: 1–74. pmid:2181826 doi: 10.1016/s0065-308x(08)60104-9
- 47. Stevens JR, Godfrey DG. Numerical taxonomy of Trypanozoon based on polymorphisms in a reduced range of enzymes. Parasitology. 1992; 104: 75–86. pmid:1614742 doi: 10.1017/s0031182000060820
- 48. Coleman PG, Welburn SC. Are fitness costs associated with resistance to human serum in Trypanosoma brucei rhodesiense? Trends Parasitol. 2004; 20: 311–315. pmid:15193561 doi: 10.1016/j.pt.2004.04.015
- 49. Barry JD, Graham SV, Fotheringham M, Graham VS, Kobryn K, Wymer B. VSG gene control and infectivity strategy of metacyclic stage Trypanosoma brucei. Mol Biochem Parasitol. 1998; 91: 93–105. pmid:9574928 doi: 10.1016/s0166-6851(97)00193-x
- 50. De Greef C, Hamers R. The serum resistance associated (SRA) gene of Trypanosoma brucei rhodesiense encodes a VSG-like protein. Mol Biochem Parasitol. 1994; 68: 277–284. pmid:7739673 doi: 10.1016/0166-6851(94)90172-4
- 51. Campillo N, Carrington M. The origin of the serum resistance associated (SRA) gene and a model of the structure of the SRA polypeptide from Trypanosoma brucei rhodesiense. Mol Biochem Parasitol. 2003; 127: 79–84. pmid:12615339 doi: 10.1016/s0166-6851(02)00306-7
- 52. Vanhamme L, Paturiaux-Hanocq F, Poelvoorde P, Nolan D, Lins L, Van den Abbeele J, et al. Apolipoprotein L-1 is the trypanosome lytic factor of human serum. Nature. 2003; 422: 83–87. pmid:12621437 doi: 10.1038/nature01461
- 53. Poelvoorde P, Vanhamme L, Van den Abbeele J, Switzer WM, Pays E. Distribution of apolipoprotein L-I and trypanosome lytic activity among primate sera. Mol Biochem Parasitol. 2004; 134: 155–157. pmid:14747153 doi: 10.1016/j.molbiopara.2003.11.006
- 54. Thomson R, Molina-Portela P, Mott H, Carrington M, Raper J. Hydrodynamic gene delivery of baboon trypanosome lytic factor eliminates both animal and human-infective African trypanosomes. Proc Natl Acad Sci U S A. 2009; 106: 19509–19514. doi: 10.1073/pnas.0905669106. pmid:19858474
- 55. Brun R, Jenni L. Human serum resistance of metacyclic forms of Trypanosoma brucei brucei, T. brucei rhodesiense and T. brucei gambiense. Parasitol Res. 1987; 73: 218–223. pmid:3588581 doi: 10.1007/bf00578507
- 56. Gibson W, Bailey M. Genetic exchange in Trypanosoma brucei: evidence for meiosis from analysis of a cross between drug resistant transformants. Mol Biochem Parasitol. 1994; 64: 241–252. pmid:7935602 doi: 10.1016/0166-6851(94)00017-4