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Veterinary Parasitology 181 (2011) 61– 68

Contents lists available at ScienceDirect

Veterinary Parasitology

jo u rn al hom epa ge : www.elsev ier .com/ locate /vetpar

rypanosome genetics: Populations, phenotypes and diversity

ndy Taita,b,∗, Liam J. Morrisona,b, Craig W. Duffya,b, Anneli Coopera,b,. Mike. R. Turnera,c,1, Annette Macleoda,b,1

Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, United KingdomHenry Wellcome Institute of Comparative Medical Sciences, Garscube, Glasgow G61 1QH, United KingdomGlasgow Biomedical Research Centre, 120, University Place, Glasgow G12 8TA, United Kingdom

r t i c l e i n f o

eywords:rypanosomeseneticsopulationsathogenesis

a b s t r a c t

In the last decade, there has been a wide range of studies using a series of molecularmarkers to investigate the genotypic diversity of some of the important species of Africantrypanosomes. Here, we review this work and provide an update of our current understand-ing of the mechanisms that generate this diversity based on population genetic analysis. Inparallel with field based studies, our knowledge of the key features of the system of geneticexchange in Trypanosoma brucei, based on laboratory analysis, has reached the point atwhich this system can be used as a tool to determine the genetic basis of a phenotype. Inthis context, we have outlined our current knowledge of the basis for phenotypic variationamong strains of trypanosomes, and highlight that this is a relatively under researched area,except for work on drug resistance. There is clear evidence for ‘strain’-specific variation intsetse transmission, a range of virulence/pathogenesis phenotypes and the ability to cross

the blood brain barrier. The potential for using genetic analysis to dissect these phenotypesis illustrated by the recent work defining a locus determining organomegaly for T. brucei.When these results are considered in relation to the body of research on the variability ofthe host response to infection, it is clear that there is a need to integrate the study of hostand parasite diversity in relation to understanding infection outcome.

. Introduction

Since the 1980s, we have known that the main species offrican trypanosomes infecting humans and livestock areenotypically diverse, with the exception of Trypanosomaivax, where the difficulties associated with growing thisarasite species in the laboratory has limited the number

f isolates available for analysis. The basis for this vari-tion has been controversial, with one view that geneticxchange between parasites was common, leading to the

∗ Corresponding author at: Institute of Infection, Immunity and Inflam-ation, College of Medical, Veterinary and Life Sciences, University oflasgow, Bearsden Rd., Glasgow G61 1QH, United Kingdom. Tel.: +44 014130 5750; fax: +44 0141 330 5422.

E-mail address: a.tait@vet.gla.ac.uk (A. Tait).1 Joint last authors.

304-4017/$ – see front matter © 2011 Elsevier B.V. All rights reserved.oi:10.1016/j.vetpar.2011.04.024

© 2011 Elsevier B.V. All rights reserved.

observed diversity (Tait, 1980; Gibson et al., 1980), whileanother view was that these parasites primarily expandedclonally with the observed variation being largely due tomutation (Tibayrenc et al., 1990). In the last few years, withthe advent of genome sequence data and the developmentof methods for the genetic characterisation (‘genotyping’)of parasites directly from blood samples, the questionregarding the role of genetic exchange in trypanosomepopulations has been further investigated. Particularlyimportant in these new studies has been the ability todevelop panels of highly polymorphic micro- and mini-satellite markers from genome sequences, so that parasitescan be rapidly genotyped by PCR based methods.

In parallel with these developments, our understandingof the mechanisms of genetic exchange, based on labora-tory crosses (reviewed by Gibson and Stevens, 1999), hasexpanded substantially, such that we are now in a position

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to use crosses and genetic analysis to determine the geneticbasis of phenotypes of relevance to the disease and itstransmission (Tait et al., 2002). Reverse genetic techniqueswith trypanosomes are well developed (Burkard et al.,2007; Ngo et al., 1998), at least for T. b. brucei, and allowstudies of the effect of gene knockouts or gene silencing byRNAi on phenotype. These are powerful technologies butare dependent on initially identifying and defining a gene(or genes) of interest and then determining the phenotypecaused by gene silencing or knockout. In contrast, forwardgenetic techniques, such as crosses, allow the investigationof the genetic basis of known phenotypes without priorknowledge of the genes involved. However, such studiesare dependent on the occurrence of naturally occurringvariation in phenotype for genetic analysis, highlightingthe need to study not only genotypic but also phenotypicvariation.

In this context, we review our current knowledge andunderstanding of the basis for genotypic diversity in fieldpopulations, the mechanism of genetic exchange and thevariation in phenotypes of relevance to the disease, coupledto recent parasite genetic analysis of one such phenotype.This review is mainly an account of recent findings on try-panosome genetics and diversity.

2. Trypanosome genetic diversity

T. brucei is found throughout the tsetse belt ofsub-Saharan Africa and comprises three morphologicallyidentical sub-species: T. b. rhodesiense, T. b. gambiense andT. b. brucei, with the first two causing significant diseasein humans and the third infecting livestock and wild ani-mals throughout the region (Fevre et al., 2008). The humaninfective sub-species occur in discrete foci of disease withT. b. gambiense found in West and Central Africa and T.b. rhodesiense in East and Southern Africa (Hoare, 1972).A combination of iso-enzyme and molecular markers hasshown that T. b. gambiense isolates can be divided into twodiscrete sub-groups, designated as Groups 1 and 2 (Gibson,1986). Although T. b. brucei is found in livestock, it is gen-erally not considered to be a major pathogen, in contrast toT. congolense and T. vivax which both cause major disease(Hoare, 1972). A combination of molecular and iso-enzymestudies have sub-divided T. congolense into three groupsor clades (Young and Godfrey, 1983; Majiwa et al., 1986;Gashumba et al., 1988), designated as Savannah, Forestand Kilifi, and one could speculate that these are differ-ent species or sub-species. In addition, a number of othertrypanosome species are important, but have not been thesubject of recent work on diversity and have thus not beenconsidered here.

In the last few years, molecular markers have beendeveloped to study the genotypic diversity of isolatesfrom most of the important species. While some analy-sis has previously been undertaken, the commonly usedapproaches/markers have been Amplified Fragment lengthPolymorphisms (AFLPs; Masiga et al., 2000; Agbo et

al., 2002; Masumu et al., 2006a; Simo et al., 2007),Mobile genetic element-Polymerase Chain Reaction (MGE-PCR; Hide and Tilley, 2001; Tilley et al., 2003) andmicro- and minisatellites (Biteau et al., 2000; MacLeod

logy 181 (2011) 61– 68

et al., 2000; Koffi et al., 2007). Essentially, the AFLPapproach analyses restriction fragment polymorphismson a genome-wide scale and therefore has a high sen-sitivity for detecting differences among strains. Howeverthis technique has a number of disadvantages; specif-ically, it requires growth of the parasites, preparationof purified DNA and is difficult to interpret in geneticterms. The MGE-PCR method overcomes the need to pre-pare DNA, but resultant electrophoretic patterns cannotbe readily interpreted genetically. By contrast, micro-and mini-satellites do not suffer from these disadvan-tages although, unless a very large number are used,they do not provide a genome-wide analysis. There isno ideal marker system, but methods should be selectedbased on the question being addressed and the natureof the material available for study. In the future, thesemethods/markers could be superseded by whole genomesequence analysis, as new technologies become availableand their costs reduce. However, this will still requirethe ‘amplification’ of parasites in rodents, potentially lim-iting their application as well as raising the possibilityof genotypic selection, which can occur during rodentamplification (McNamara et al., 1995; Jamonneau et al.,2003).

Detection of diversity, using these molecular methods,has been undertaken primarily for the three sub-species ofT. brucei but, more recently, for T. congolense (Savannah)and T. vivax. Within T. brucei, the general picture is that T.b. brucei is highly diverse, T. b. rhodesiense (depending onthe focus) ranges from showing very low levels of diversityto high levels, whereas T. b. gambiense shows low levelsof diversity within a focus, but strains from different geo-graphical foci are very distinct. These broad conclusionsare supported, irrespective of whether microsatellite, AFLPor MGE-PCR methods of analysis are used. Recent studiesof T. congolense (Savannah), using molecular markers andmultiple isolates, have shown high levels of diversity bothwithin a single geographical region (Morrison et al., 2009a)and between regions (Masumu et al., 2006a). However,limited information is available on the diversity betweendifferent geographically separated populations of T. con-golense (Savannah) or within the Kilifi and Forest ‘clades’of T. congolense. Thus, our knowledge of diversity in T. con-golense is relatively limited, apart from the iso-enzymestudies in the 1980s (Young and Godfrey, 1983; Gashumbaet al., 1988) but, where this species has been studied, itappears to be highly diverse. Because of the difficulty ofgrowing T. vivax in laboratory rodents or culture, knowl-edge of the strain diversity of this species has been limited(Gardiner, 1989; Fasogbon et al., 1990), except for a recentstudy using microsatellite markers amplified directly frominfected blood isolated from horses and cattle in a singlelocation (Duffy et al., 2009). The latter study showed thatthere was limited diversity as many isolates have identicalgenotypes. The findings from all the studies on diversityraise two main questions, what is the basis for the observeddiversity and why are differences in diversity found in dif-

ferent foci of the same sub-species as well as betweendifferent species. One central question is whether this vari-ation is generated by genetic exchange, and this aspect isaddressed in the next section.

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. Population genetics and molecular epidemiology

In the early 1990s, there was considerable debate abouthe role of genetic exchange in field populations of Africanrypanosomes. Tibayrenc et al. (1990) proposed that mostarasitic protozoa expand clonally with limited geneticxchange, while others (Tait, 1980; Gibson et al., 1980) took

different view, proposing that, in some populations of T.rucei, mating occurred regularly. These different hypothe-es were based on the analysis of the observed frequency ofifferent genotypes compared with those predicted if ran-om mating was occurring as well as the analysis of linkagequilibrium (LE; the predicted frequency of the combina-ions of alleles at pairs of loci, assuming random mating). Ashe authors were using different populations of isolates andsed different definitions of what constituted a population,here was no real resolution to this debate. Clonal popula-ions will show multiple isolates with the same genotype,igh levels of heterozygosity, departure from LE (LD; link-ge disequilibrium) and frequencies of genotypes at singleoci that do not agree with those predicted by random mat-ng. In contrast, a panmictic population will have few, ifny, identical genotypes, and the frequencies of genotypest single loci and alleles at pairs of loci will be in agree-ent with the proportions predicted for a random mating

opulation. Thus, the population structure can either belonal or panmictic. A third population structure was pro-osed by Maynard Smith et al. (1993), who were workingn bacterial genetics (of an epidemic population), in whichne or a few genotypes expand clonally due to a partic-lar selective advantage but mask an underlying randomating population. They re-analysed data from a T. bru-

ei population (from several hosts) in the Lambwe valleyKenya) and showed that this population had an epidemicopulation structure. This dataset partly resolved the dif-erent conclusions about the role of genetic exchange inhe field, as an epidemic population combines aspects oflonal expansion with underlying genetic exchange. Sub-equent analysis using micro and mini-satellite markersnd geographically discrete populations has led to a furtheresolution of the debate.

The T. brucei group of sub-species has been most inten-ively studied using microsatellite markers, with a focusn the human-infective sub-species. Studies of T. b. gambi-nse Group 1 (Morrison et al., 2008; Koffi et al., 2009) fromisease foci in West and Central Africa have shown thatopulations isolated from geographically discrete areasre clonal with multiple isolates having the same geno-ype, LD and genotype frequencies that do not conformo those predicted by random mating. An interesting find-ng is that the genotypes from geographically distinct foci

ere very distinct, giving measures of genetic distance thatre comparable with those between sub-species. The basisor this high level of divergence is not understood, butt could be postulated that, originally, there was a singleopulation throughout the region which has subsequentlyub-divided, with each focus evolving independently and

o diverging. This proposal would be consistent with evolu-ionary theories of clonal organisms (Balloux et al., 2003).hylogenetic studies could test this hypothesis but haveot yet been undertaken. Studies on T. b. rhodesiense and T.

logy 181 (2011) 61– 68 63

b. brucei in Uganda have provided evidence that the formerexpands clonally with limited genetic exchange, while thelatter has an epidemic population structure with geneticexchange playing a significant role in generating diversity(MacLeod et al., 2000). However, studies of a separate focusof human sleeping sickness in Malawi show that there isa much higher degree of diversity in these parasites thanthose from Uganda, and these data are suggestive of a rolefor genetic exchange in generating diversity in T. b. rhode-siense in Malawi (Duffy et al., unpublished data). Thus,the picture that emerges from these more recent stud-ies is that genetic exchange plays a limited role in T. b.gambiense populations, a variable role in T. b. rhodesiensepopulations and probably a significant role in T. b. brucei,although there are relatively few extensive studies of thelatter sub-species. Furthermore, these studies emphasizethe sub-structuring of these populations by showing sig-nificant genetic isolation between geographically separatepopulations.

Recent population analysis of T. vivax and T. con-golense from a single discrete area in The Gambia providesstrong evidence for different population structures in thetwo species. Genotyping of 84 isolates of T. congolense(Savannah) with seven microsatellite markers shows thatthey comprised 80 different multilocus genotypes (MLG),suggesting that genetic exchange might be occurring(Morrison et al., 2009a). However, when the data wereanalysed to test for random mating, there was evidencefor LD, and the genotype frequencies show a deficit ofheterozygotes. The population has none of the proper-ties of a clonal population. To investigate the reasons forthe observed LD, the population was tested for crypticsub-structuring and found to be sub divided into four sub-populations with one of these in LE and the others showingapproaches to LE. Thus, genetic exchange occurs in T. con-golense (Savannah). The reasons for this sub-structuring arenot clear, but are not associated with the species of host ortime of sampling.

A similar analysis was undertaken using isolates of T.vivax from the same set of samples (Duffy et al., 2009).A total of 31 isolates were genotyped with 8 microsatel-lite markers, and the combinations of alleles at each locusfor each sample were used to generate MLGs. Only 9 MLGswere defined, and one of these was the same in 15 of thesamples, suggesting a clonal population structure. A fur-ther test was conducted using standard population geneticanalysis, which showed an excess of heterozygotes, a highlevel of LD and a lack of agreement between the observedgenotype frequencies and those predicted for a randomlymating population, further confirming that this populationis clonal. For both species, it will be important to analyseadditional populations in other areas of Africa to establishwhether the findings from The Gambia apply generally, asit is possible that in different epidemiological situationsthe role of genetic exchange might vary. Therefore, under-standing the role of genetic exchange in field populationswill not only provide information on how genetic diversity

arises and on the consequent ability to adapt to selectivepressures, but will also increase our understanding of therole of mutation versus recombination in the evolution oftrypanosomes.

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4. Mechanisms of genetic exchange

Since 1986, it has been known that laboratory crossescan be undertaken between different strains of T. brucei (seeJenni et al., 1986), but it has only been relatively recentlythat the mechanism and other key features have beenelucidated. It is now clear that mating takes place in theepimastigote stages in the salivary glands of the tsetse fly(Tait et al., 2007; Gibson et al., 2008) and that there is strongevidence for the occurrence of meiosis (MacLeod et al.,2005a). Earlier studies had shown that mating was not anobligatory part of the life cycle (Schweizer et al., 1988)and this aspect has been confirmed more recently usingparental trypanosomes with fluorescent gene tags andgenetic marker analysis of the progeny (Gibson et al., 2008).To date, some 12 crosses between different strains andsub-species have been undertaken, with genetic exchangebeing demonstrated between distinct strains of T. b. brucei,T. b. brucei and Group 2 T. b. gambiense, T. b. rhodesiense andT. b. brucei and T. b. rhodesiense and Group 2 T. b. gambiense(see Gibson and Stevens, 1999; MacLeod et al., 2007). Thus,there appear to be limited barriers to mating, even betweensub-species under laboratory conditions, although this hasnot been tested with T. b. gambiense Type 1. However, todate, there is no evidence from the field to suggest thatmating between the sub-species occurs (MacLeod et al.,2000), but this issue has not been systematically studied.As well as cross fertilisation between different parasitestrains, self-fertilisation also occurs and, although this pro-cess was originally thought to only occur in the presenceof cross fertilisation (Tait et al., 1996), recent data haveshown that self-fertilisation can occur when a single strainis transmitted through tsetse flies (Peacock et al., 2009). Thenon-obligatory mating system and the ability to undergoself fertilisation suggests that trypanosomes are uniquelyadaptable to environmental change and new niches; asthey can propagate clonally without disrupting combina-tions of alleles, they can generate novel combinations ofalleles by mating or become homozygous for ‘successful’alleles by self fertilisation.

Earlier studies had provided evidence that was consis-tent with the progeny of crosses being the equivalent ofan F1 produced by the parental strains undergoing meio-sis (Turner et al., 1990; Gibson, 1995). More recently, theseconclusions have been rigorously tested by microsatellitemarker analysis of >30 progeny clones from two indepen-dent crosses. Alleles at heterozygous loci on one parentalstrain, each located on one of the 11 housekeeping chro-mosomes, segregate into the progeny independently andin Mendelian proportions, thus strongly supporting theoccurrence of meiosis (MacLeod et al., 2005a). All of theprogeny analysed to date appear to be the products ofa single round of mating. Furthermore, genetic maps ofboth T. b. brucei and T. b. gambiense Group 2 have beenconstructed using > 150 microsatellite markers and theresultant linkage groups align with the physical map ofthe genome provided by the T. b. brucei genome sequence

(MacLeod et al., 2005b; Cooper et al., 2008). The con-struction and analysis of the genetic maps illustrated twofurther standard properties of the genetic system, namelycrossing-over between pairs of homologous chromosomes

logy 181 (2011) 61– 68

and regions of high and low recombination (hot and coldspots) along a chromosome. Thus, overall, T. brucei appearsto have a conventional diploid, Mendelian genetic systemin common with many other eukaryotes. It should be notedthat no crosses have been reported with T. b. gambienseGroup 1, and, to our knowledge, no crosses have beenreported with either T. congolense or T. vivax. The recentreport of the ability to replicate the whole life cycle of T.congolense in vitro (Coustou et al., 2010) offers the oppor-tunity of testing whether crosses with this species could beundertaken.

While many progeny clones in T. brucei crosses arediploid, a proportion of these are triploid or even tetraploidwith the frequency varying between different crosses(Gibson and Stevens, 1999). For example, in the crossesbetween two T. b. brucei strains (TREU 927 and STIB247), no triploids have ever been observed, but in a crossbetween T. b. gambiense and T. b. rhodesiense, 6/12 progenywere triploid (Gibson and Bailey, 1994). Originally, it wasthought that this phenomenon only occurred in inter sub-species crosses, but, recently, both tetraploid and triploidprogeny were reported in a purely T. b. brucei cross (Gibsonet al., 2008). While this can be explained by some cells notundergoing meiosis, but still being stimulated to fuse, it isunclear why it occurs at quite a high frequency in somecrosses. No triploid T. brucei have been described in themarker analysis of field isolates and, so, it appears that thisdoes not happen in natural populations, although it couldbe that polyploidy trypanosomes are not viable in theirnatural hosts. However, it should be pointed out that sim-ilar deviations in ploidy are found in human conceptusesbut result in miscarriage. Thus, this may be a general phe-nomenon but is only readily seen in trypanosomes, becauseof their tolerance of changes in ploidy. This phenomenonrequires further investigation.

5. Phenotypic diversity

The most extensively studied trypanosome phenotypehas been drug resistance due to its importance for thecontrol and treatment of both animal and human try-panosomosis. Variation in drug response to most of thecommonly used trypanocidal drugs has been reportedbetween different strains of the T. brucei group of sub-species and T. congolense. This diversity has been reviewedelsewhere (Matovu et al., 2001; Delespaux and de Koning,2007), and most of the analysis of mechanisms has beenundertaken using biochemical and molecular approachesrather than genetic analysis. Less attention has beenfocussed on strain variation in other phenotypes, althoughthere are reports of differences in virulence (Masumuet al., 2006b; Holzmuller et al., 2008; Balmer et al., 2009),pathogenicity (Morrison et al., 2009b) and tsetse transmis-sibility (Welburn et al., 1995). It is important to identifythe parasite genes that determine these traits and, fromthis begin to understand, the molecular determinants ofthe disease and its transmission.

There have been relatively few reports on parasite strainvariation in relation to the ability to infect and developin the tsetse fly, yet, this could be a significant factor indetermining which strains are likely to spread rapidly. A

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omparison between the transmission (by Glossina m. mor-itans) of a set of T. b. rhodesiense and T. b. brucei isolatesWelburn et al., 1995) has shown that there were significantifferences in the transmission index (TI = the ratio of midut to salivary gland infections). The results showed a cleartatistically significant difference in the mean TI betweenhe two sub-species with T. b. rhodesiense, giving lower val-es, but, interestingly, there was also variation betweentrains within each sub-species although these differencesere smaller. As far as we are aware, there has not been fur-

her research on the basis for these differences. A similarnding of strain variation in tsetse transmission has been

ound for T. congolense (see Masumu et al., 2006c), althoughrimarily in terms of mid-gut infectivity.

Clearly, the outcome of an infection will be determinedy both host and parasite factors, and the former haveeen studied extensively in many host–pathogen systems.his is also true for trypanosome infections, with stud-es implicating a range of host parameters that determinehe outcome of infection (MacLean et al., 2004, 2007; Hillt al., 2005; Courtin et al., 2008). However, the focus ofhis review is on the parasite determinants and, so, theost component will not be considered in any detail. This

s a relatively neglected area, despite its potential impor-ance. There are several studies that implicate variationn the parasite as the determinant of clinical outcomeJamonneau et al., 2000; Garcia et al., 2006), but as botharasite and host vary in such field studies, there is uncer-ainty as to the cause of the variation. Thus, much of thevailable data comes from studies using inbred mice ashe host, making the assumption that the findings applyo the natural host. Examples from T. brucei include thetudy of two different strains of T. b. brucei showing sig-ificant differences in organomegaly, reticulocytosis andnaemia (Morrison et al., 2009b). The basis for this vari-tion was investigated by analysing the response of theost to the infection with the two strains using microarraynalysis of RNA from the spleens. The main pathways thatere differentially regulated involved the innate immune

ystem (IL10 signalling, LXR/RXR signalling and alterna-ive macrophage activation), suggesting that strain specific

odulation of these pathways is a key component of theathogenesis. Differences in virulence between anotherair of T. b. brucei strains have also been observed (Balmert al., 2009) with significant differences found in hosturvival, thrombocytopenia, anaemia and hypoglycaemiaeasured in outbred mice. Interestingly, the less virulent

ine became virulent after multiple passage in mice andixed infections with virulent and avirulent strains lead to

he suppression of the virulent phenotype. In another studyHolzmuller et al., 2008), 10 strains of T. b. gambiense fromote d’Ivoire demonstrated variation in virulence (definedy level of parasitaemia) and pathogenicity (defined by sur-ival). Variation in virulence/pathogenicity has also beenescribed in studies of the three different clades (Savan-ah, Kilifi and Forest) of T. congolense in both bovine andurine hosts, in which, importantly, the infection param-

ters for both hosts showed a good concordance (Bengalyt al., 2002a,b). Additionally, a further study of 31 strains of. congolense (Savannah) from several locations in Zambiahowed a range of infection parameters, with the strains

logy 181 (2011) 61– 68 65

being classified into three groups: virulent, moderatelyvirulent and low virulence (Masumu et al., 2006b). Theparameters that differed between the groups were sur-vival time, pre-patent period and packed cell volume (PCV).Thus, there was both within and between clade diversity.

A key feature of T. brucei is the ability to penetratethe blood brain barrier, leading to one of the major clin-ical signs of the human disease, and field studies havereported (Jamonneau et al., 2004) infected patients who donot progress to stage 2-disease (i.e. penetration of the bloodbrain barrier). An in vitro system has been developed tomeasure the penetration of brain microvascular endothe-lial cells by T. brucei to study this process (Grab et al.,2004), and inhibitor studies have implicated a parasitesecreted cysteine proteinase (brucipain) as the moleculethat mediates penetration (Nikolskaia et al., 2006). Fromthe perspective of parasite diversity, experiments wereundertaken with T. b. rhodesiense (previously described as T.b. gambiense, in error) and two strains of T. b. brucei to showthat the human-infective sub-species crosses the endothe-lial cells six times as effectively as the T. b. brucei strains.Interestingly, T. b. rhodesiense has some eight-fold higherbrucipain activity, raising the possibility that this activitymay be responsible for the increased ability of T. b. rhode-siense to penetrate the endothelial monolayer (Nikolskaiaet al., 2006).

Taken together, there is clear evidence for parasitestrain/sub-species variation in a range of phenotypes thataffect transmission, disease severity and disease treatment.Apart from considerable advances in our understanding ofdrug resistance (disease treatment), we have very limitedknowledge of the genetic basis for the other phenotypes;yet they are likely to be important in terms of understand-ing the disease and its spread. Furthermore, identifying theparasite genes involved could, potentially have therapeuticimplications.

6. Genetic analysis of phenotypic variation

There is a considerable body of data on the variationin the host response to infection and the genetic basis oftolerance or ‘resistance’ to infection both in human andlivestock disease (Kemp et al., 1997). In terms of human dis-ease, a number of association studies have been undertakenand evidence for specific host genetic variants obtained(Courtin et al., 2008). In the case of animal trypanosomo-sis (using T. congolense), genetic analyses of both cattleand mice have been undertaken to identify loci that aremajor determinants of disease outcome (Iraqi et al., 2000;Hanotte et al., 2003; Hill et al., 2005). With the availabil-ity of single nucleotide polymorphism (SNP) arrays forhumans, cattle and mice, the resolution of such mappingstudies will be increased by using whole genome scans asan approach and it is likely that further the host gene vari-ants responsible for disease outcome could be identified.

The availability of genetic maps of T. brucei has openedup the possibility of undertaking analyses to identify the

loci that determine phenotypes of relevance to diseasetransmission and pathogenesis/virulence. One study (fromour laboratory) has been undertaken to map the locithat determine strain specific differences in organomegaly,

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reticulocytosis and anaemia (Morrison et al., 2010). Theprogeny from a cross between two strains that differedin these phenotypes were scored (in mice) for each ofthe parameters, and they were shown to segregate in asemi-quantitative manner. This information implied thatthese phenotypes are determined by allelic variation ofseveral parasite genes and, so, a genetic linkage analy-sis was undertaken assuming that these were quantitativetrait loci (QTL). Splenomegaly and hepatomegaly showedevidence for a highly significant QTL (LOD scores > 7) onchromosome 3, accounting for 66% and 64%, respectivelyof the phenotypic variance, thus demonstrating a par-asite gene, or genes, determining a major componentof these phenotypes. The identified region on chromo-some 3 included >300 genes; therefore, further analysisis required to identify the individual genes involved. Sev-eral approaches are available including, fine-scale mappingwith more markers, analysis of more progeny clones toidentify crossovers within the designated region, analysisof stage-specific expression and, possibly, reverse genetics.While this locus was the most significant, additional signif-icant QTLs were identified for reticulocytosis, anaemia andorganomegaly on other chromosomes. Similar approachesto those described are needed to fully confirm the signifi-cance of these loci. The importance of this work is two fold.Firstly it establishes unequivocally that there are parasiteloci determining pathogenesis and, secondly, that geneticanalysis can be used to map such loci in trypanosomes. Suchfindings need to be considered in studies of the host geneticdeterminants, for which much of the work is based on usinga single parasite ‘strain’.

7. Conclusions and future prospects

The genotypic diversity within some of the majorspecies and sub-species of trypanosomes varies substan-tially, ranging from low, in the case of T. gambiense Group1 and T. vivax, to high, in the case of T. b. brucei and T. con-golense. This variation reflects the role of genetic exchangein the populations studied. The recent sequencing of manyprotist genomes has led to the analysis of genes that areassociated with meiosis and an argument that the pres-ence of such genes indicates the existence of a sexual cycle– the simple ‘lose it or use it’ premise (Schurko and Logsdon,2008). The meiosis-associated genes (‘meiotic tool box’) arepresent in all the trypanosome species and sub-species dis-cussed here, but, clearly, the presence of these genes doesnot necessarily imply they are functional in the species withclonal population structures. Possible explanations for thisapparent contradiction could be that (i) these species doundergo genetic exchange in certain epidemiological sce-narios, (ii) these genes have alternative functions, or (iii)there has not been sufficient time for mutations and dele-tions to occur since the species started to expand clonally.

In parallel with the population-based analyses, ourunderstanding of the process of genetic exchange, at leastfor T. brucei, has advanced substantially to a point that

genetic maps of the parasite have been constructed so thatgenetic analysis can be undertaken. There are still somesignificant unknowns, such as whether haploid gametesoccur, the basis for the high proportion of non-diploid

logy 181 (2011) 61– 68

progeny, the nature of the process that leads to cell fusionfor the formation of progeny and the nature of the signalsthat initiate meiosis. The population-based identificationof mating in T. congolense (Savannah) also raises a numberof questions, such as whether this can be reproduced in thelaboratory, whether the genetic system has a similar mech-anism to that in T. brucei and whether the other clades of T.congolense also undergo genetic exchange.

The basis for the variation in phenotypes, such astsetse transmission, virulence and pathogenesis, is not wellunderstood and, except for the penetration of the bloodbrain barrier, the genes involved largely remain uniden-tified. However, in terms of determining how disease iscaused and the parasite is transmitted, it would be impor-tant to identify the key parasite mechanisms. Furthermore,knowledge of the parasite genes involved could provideavenues for new intervention and treatment strategies.The advances in trypanosome laboratory genetics offer anapproach to identify such genes and offer promise for thefuture. An alternative approach would be to undertakeassociation analyses between genotype and phenotype. Forexample, if large numbers of virulent and avirulent strainswere sequenced, polymorphisms that specifically associ-ated with one phenotype or the other would allow theidentification of candidate genes. Such approaches havebeen successfully undertaken to identify specific genesresponsible for non-infectious diseases in, for example, thedog (Karlsson et al., 2007) and cattle (Charlier et al., 2008).The advances in sequencing technologies would allow thegenotyping/sequencing of large numbers of strains to beundertaken cheaply and rapidly and, so, this is a prospectfor the future. Whether such analyses would be mosteffectively undertaken with populations of clonal parasitespecies or would require the use of panmictic populationsis a matter for discussion. The demonstration of variationin virulence phenotypes in T. b. gambiense (see Holzmulleret al., 2008), for example, for which genotypic diversity islow, could allow rapid association with genotypic polymor-phism but might suffer from a lack of resolution, as thehaplotypes associated with the phenotype could be largedue to the lack of recombination. In contrast, an analysisof T. congolense (with high levels of polymorphism) wouldreduce the size of the haplotypes but increase the levels of‘polymorphic noise’, making any association more difficultto establish statistically.

The studies of disease progression and pathogenesissuggest that parasite diversity plays a significant role inthe outcome of infection and will need to be consideredin relation to host diversity. We now have the tools andtechnologies to define the host and parasite loci that areinvolved and to identify the alleles that are responsiblefor different phenotypes. Thus, it is possible to envisage ahost–parasite ‘interactome’ that would begin to define themultiple routes to symptomatic and asymptomatic infec-tion. Similar approaches could also be taken to understandtsetse transmission, in order to define high and low trans-mission scenarios.

Conflict of interest

The authors declare that there is no conflict of interest.

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cknowledgements

AT, CMR and AM would like to acknowledge the sup-ort of the Wellcome Trust for the work in their laboratory.M is a Wellcome Trust Career Development Fellow, LJM

s a Royal Society University Research Fellow and CWD isupported by the Wellcome Trust 4 year PhD programmewarded to Glasgow University.

eferences

gbo, E.E.C., Majiwa, P.A.O., Claassen, H.J.H.M., Tepas, M.F.W., 2002. Molec-ular variation of Trypanosoma brucei subspecies as revealed by AFLPfingerprinting. Parasitology 124, 249–358.

alloux, F., Lehmann, L., de Meeus, T., 2003. The population genetics ofclonal and partially clonal diploids. Genetics 164, 1635–1644.

almer, O., Stearns, S.C., Schotzau, A., Brun, R., 2009. Intraspecific compe-tition between co-infecting parasite strains enhances host survival inAfrican trypanosomes. Ecology 90, 3367–3378.

engaly, Z., Sidibe, I., Boly, H., Sawadogo, L., Desquesnes, M., 2002a.Comparative pathogenicity of three genetically distinct Trypanosomacongolense-types in inbred Balb/c mice. Vet. Parasitol. 105, 111–118.

engaly, Z., Sidibe, I., Ganaba, R., Desquesnes, M., Boly, H., Sawadogo, L.,2002b. Comparative pathogenicity of three genetically distinct typesof Trypanosoma congolense in cattle: clinical observations and haema-tological changes. Vet. Parasitol. 108, 1–19.

iteau, N., Bringaud, F., Gibson, W., Truc, P., Baltz, T., 2000. Characteri-zation of Trypanozoon isolates using a repeated coding sequence andmicrosatellite markers. Mol. Biochem. Parasitol. 105, 187–202.

urkard, G., Fragoso, C.M., Roditi, I., 2007. Highly efficient stable transfor-mation of bloodstream forms of Trypanosoma brucei. Mol. Biochem.Parasitol. 153, 220–223.

harlier, C., Coppieters, W., Rollin, F., Desmecht, D., Agerholm, J., Cam-bisano, S., Carta, N., Dardano, E., Dive, S., Fasquelle, M., Frennet, C.J.,Hanset, C., Hubin, R., Jorgensen, X., Karim, C., Kent, L., Harvey, M.,Pearce, K.B., Simon, R., Tama, P., Nie, N., Vandeputte, H., Lien, S.,Longeri, S., Fredholm, M., Harvey, M.R., Georges, J.M., 2008. Highlyeffective SNP-based association mapping and management of reces-sive defects in livestock. Nat. Genet. 40, 449–454.

ooper, A., Tait, A., Sweeney, L., Tweedie, A., Morrison, L., Turner, C.M.R.,MacLeod, A., 2008. Genetic analysis of the human infective try-panosome Trypanosoma brucei gambiense: chromosomal segregation,crossing over, and the construction of a genetic map. Genome Biol. 9,R103.

ourtin, D., Berthier, D., Thevenon, S., Dayo, G.-K., Garcia, A., Bucheton, B.,2008. Host genetics in African trypanosomiasis. Infect. Gen. Evol. 8,229–238.

oustou, V., Guegan, F., Plazolles, N., Baltz, T., 2010. Complete in vitro lifecycle of Trypanosoma congolense: development of genetic tools. PLoSNegl. Trop. Dis. 4, e618.

elespaux, V., de Koning, H.P., 2007. Drugs and drug resistance in Africantrypanosomiasis. Drug Res. Updates 10, 30–50.

uffy, C.W., Morrison, L.J., Black, A., Pinchbeck, G.L., Christley, R.M.,Schoenefeld, A., Tait, A., Turner, C.M.R., MacLeod, A., 2009. Try-panosoma vivax displays a clonal population structure. Int. J. Parasitol.39, 1475–1483.

asogbon, A.I., Knowles, G., Gardiner, P.R., 1990. A comparison of the isoen-zymes of Trypanosoma (duttonella) vivax isolates from East and WestAfrica. Int. J. Parasitol. 20, 389–394.

evre, E.M., Wissmann, B.V., Welburn, S.C., Lutumba, P., 2008. The burdenof Human African Trypanosomiasis. PLoS Negl. Trop. Dis. 2, e333.

arcia, A., Courtin, D., Solano, P., Koffi, M., Jamonneau, V., 2006. HumanAfrican trypanosomiasis: connecting parasite and host genetics.Trends Parasitol. 22, 405–409.

ardiner, P.R., 1989. Recent studies of the biology of Trypanosoma vivax.Adv. Parasitol. 28, 229–317.

ashumba, J.K., Baker, R.D., Godfrey, D.G., 1988. Trypanosoma congolense:the distribution of enzymic variants in East and West Africa. Parasitol-ogy 96, 475–486.

ibson, W.C., Marshall, T.F., de, C., Godfrey, D.G., 1980. Numerical anal-

ysis of enzyme polymorphism: a new approach to the epidemiologyand taxonomy of trypanosomes of the subgenus Trypanozoon. Adv.Parasitol. 18, 175–246.

ibson, W.C., 1986. Will the real Trypansoma b. gambiense please stand up.Parasitol. Today 2, 255–257.

logy 181 (2011) 61– 68 67

Gibson, W.C., Bailey, M., 1994. Genetic exchange in Trypanosoma brucei:evidence for meiosis from analysis of a cross between drug-resistanttransformants. Mol. Biochem. Parasitol. 64, 241–252.

Gibson, W.C., 1995. The significance of genetic exchange in trypanosmes.Parasitol. Today 11, 465–468.

Gibson, W.C., Stevens, J., 1999. Genetic exchange in the Trypanosomatidae.Adv. Parasitol. 43, 1–45.

Gibson, W., Peacock, L., Ferris, V., Williams, K., Bailey, M., 2008. The use ofyellow fluorescent hybrids to indicate mating in Trypanosoma brucei.Parasites Vectors 1, 4.

Grab, D.J., Nikolskaia, O., Kim, Y.V., Lonsdale-Eccles, J.D., Ito, S., Hara, T.,Fukuma, T., Nykrko, E., Jim, K.J., Stins, M.F., Delannoy, M.J., Kim, K.S.,2004. African trypanosome interactions with an in vitro model of thehuman blood–brain barrier. J. Parasitol. 90, 970–979.

Hanotte, O., Ronin, Y., Agaba, M., Nilsson, P., Gelhaus, A., horstmann, R.,Sugimoto, Y., Kemp, S., Gibson, J., Korol, A., Soller, M., Teale, A., 2003.Mapping of quantitative trait loci controlling trypanotolerance in across of tolerant West African N’Dama and susceptible East AfricanBoran cattle. Proc. Natl. Acad. Sci. U.S.A. 100, 7443–7448.

Hide, G., Tilley, A., 2001. Use of mobile genetic elements as tools for molec-ular epidemiology. Int. J. Parasitol. 31, 599–602.

Hill, E.W., O’Gorman, G.M., Agaba, M., Gibson, J.P., Hanotte, O., Kemp,S.J., Naessens, J., Coussens, P.M., Machugh, D.E., 2005. Understandingbovine trypanosomiasis and trypanotolerance: the promise of func-tional genomics. Vet. Immunol. Immunopathol. 105, 247–258.

Hoare, C.A., 1972. The Trypanosomes of Mammals A Zoological Mono-graph. Blackwell Scientific Publications, Oxford.

Holzmuller, P., Biron, D.G., Courtois, P., Koffi, M., Bras-Goncalves, R., deDauloue, S., Solano, P., Cuny, G., Vincendeau, P., Jamonneau, V., 2008.Virulence and pathogenicity patterns of Trypanosoma brucei gambi-ense field isolates in experimentally infected mouse: differences inhost immune response modulation by secretome and proteomics.Microbes Infect. 10, 79–86.

Iraqi, F., Clapcott, S.J., Kumari, P., Haley, C.S., Kemp, S.J., Teale, A.J., 2000.Fine mapping of trypanosomiasis resistance loci in murine advancedintercross lines. Mamm. Genome 11, 645–648.

Jamonneau, V., Garcia, A., Frezil, J.L., N’guessan, P., N’dri, L., Sanon, R.,Laveissiere, C., Truc, P., 2000. Clinical and biological evolution ofhuman trypanosomiasis in Cote d’Ivoire. Ann. Trop. Med. Parasitol.94, 831–835.

Jamonneau, V., Barnabe, C., Koffi, M., Sane, B., Cuny, G., Solano, P., 2003.Identification of Trypanosoma brucei circulating in a sleeping sick-ness focus in Cote d’Ivoire: assessment of genotype selection by theisolation method. Infect. Genet. Evol. 3, 143–149.

Jamonneau, V., Ravel, S., Garcia, A., Koffi, M., Truc, P., Laveissiere, C.,Herder, S., Grebaut, P., Cuny, G., Solano, P., 2004. Characterizationof Trypanosoma brucei s.l. infecting asymptomatic sleeping-sicknesspatients in Cote d’Ivoire: a new genetic group? Ann. Trop. Med. Para-sitol. 98, 329–337.

Jenni, L., Marti, S., Schweizer, J., Betschart, B., LePage, R.W.F., Wells, J.M.,Tait, A., Pays, E., Paindavoine, P., Steinert, M., 1986. Hybrid formationbetween African trypanosomes during cyclical transmission. Nature322, 173–175.

Karlsson, E.K., Baranowska, I., Wade, C.M., Salmon Hillbertz, N.H., Zody,M.C., Anderson, N., Biagi, T.M., Patterson, N., Pielberg, G.R., Kulbokas,E.J., 3rd, Comstock, K.E., Keller, E.T., Mesirov, J.P., von Euler, H., Kampe,O., Hedhammar, A., Lander, E.S., Andersson, G., Andersson, L., Lindblad-Toh, K., 2007. Efficient mapping of Mendelian traits in dogs throughgenome-wide association. Nat. Genet. 39, 1321–1328.

Kemp, S.J., Iraqi, F., Darvasi, A., Soller, M., Teale, A.J., 1997. Localization ofgenes controlling resistance to trypanosomiasis in mice. Nat. Genet.16, 194–196.

Koffi, M., Solano, P., Barnabé, C., de Meeûs, T., Bucheton, B., Cuny, G.,Jamonneau, V., 2007. Genetic characterisation of Trypanosoma bru-cei s.l. using microsatellite typing: new perspectives for the molecularepidemiology of human african trypanosomosis. Infect. Genet. Evol.7, 675–684.

Koffi, M., De Meeûs, T., Bucheton, B., Solano, P., Camara, M., Kaba, D., Cuny,G., Ayala, F.J., Jamonneau, V., 2009. Population genetics of Trypanosomabrucei gambiense, the agent of sleeping sickness in Western Africa.Proc. Natl. Acad. Sci. U.S.A. 106, 209–214.

MacLean, L., Chisi, J.E., Odiit, M., Gibson, W.C., Ferris, V., Picozzi, K.,Sternberg, J.M., 2004. Severity of human african trypanosomiasis in

East Africa is associated with geographic location, parasite genotype,and host inflammatory cytokine response profile. Infect. Immun. 72,7040–7044.

MacLean, L., Odiit, M., MacLeod, A., Morrison, L., Sweeney, L., Cooper, A.,Kennedy, P.G., Sternberg, J.M., 2007. Spatially and genetically distinct

Parasito

of human serum-resistant and human serum-sensitive field isolates.

68 A. Tait et al. / Veterinary

African trypanosome virulence variants defined by host interferon-gamma response. J. Infect. Dis. 196, 1620–1628.

MacLeod, A., Tweedie, A., Welburn, S.C., Maudlin, I., Turner, C.M., Tait, A.,2000. Minisatellite marker analysis of Trypanosoma brucei: reconcilia-tion of clonal, panmictic, and epidemic population genetic structures.Proc. Natl. Acad. Sci. U.S.A. 97, 13442–13447.

MacLeod, A., Tweedie, A., McLellan, S., Hope, M., Taylor, S., Cooper, A.,Sweeney, L., Turner, C.M.R., Tait, A., 2005a. Allelic segregation andindependent assortment in T. brucei crosses: proof that the geneticsystem is Mendelian and involves meiosis. Mol. Biochem. Parasitol.143, 12–19.

MacLeod, A., Tweedie, A., McLellan, S., Taylor, S., Hall, N., Berriman, M.,El-Sayed, N.M., Hope, M., Michael, C., Turner, R., Tait, C.M.R. A., 2005b.The genetic map and comparative analysis with the physical map ofTrypanosoma brucei. Nucleic Acids Res. 33, 6688–6693.

MacLeod, A., Turner, M., Tait, A., 2007. The system of genetic exchange inTrypanosoma brucei and other Trypanosomatids. In: Barry, D., McCul-loch, R., Mottram, J., Acosta-Serrano, A. (Eds.), Trypanosomes After theGenome. Horizon Bioscience, pp. 71–91.

Majiwa, P.A.O., Hamers, R., Vanmmeirvenne, N., Matthyssens, G., 1986.Evidence for genetic diversity in Trypanosoma congolense. Parasitology93, 291–304.

Masiga, D.K., Tait, A., Turner, C.M.R., 2000. The use of the amplified frag-ment length polymorphism (AFLP) technique in parasite genetics.Parasitol. Today 16, 350–353.

Masumu, J., Geysen, D., Vansnick, N.E., Geerts, S., Van den Bossche, P.,2006a. A modified AFLP for Trypanosoma congolense isolate charac-terisation. J. Biotechnol. 125, 22–26.

Masumu, J., Marcotty, T., Geysen, D., Geerts, S., Vercruysse, J., Dorny, P., Vanden Bossche, P., 2006b. Comparison of the virulence of Trypanosomacongolense strains isolated from cattle in a trypanosomiasis endemicarea of eastern Zambia. Int. J. Parasitol. 36, 497–501.

Masumu, J., Marcotty, T., Ndeledje, N., Kubi, C., Geerts, S., Vercruysse, J.,Dorny, P., Van den Bossche, P., 2006c. Comparison of the transmissibil-ity of Trypanosoma congolense strains, isolated in a trypanosomiasisendemic area of eastern Zambia, by Glossina morsitans morsitans. Par-asitology 133, 331–334.

Matovu, E., Seebeck, T., Enaru, J.C.K., Kaminsky, R., 2001. Drugresistance in Trypanosoma brucei spp., the causative agents of sleep-ing sickness in man and nagana in cattle. Microbes. Infect. 3,763–770.

Maynard Smith, J., Smith, N.H., O’Rourke, M., Spratt, B.G., 1993. How clonalare bacteria? Proc. Natl. Acad. Sci. U.S.A. 90, 4384–4388.

McNamara, J.J., Bailey, J.W., Smith, D.H., Wakhooli, S., Godfrey, D.G., 1995.Isolation of Trypanosoma brucei gambiense from northern Uganda:evaluation of the kit for in vitro isolation (KIVI) in an epidemic focus.Trans. R. Soc. Trop. Med. Hyg. 89, 388–389.

Morrison, L.J., Tait, A., McCormack, G., Sweeney, L., Black, A., Truc, P.,Likeufack, A.C.L., Turner, C.M., Macleod, A., 2008. Trypanosoma bru-cei gambiense type 1 populations from human patients are clonaland display geographical genetic differentiation. Infect. Genet. Evol.

8, 847–854.

Morrison, L.J., Tweedie, A., Black, A., Pinchbeck, G., Christley, R., Schoene-feld, A., Hertz-Fowler, C., MacLeod, A., Turner, C.M.R., Tait, A., 2009a.Discovery of mating in the major African livestock pathogen Try-panosoma congolense. PLoS One 4, e5564.

logy 181 (2011) 61– 68

Morrison, L.J., McLellan, S., Sweeney, L., Chi Chan, C., MacLeod, A., Turner,C.M.R., Tait, A., 2009b. Role for parasite genetic diversity in differentialhost responses to Trypanosoma brucei infection. Infect. Immun. 78,1096–1108.

Morrison, L.J., Tait, A., McLellan, S., Sweeney, L., Turner, C.M.R., MacLeod,A., 2010. A major genetic locus in Trypanosoma brucei is a determinantof host pathology. PLoS Negl. Dis. 3, e557.

Ngo, H., Tschudi, C., Gull, K., Ullu, E., 1998. Double stranded RNA inducesmRNA degradation in Trypanosoma brucei. Proc. Natl. Acad. Sci. U.S.A.95, 14687–14692.

Nikolskaia, O.V., Lima, A.P.C.A., Kim, Y.V., Lonsdale-Eccles, J.D., Fukuma, T.,Scharfstein, J., Grab, D.J., 2006. Blood–brain barrier traversal by Africantrypanosomes requires calcium signaling induced by parasite cysteineprotease. J. Clin. Invest. 116, 2739–2747.

Peacock, L., Ferris, V., Bailey, M., Gibson, W., 2009. Intraclonal matingoccurs during tsetse transmission of Trypanosoma brucei. ParasitesVectors 2, 43.

Schurko, A.M., Logsdon, J.M.J., 2008. Using a meiosis detection toolkit toinvestigate ancient asexual “scandals” and the evolution of sex. Bioes-says 30, 579–589.

Schweizer, J., Tait, A., Jenni, L., 1988. The timing and frequency of hybridformation in African trypanosomes during cyclical transmission. Par-asitol. Res. 75, 98–101.

Simo, G., Cuny, G., Demonchy, R., Herder, S., 2007. Trypanosoma bruceigambiense: study of population genetic structure of central Africanstocks using amplified fragment length polymorphism (AFLP). Exp.Parasitol. 118, 172–180.

Tibayrenc, M., Kjellberg, F., Ayala, F.J., 1990. A clonal theory of parasiticprotozoa: the population structures of entamoeba, giardia, leishma-nia, naegleria, plasmodium, trichomonas, and trypanosoma and theirmedical and taxonomical consequences. Proc. Natl. Acad. Sci. U.S.A.87, 2414–2418.

Tait, A., 1980. Evidence for diploidy and mating in trypanosomes. Nature287, 536–538.

Tait, A., Buchanan, N., Hide, G., Turner, C.M.R., 1996. Self-fertilisation inTrypanosoma brucei. Mol. Biochem. Parasitol. 76, 31–42.

Tait, A., Masiga, D., Ouma, J., MacLeod, A., Sasse, J., Melville, M., Lindegard,G., McIntosh, A., Turner, C.M.R., 2002. The genetic analysis of complexphenotypes in T. brucei. Phil. Trans. Roy. Soc. B 367, 89–99.

Tait, A., MacLeod, A., Tweedie, A., Masiga, D., Turner, C.M.R., 2007. Geneticexchange in T. brucei: evidence for mating prior to metacyclic stagedevelopment. Mol. Biochem. Parasitol. 151, 133–136.

Tilley, A., Welburn, S.C., Fevre, E.M., Feil, E.J., Hide, G., 2003. Trypanosomabrucei: Trypanosome strain typing using PCR analysis of mobilegenetic elements (MGE-PCR). Exp. Parasitol. 104, 26–32.

Turner, C.M.R., Sternberg, J., Buchanan, N., Smith, E., Hide, G., et al., 1990.Evidence that the mechanism of gene exchange in Trypanosoma bru-cei involves meiosis and syngamy. Parasitology 101, 377–386.

Welburn, S.C., Maudlin, I., Milligan, P.J.M., 1995. Trypanozoon: infectivity tohumans is linked to reduced transmissibility in tsetse. I. Comparison

Exp. Parasitol. 81, 404–408.Young, C.J., Godfrey, D.G., 1983. Enzyme polymorphism and the distribu-

tion of Trypanosoma congolense isolates. Ann. Trop. Med. Parasitol.77, 467–481.