DNA metabolism and genetic diversity in Trypanosomes

Mutation Research 612 (2006) 40–57 www.elsevier.com/locate/reviewsmr Community address: www.elsevier.com/locate/mutres

Review

DNA metabolism and genetic diversity in Trypanosomes Carlos Renato Machado a, Luiz Augusto-Pinto a, Richard McCulloch b, Santuza M.R. Teixeira a,* a

Department of Biochemistry and Immunology, Universidade Federal de Minas Gerais, Belo Horizonte, Brazil b Wellcome Centre for Molecular Parasitology, University of Glasgow, Anderson College, 56 Dumbarton Road, Glasgow G11 6NU, UK Received in revised form 14 March 2005; accepted 19 May 2005 Available online 22 July 2005

Abstract Trypanosomes are protozoan parasites that cause major diseases in humans and other animals. Trypanosoma brucei and Trypanosoma cruzi are the etiologic agents of African and American Trypanosomiasis, respectively. In spite of large amounts of information regarding various aspects of their biology, including the essentially complete sequences of their genomes, studies directed towards an understanding of mechanisms related to DNA metabolism have been very limited. Recent reports, however, describing genes involved with DNA recombination and repair in T. brucei and T. cruzi, indicated the importance of these processes in the generation of genetic variability, which is crucial to the success of these parasites. Here, we review these data and discuss how the DNA repair and recombination machineries may contribute to strikingly different strategies evolved by the two Trypanosomes to create genetic variability that is needed for survival in their hosts. In T. brucei, two genetic components are critical to the success of antigenic variation, a strategy that allows the parasite to evade the host immune system by periodically changing the expression of a group of variant surface glycoproteins (VSGs). One component is a mechanism that provides for the exclusive expression of a single VSG at any one time, and the second is a large repository of antigenically distinct VSGs. Work from various groups showing the importance of recombination reactions in T. brucei, primarily to move a silent VSG into an active VSG expression site, is discussed. T. cruzi does not use the strategy of antigenic variation for host immune evasion but counts on the extreme heterogeneity of their population for parasite adaptation to different hosts. We discuss recent evidence indicating the existence of major differences in the levels of genomic heterogeneity among T. cruzi strains, and suggest that metabolic changes in the mismatch repair pathway could be an important source of antigenic diversity found within the T. cruzi population. # 2005 Elsevier B.V. All rights reserved. Keywords: DNA repair; Recombination; Genetic variability; Trypanosoma

* Corresponding author. Tel.: +55 31 3499 2665; fax: +55 31 3499 2614. E-mail address: [email protected] (Santuza M.R. Teixeira). 1383-5742/$ – see front matter. # 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.mrrev.2005.05.001

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Contents 1. 2. 3. 4.

5.

6.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T. cruzi and Chagas disease. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T. brucei and sleeping sickness . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genetic diversity in T. brucei and T. cruzi. . . . . . . . . . . . . . . . . . . . . . . . 4.1. Antigenic variation in T. brucei . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Population structure and genetic diversity of T. cruzi . . . . . . . . . . . DNA metabolism and the generation of genetic diversity in Trypanosomes. 5.1. Mismatch repair and genetic diversity in T. cruzi . . . . . . . . . . . . . . 5.2. DNA recombination and T. cruzi genetic diversity . . . . . . . . . . . . . 5.3. DNA recombination and antigenic variation in T. brucei . . . . . . . . . Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction The protozoan parasites Trypanosoma brucei and Trypanosoma cruzi are of considerable medical importance, since these species are the etiologic agents of sleeping sickness (African Trypanosomiasis) and Chagas disease (American Trypanosomiasis), respectively [1]. Both are hemoflagellates of the order kinetoplastida, family Trypanosomatidae, characterized by the presence of a single flagellum and a mitochondrion whose genome is arranged in a unique network known as the kinetoplast DNA [2]. As members of a family of highly divergent eukaryotes [3], Trypanosomes utilize several distinctive biological strategies, including polycistronic transcription of most of their genomes [4–5], RNA polymerase Imediated transcription of protein-coding genes [6], RNA trans-splicing to generate mature, capped mRNA molecules of nuclear genes [7] and extensive RNA editing to generate functional mRNAs of mitochondrial genes [8]. Although the development of powerful genetic tools [9] has allowed rapid progress towards understanding various aspects of their biology, as well as an intricate appreciation of their population genetics [10], studies related to core functions involved in DNA metabolism, particularly DNA repair and recombination, have been limited. Recent work, however, together with the imminent unveiling of the genome sequences of T. brucei and T. cruzi (and the related kinetoplastid parasite Leishmania major), indicate that, despite the fact that the two parasites present a high degree of homology for various genes

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involved in DNA metabolism, they utilize different pathways to generate genetic diversity. It seems likely that these differences are related to the strikingly different strategies that each parasite has evolved to cope with the host immune responses.

2. T. cruzi and Chagas disease Chagas disease, or American Trypanosomiasis, is caused by the intracellular protozoan parasite T. cruzi. The T. cruzi life cycle is developed among small wild mammals in an enzootic sylvatic transmission cycle established in the American continent. Vectorial transmission occurs through the contact of mammals with the feces of infected triatomines of the family Reduvidae. Infective trypomastigotes, present in the feces, are inoculated through skin punctures or mucous membranes of the mammalian host (‘‘posterior’’ transmission). After reaching the bloodstream, trypomastigotes invade a variety of cells and convert into amastigote forms, which replicate in the cytosol by binary division. After multiple rounds of replication, amastigotes transform into trypomastigotes that are released by rupture of the host cell plasma membrane. Trypomastigotes and amastigotes present in the blood of the infected mammal complete the cycle when they are taken up in a blood meal by a reduviid bug. Following an incubation period of 7–9 days, classical symptoms (including fever, lymph node enlargement and subcutaneous edema) of acute phase Chagas disease begin, during which time the parasite undergoes

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an intense period of multiplication and invades the bloodstream and several organs. The infection then proceeds to a chronic phase, which has an unpredictable clinical course that ranges from absence of symptoms to severe disease. The major affected organs are the heart, colon and esophagus, characterizing the digestive and cardiac forms of Chagas disease, or both [11]. In endemic countries, it is estimated that 16–18 million people are infected by T. cruzi and the ‘‘domiciliation’’ of the triatomines exposes at least 90 million individuals to the risk of infection in regions extending from Chile in South America to areas of the Southern states in the USA. With no vaccine or effective treatment available for T. cruzi in large-scale public health interventions, the main control strategy relies on prevention of transmission, principally by eliminating the domestic insect vectors and control of transmission by blood transfusions (for general information on T. cruzi and Chagas disease, including a detailed description of the parasite life cycle see: http://www.who.int/tdr/diseases/chagas/).

3. T. brucei and sleeping sickness African Trypanosomiasis is a disease popularly known as sleeping sickness in humans and N’gana in cattle. Three subspecies of the pathogen T. brucei are transmitted to mammals through the bite of tsetse flies (of the species Glossina; ‘‘anterior’’ transmission), and two cause human disease [12]. The T. brucei life cycle (http://www.who.int/tdr/diseases/tryp/) alternates between multiple forms of the parasite during its development in the gut and salivary glands of the tsetse fly, and a smaller number of forms in the bloodstream and tissue vasculature of the mammal (also reviewed in [13]). The clinical manifestations of T. brucei infection in humans are dependent on the sub-species involved. T. brucei rhodesiense, found in Southern and Eastern Africa, causes highly virulent, acute infections, whereas infections by T. brucei gambiense, common in Central and Western Africa, are delayed and chronic. In both cases, symptoms begin with fever, headache, joint pain, itching and lethargy, and then, as the parasites cross the blood– brain barrier, irritability, confusion, slurred speech, poor coordination and profound fatigue ensue. Untreated, coma and death invariably follow.

It is estimated that about 60 million people in 36 nations live in conditions ripe for African sleeping sickness. Although as many as 45,000–50,000 cases are reported each year, probably 10 times that many people are actually infected. Many die undiagnosed, and only 10% are treated, even thought there are effective treatments for African sleeping sickness, particularly if the disease is diagnosed early. (World Health Organization, http://www.who.int/tdr/diseases/ tryp/).

4. Genetic diversity in T. brucei and T. cruzi Knowledge about the mechanisms employed by Trypanosomes to generate genetic diversity is a crucial step towards understanding the immunological and ecological relationships between African and American Trypanosomes and their hosts. In T. brucei, genetic diversity plays a central role in the evasion of the mammalian immune responses through a mechanism called antigenic variation (see below). However, other strategies for immune evasion, in many different protozoan parasites, have been suggested [14] and these may also be dependent on genetic diversification. Moreover, it has been little explored how genetic diversity might influence more subtle interactions with the host, including adaptation to new hosts or changes within a given host. Any considerations of such issues are dependent on the life cycle and parasitic strategy of the organism. T. brucei lives in the blood and lacks any intracellular stage, and thus the parasite is a target for antibody-mediated elimination. T. cruzi, in contrast, rapidly invades a variety of host cell types where it multiplies away from the damaging elements of the host humoral immune response. 4.1. Antigenic variation in T. brucei Antigenic variation is the immune evasion strategy evolved by African Trypanosomes in which it is clear that genetic variability is critical. This general strategy, consisting of periodic changes in exposed antigens, is used by a diverse array of microbes [13,15]. In T. brucei, antigenic variation consists of changes in the expression of variant surface glycoproteins (VSGs) [16–18]. VSGs are attached to the surface membrane via a GPI anchor [19], and form a

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tightly packed ‘coat’ which shields invariant surface molecules from the host immune system and protects against the alternative complement response. Switching the expressed VSG is needed to allow the parasite population to escape immune killing mediated by antibodies directed against the previously expressed VSG. Two genetic components are critical to the success of T. brucei antigenic variation: a mechanism for the exclusive expression of a single VSG at any one time, and a huge repository of antigenically distinct VSGs. VSG genes are expressed in the mammal only when present in subtelomeric transcription units termed the bloodstream VSG expression sites (ES), which are transcribed by RNA polymerase I [6] and are silenced in the tsetse fly [20]. Surprisingly, T. brucei has not one such ES, but around 20. As only one ES is transcriptionally active at one time, T. brucei has evolved a mechanism [21] for silencing the majority of the ESs and selecting one ES for VSG expression, which remains active only in bloodstream forms. The molecular basis of this mechanism, named allelic exclusion, appears to involve a discrete sub-nuclear entity, termed the expression site body, which contains the active ES [22] (Fig. 1). Much remains to be learned about the expression site body, including its molecular composition, existence throughout the T. brucei life cycle [23] and, crucially, its potential role in VSG switching. One route that T. brucei uses to switch the expressed VSG is to silence the active ES and activate a silent ES. This transcriptional switching reactions appears to be a coordinated process, in that attempts to select for cells which transcribe stably more than one ES generated only switching intermediates that express transiently two [24], but not three [25], ESs. Whether the expression site body has a regulatory function in VSG switching or is simply the site of ES transcription is unknown. The fact that the inactive ESs are partly transcribed [26–28], and that transcription elongation and processing of these multi-genic transcription units appears to contribute to the regulation of VSG expression [18,23,26,29], must also be considered in understanding this complex process. No clear correlation can be made between DNA rearrangements or sequence changes in the ES and transcriptional VSG switching [30–32], meaning that this antigenic variation reaction does

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Fig. 1. VSG transcriptional control and the expression site body of Trypanosoma brucei. The long slender replicative form of T. brucei found in the mammalian bloodstream is depicted, and the insert shows a cartoon of the nucleus (light blue), containing the nucleolus (light grey) and a subnuclear entity termed the expression site body (ESB; dark blue). For simplicity, only four T. brucei chromosomes are depicted in the nucleus, and the inserts depict two telomeric loci of variant surface glycoprotein (VSG) gene expression, termed the expression sites (ES), that are present at the chromosome ends. The VSG (grey box) is always found adjacent to the telomere (vertical line), and is co-transcribed with a number of expression site associated genes (black boxes) and flanked upstream by an array of degenerate 70 bp repeats (vertically hatched box). The actively transcribed ES is localised to the ESB, and transcription (dotted line) initially occurs as a multigene transcript encompassing all the protein-coding sequence. Inactive ESs do not co-localise with the ESB, but T. brucei can switch the transcriptional status between ES, thus executing a change in expressed VSG. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

not involve genetic variation directly. However, ESs are complex structures containing multiple non-VSG genes, termed the expression site associated genes (ESAGs), and differences in both the number and sequence of ESAGs encoded by different ESs are apparent [33]. It is therefore an interesting question whether or not these genetic variations, revealed by transcriptional switching, might have implications for T. brucei growth, infectivity or host adaptation [34]. ESAG6 and ESAG7 together encode a heterodimeric, VSG-related surface receptor for host transferrin [35–38]. ESAG6/7 heterodimers encoded from different ESs have been shown to have differing affinities for transferrins from different mammals [39,40]. Initially, the sequence variations in the different encoded transferrin

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Fig. 2. VSG switching by gene conversion in T. brucei. (a) Switching of the expressed VSG by gene conversion involves copying a silent VSG (red box) into the ES (see (b) replacing the resident VSG (blue box)). The silent VSG sequences in these reactions can be present in arrays on the megabase chromosomes, at the telomeres of minichromosomes (which contains 177 bp repeated sequence; light grey box) or in an inactive ES (only the bloodstream stage version is diagrammed). The amount of sequence copied during gene conversion is illustrated, and normally encompasses the VSG ORF and extends upstream to the 70 bp repeats (vertically hatched box). This can vary (depicted by a dotted line), however: the upstream conversion limit can extend to different parts of the 70 bp repeats, or into the conserved expression site associated genes, whilst VSG gene conversions encompassing the downstream repeats that make up the telomere have been also described. (b) Segmental gene conversion of VSG genes. Switching of the expressed VSG can occur by gene conversions of segments from multiple VSG pseudogenes (here shown as three genes, depicted by red, blue and green boxes with crosses), a process often referred to as mosaic VSG formation. These reactions normally involve the poorly conserved open reading frame sequences of VSGs. To be expressed, the newly created VSG must become present in the ES (see Fig. 1 but whether or not the reaction occurs within the actively transcribed ES (as drawn) is unclear. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

receptors, which are concentrated in a small region, were proposed to represent a form of antigenic variation, analogous to the VSG system [41]. However, as the variable regions appear not to be surface exposed [42], the differing affinities for host transferrin have been interpreted more recently as a means by which T. brucei adapts to different hosts [39], or may ensure efficient iron uptake in the face of host antibodies against ESAG6/7 [41,43,44]. ESassociated host-adaptation can also be invoked for another VSG-related ESAG, the serum-resistanceassociated protein (SRA). T. brucei brucei, unlike T. brucei rhodesiense, is unable to infect humans because, at least in part, it is lysed by apolipoprotein L-1 (apoL1) in human serum [45]. T. brucei rhodesiense is able to survive in humans by expressing SRA, which binds apoL1 and neutralizes its lytic activity. Xong et al. [46] showed that SRA in T. b. rhodesiense can be encoded by an ES, and the resistance of the parasite to human serum was dependent on whether or not the ES was being expressed. However, SRA-like genes may not be limited to the ES. T. b. gambiense is resistant to lysis

by human serum but lacks an SRA gene [47], meaning that SRA-independent mechanisms for human serum resistance must also occur [48,49]. Experimental evidence suggests also that mammalian serum may contain trypanolytic factors other than apoL1 [50], but the potential mechanisms of trypanosome resistance to these are unknown. T. brucei antigenic variation occurs primarily not through transcriptional switches amongst the ESs, but by reactions which select a silent VSG gene, or genes, and generate a new VSG coat following recombination reactions into the ES. Long-standing, indirect estimates suggested that T. brucei contains a silent store of around 1000 VSG genes [51]. Sequencing the T. brucei genome, and annotation of the VSG repertoire (L. Marcello, J.D. Barry, personal communication), confirms that this is a fairly accurate estimate. The T. brucei genome is composed of 11 diploid megabase chromosomes [52], a large number (perhaps 100–200) of minichromosomes [53,54] and a set of intermediate-sized chromosomes [55]. Silent VSGs are found on all types of chromosome. A number of recombination reactions have been

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described which can move a silent VSG into the ES for activation (recently reviewed by [13]). The most commonly observed is VSG gene conversion, or duplicative transposition, where a copy is made from a silent VSG and replaces the VSG resident in the ES (Fig. 2a). For a phenotypic VSG switch, this reaction should occur into the active ES, but some putative examples of phenotypically silent gene conversions of telomeric VSGs have also been described [56], which may reflect the predominant occupation of T. brucei subtelomeres by sequences associated with antigenic variation [57]. Reciprocal recombination events, in which the VSG-containing ends of two chromosomes are swapped, have also been described (not shown). Finally, new VSGs can be created by ‘mosaic’ VSG formation, where a new VSG sequence is created from a number of silent VSGs or VSG pseudogenes (Fig. 2b). This has been considered to be a rare event, since such mosaic VSGs were isolated late in infections [58,59]. In fact, the complete genome of T. brucei now demonstrates that most of the T. brucei VSG repertoire is defective (composed either of VSG pseudogenes or VSG fragments; L. Marcello, J.D. Barry, personal communication) and therefore mosaic VSG formation most likely makes a very significant contribution to antigenic variation, as has been predicted [60]. Genetic variability associated with immune evasion in T. brucei operates at a number of levels. Firstly, there is the overt variation associated with the movement of VSGs into the ESs, from a number of loci around the genome, specifically to avoid antiVSG antibodies generated the course of an infection. Beyond this, however, we know less about how comparable the VSG repertoires are between different strains, subspecies and species of Trypanosoma (although there appears to be great diversity [61]), or the potential for genetic exchange between Trypanosomes [62,63] to diversify this further [64]. By addressing these questions we may be able to understand how the repertoire arose in evolution, and the rate at which it is diversifying. 4.2. Population structure and genetic diversity of T. cruzi T. cruzi does not use a strategy of antigenic variation for host immune evasion and the relation-

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ship between genetic variability and immune evasion or host adaptation has been less characterized. However, the T. cruzi life cycle requires the development, in the mammalian host, of a nonreplicative extracellular blood stage and a replicative intracellular form. The parasite evades macrophagemediated destruction of intracellular stages by escaping from the phagolysosome and by invading non-phagocytic cells, as well as by modulating the pattern of expression of cytokines [65,66]. It also possesses molecules that mimic those of host cells, inhibiting specific immune recognition and eliciting autoimmunity [67]. Thus, a variable repertoire of surface molecules and antigenic components that are highly polymorphic would represent a useful arsenal that the T. cruzi population can count on to increase the success rate of the cell invasion process and parasite survival. In the absence of antigenic variation, such genetic diversity must be generated by diverse alleles at defined loci, in which case it is a property of the population called polymorphism. T. cruzi has an heterogeneous population, composed of a pool of strains. Early studies of T. cruzi isolates from different origins had demonstrated the presence of a large range of strains with distinct characteristics. This striking intra-specific variation has been extensively documented by molecular inferences and biological characterization, which includes morphology of blood forms, curves of parasitemia, virulence, pathogenicity, sensitivity to drugs, antigenic profile, growth rate, metacyclogenesis and tissue tropism (reviewed in [10,68]). In spite of the broad genetic diversity observed among different isolates, two major lineages of the parasite, named T. cruzi I and II, have been identified [69]. These divergent lineages occupy distinct ecological environments: the silvatic cycle (T. cruzi I) and domestic cycle (T. cruzi II) of Chagas disease [70,71] as well as distinct silvatic host associations [72]. In contrast to T. cruzi I strains, which induce low parasitism in human chagasic patients, T. cruzi II causes human infections with high parasitemia in endemic areas of southern countries in South America [71]. Several molecular markers, including the 24S ribosomal gene (rDNA) and miniexon genes [73], permit a clear distinction between strains belonging to T. cruzi I or II. Others analyses [74] based on RAPD/ MLEE permitted the division of T. cruzi II into five

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sub-groups: T. cruzi IIa, IIb, IIc, IId and IIe. Still, the zymodeme patterns described by Miles et al. [75] and the studies by Machado and Ayala [76,77] and Augusto-Pinto et al. [78], looking at various protein coding genes, indicated the existence of three or more subdivisions in the T. cruzi population, with one lineage mainly composed of hybrid strains. The biological and molecular aspects underlying T. cruzi population structure, particularly regarding the hybrid strains are under intense debate. Together with the epidemiology of Chagas disease these issues are beyond the scope of the present article and have been extensively addressed in recent reviews [10,68]. However, there is emerging evidence, through the characterization of genes involved with DNA metabolism (see below), indicating that different T. cruzi isolates present different levels of genotypic heterogeneity, which may have a major impact in Chagas disease. In addition to the high population polymorphism, the T. cruzi genome contains a large number of gene families encoding proteins with various functions, as well as pseudogenes. Duplicated genes arise frequently in eukaryotic genomes, either via local recombination events that generate tandem duplications, larger-scale events that duplicate chromosomal regions or entire chromosomes, or genome-wide events that result in complete genome duplication (polyploidization) [79]. Because duplicated genes are believed to be initially redundant in function, it is commonly thought that one member of the pair will usually become silenced by degenerative mutation. The function of a duplicated gene will be maintained if it is under positive selection, such as some drug resistance [80], or if it generates new functions [81]. A striking example of gene duplication that may be related to the parasite’s immune evasion strategy is the heterogeneity and functional diversity of the trans-sialidase and mucin gene families. T. cruzi has 100s of genes encoding trans-sialidases, which catalyzes the transfer of sialic acid from host glycoconjugates to several distinct mucin-like molecules located on the parasite surface membrane. It has been shown that sequence divergence of members of the trans-sialidase family results in drastic changes in enzymatic activity as well as in distinct patterns of gene expression during the parasite life cycle [82].

5. DNA metabolism and the generation of genetic diversity in Trypanosomes If genetic diversity is essential for survival of both parasites in their host, in T. brucei, primarily for antigenic variation during the course of a single infection and, in T. cruzi, for antigenic diversity (allelic polymorphism) in the population, which are the mechanisms developed by these parasites to generate the required genomic variation? In any population, mutation rates must be carefully balanced between the need to create diversity to be offered as a substrate for adaptive evolution and to maintain core cellular functions [83,84]. The DNA repair machinery is a key component in striking this balance, in that it prevents the establishment of detrimentally high genomic mutation rates. But, if DNA repair does not fail at some level, then populations do not evolve. Under this perspective, the components of the DNA recombination and repair machinery in T. brucei and T. cruzi must be understood in order to examine the mechanisms involved with the generation of genetic diversity. 5.1. Mismatch repair and genetic diversity in T. cruzi Studies by Perez et al. [85] and Olivares et al. [86], characterizing a T. cruzi apurinic/apyrimidinic endonuclease gene, and by Farez-Vidal et al. [87], characterizing a uracil-DNA glycosylase gene, were the first to describe the components of a DNA repair pathway in this parasite. To date, no evidence has connected these components of the base excision repair machinery with T. cruzi genetic variablity. Later work began to examine the mismatch repair (MMR) pathway in T. cruzi and resulted in the cloning and characterization of the MSH2 gene [88]. MMR has a key function in recognizing and repairing base mismatches and frame shift mismatches that escape DNA polymerase proofreading during DNA replication. In addition, MMR can also recognize mispaired bases that arise during recombination, and this can have two outcomes: repair of the mismatched base can cause gene conversion, or the MMR machinery can abort the recombination reaction. The machinery of MMR is well-conserved in evolution, and the role of this repair pathway in

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decreasing the mutation rate and constraining recombination is well established [89–93]. In some types of human cancers, it is clear that the loss of MMR activity plays a determinant role [94–96]. Pathogenic and commensal bacteria with elevated rates of mutations, often due to having a defective MMR machinery, are found frequently in natural isolates, and the rate of mutation in bacterial populations can vary in experimental systems placed under environmental adaptation. These studies indicate that it may be possible for bacterial populations to select for an increased mutation rate to permit greater environmental or host adaptability [97–105]. Similar modulations of mutation rate have not been described unequivocally in eukaryotic organisms [106,107]. In an attempt to better understand the population structure of T. cruzi, we have asked whether differences in DNA repair efficiency may exist among the strains that compose the main lineages that can be identified within the parasite population. We began to investigate the influence of MMR on generating genetic diversity in T. cruzi with the analyses of singlenucleotide polymorphisms (SNP) of the TcMSH2 gene. Augusto-Pinto et al. [78] demonstrated that TcMSH2 SNPs correlate with the existence of three phylogenetic lineages in T. cruzi. More importantly, T. cruzi strains belonging to each of the main lineages present differences in MMR efficiency. Microsatellite loci of strains representative of each haplogroup, cultured in the presence of hydrogen peroxide, displayed allelic variation in T. cruzi II strains, while no such microsatellite instability was found in strains belonging to the T. cruzi I lineage. Also, cells from MSH2-defined haplogroups B and C (all of them having T. cruzi II and hybrid markers) were considerably more resistant to cisplatin treatment, a characteristic known to be conferred by deficiency of MMR in eukaryotic cells [108,109]. These studies suggest that, at least under genotoxic stress conditions, strains belonging to the T. cruzi I lineage have a more efficient MMR activity than T. cruzi II strains [78]. This proposition implies that the two T. cruzi populations will have distinct genetic diversity, since MMR efficiency is directly associated with mutation rate [89,110]. Preliminary work on the analysis of DNA sequences derived from two T. cruzi multi-gene families encoding the TcRBP48 antigen [111] and

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the amastin surface protein [112] corroborates this hypothesis [113]. This work is also aimed to addressing more directly the relationship between T. cruzi genetic diversity and immune evasion and host adaptation since these multi-gene families encode surface proteins or parasite antigens recognized by sera from chagasic patients. Sequence analyses of PCR-amplified genomic fragments derived from strains belonging to T. cruzi I, II lineages and strains showing characteristics of hybrid strains, showed that the distribution of sequence variation in these samples correlated well with the putative division of T. cruzi into three lineages, as defined by polymorphisms found in the TcMHS2 gene. Most importantly, both multi-copy gene families, encoding TcRBP48 and amastins, were found to be much less variable in strains belonging to the T. cruzi I lineage compared with their counterparts from T. cruzi II and hybrid strains [113]. Taken together, these findings indicate that metabolic changes in the MMR pathway could be an important source of antigenic diversity found within the T. cruzi population (Fig. 3). Whether or not this links directly with immune evasion, or with host adaptation, remains to be determined. However, proteomic analyses recently described by Buscaglia et al. [114], suggest that diversity of the surface antigens that are part of the large family of the mucinlike glycoproteins expressed in trypomastigotes may represent a refined T. cruzi strategy to elude the mammalian-host immune system. A recent work describing the antibody response to recombinant trans-sialidase proteins encoded by distinct copies of SA85-1 genes also suggest that in different mouse strains and human subjects some members of this large protein family are more immunogenic than others [115]. Results from two other recent studies where parasite infectivity and the immune response to antigens of parasite strains belonging to T. cruzi I and II lineages were compared indicate that strains belonging to the T. cruzi II lineage (which appears less efficient at MMR) display increased infectivity in experimental animals [116] and are present in the majority of chronic chagasic patients [117]. Di Noia et al. [117] tested the reactivity of recombinant TSSA mucins, derived from T. cruzi I and II parasites, with sera from chagasic individuals, and found that the high prevalence of antibodies anti-TSSA is restricted to the

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5.2. DNA recombination and T. cruzi genetic diversity

Fig. 3. Differential MMR efficiency and distinct genetic variety within the Trypanosoma cruzi population. Parasites from MSH2 haplogroup C, presenting markers characteristic of T. cruzi II strains are found associated with the domestic cycle and are responsible for most cases of human disease. Together with parasites belonging to MSH2 haplogroups B or B/C, some of them presenting characteristics of hybrid strains, they seem to carry a less efficient MMR when compared with parasites belonging to MSH2 haplogroup A (all of them showing molecular markers typical of T. cruzi I strains, a group associated with the sylvatic cycle of the parasite). Since MMR efficiency is directly associated with mutation rate we speculate that strains from T. cruzi II and hybrid strains (MSH2 B and C) present a higher mutation rate, compared to T. cruzi I strains (MSH2 A). This might be reflected in the increased genetic diversity verified in the sequences of multi-copy gene families, which were found to be much less variable in strains belonging to the T. cruzi I lineage compared to the corresponding loci in the genome of T. cruzi II and hybrid strains.

T. cruzi II isoform of this antigen. According to their results, no human infection by T. cruzi I strains could be observed. Similarly, Risso et al. [116] showed that whereas infection with T. cruzi II isolates induces 100% mortality in mice, infection with several T. cruzi I strains result in chronic disease, even with a 1000fold higher inoculum. These latter authors also showed that the expression of, and host immune response against, the T. cruzi trans-sialidase is diverse: whereas T. cruzi II strains secrete higher amounts of trans-sialidase, neutralizing antibodies against the protein were detected only in T. cruzi I-infected mice.

If MMR emerges as a DNA metabolism pathway involved in the generation of genetic diversity of T. cruzi, recombination might be considered a repair pathway that reduces this variability. Analyses of several genetic loci among different parasite populations have demonstrated that T. cruzi presents drastic departures from Hardy–Weinberg expectations and strong linkage disequilibrium (reviewed in [10]). These findings have been interpreted as meaning that T. cruzi has a clonal population structure in which sexual reproduction is rare, or even absent. Under this perspective, each clone represents an independent lineage that divides by binary division and evolves by mutation only from an ancient ancestor [118]. One expectation from this is that T. cruzi, as an asexual organism, should present the ‘‘Meselson effect’’: in the absence of sexual recombination, the two alleles at any locus evolve independently, and only functional constraints prevent unlimited divergence. Such high levels of allele divergence have indeed been found in the asexual Bdelloid rotifers [119]. In T. cruzi, however, isoenzyme and microsatellite data have shown an excess of allelic homozygosity [75,120]. Gene sequencing revealed very little divergence between two alleles in strains belonging to T. cruzi I or II lineages [76]. Contrasting with the asexual rotifers, allelic divergence in T. cruzi was observed only in nuclear genes from some strains presenting hybrid characteristics, such as the CL Brener strain chosen as the reference strain for the genome project. It has been proposed that four different mechanisms could be involved in decreasing the T. cruzi allele divergence: automixing, sexual exchange, gene conversion and mitotic recombination [76]. Automixing is a phenomenon that has never been detected in trypanosomatids. Gaunt et al. [121], on the other hand, have demonstrated unequivocally that genetic exchange can occur in T. cruzi. Using transgenic parental strains, these authors provided evidence, using isoenzyme and karyotype analyses, for fusion of parental genotypes and loss of alleles and homologous recombination in the resulting hybrids. Evidence based on nucleotide sequences has also indicated the presence of hybrids in natural

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populations [76]. However, since the major characteristics of the hybrid strains is their heterozygosity [76], genetic exchange cannot explain the high levels of allelic homozygosity observed in T. cruzi. In fact, the presence of hybrids results in increased divergence. It seems likely that loss of heterozygosity and gene conversion, either during genetic exchange or in the resulting hybrids, as well as mitotic recombination, act to limit the ‘‘Meselson effect’’, decreasing the genetic diversity within and between individuals. The possibility that these processes may not be associated directly with genetic exchange can be seen in the work by Schon and Martens [122], who reported that the ancient asexual ostracod species Darwinula stevensoni shows very low nucleotide divergence in three nuclear regions. Likelihood permutation tests confirmed the presence of gene conversion in the multicopy internal transcribed spacer sequence, but rejected rare or cryptic forms of sex as a general explanation for the low genetic diversity in the organism [122]. It thus seems likely that examining the functions of recombination in T. cruzi may be very informative with regard to genetic diversity, and genetic exchange, but little work has been done to date. Studies in many organisms, including T. brucei (see below) of the Rad51 gene, which encodes a key element in homologous recombination, has provided a valuable starting point. Sequence analyses of Rad51 from several organisms have shown that this gene is one of the most conserved genes described. A phylogenetic tree constructed with Rad51 sequences derived from T. brucei, Leishmania and the available Rad51 sequence obtained from the T. cruzi genome project database (www.genedb.orgg), as well as from several other organisms, confirms the high level of homology among these sequences. It also shows a close relationship between the sequences derived from trypanosomatids, which are grouped in a well-defined cluster presenting a highly significant bootstrap value (Fig. 4). Such high level of homology observed among Trypanosome Rad51 sequences may be related to the importance of recombination for parasite DNA metabolism. Studies investigating T. cruzi resistance to exposure to gamma irradiation point toward a crucial role of DNA recombination for parasite survival. Early attempts to develop protocols for parasite attenuation

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[123] indicated that most T. cruzi strains display an unusually high resistance to ionizing radiation. These findings have been confirmed by testing the sensitivity to gamma irradiation of epimastigote cultures from different T. cruzi lineages: all strains survived doses as high as 1000 Gy. (C.G. Silva, S.M.R. Teixeira, C.R. Machado, unpublished). Moreover, exposure to these levels of gamma radiation resulted in high proportions of chromosome breakage events followed by rapid kinetics of chromosomal repair as assessed by pulse field gel electrophoresis. Gamma irradiation also led to up-regulation of the expression of T. cruzi Rad51 gene whose product, among other functions, has a pivotal role in repairing double-strand breaks [124]. 5.3. DNA recombination and antigenic variation in T. brucei The genetic factors that underpin recombinogenic antigenic variation in T. brucei have begun recently to be elucidated. This pathway of VSG switching appears to be predominantly a form of homologous recombination, in keeping with molecular descriptions of the process [125]. Mutation of the T. brucei Rad51 gene impairs VSG switching [126]. In contrast, mutation of the T. brucei genes encoding either component of the KU70/80 heterodimer, a major factor in a competing from of DNA double strand break repair found in other organisms termed non-homologous end-joining [127], has no effect on VSG switching [128]. However, two observations suggest that describing VSG recombination simply as a form of RAD51-dependent recombination does not encapsulate either the number of potential recombination pathways that can provide for VSG switching, nor the potential genetic complexity of the RAD51 recombination reaction that is used (at least predominantly). The first observation is that T. brucei RAD51 mutants remain capable, albeit at a reduced rate, of catalysing recombination [125] and recombinationbased VSG switching, meaning that RAD51-independent recombination reactions must therefore be present. One such pathway has been described that acts on DNA substrates 7–15 bp in length, often containing base mismatches [129]. This reaction is clearly distinct from T. brucei RAD51-dependent recombination, which requires minimally around

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Fig. 4. Phylogenetic relationships between RAD51 and recA protein sequences showed in a Neighbor-joining (NJ) tree for Dayhoff PAM distances. Parsimony trees were also constructed, which produced very similar results (not shown). Numbers above each branch represent the number of times the branch was found in 1000 bootstrap replicas. The species and accession number in Genbank for each taxa are: Trypanosoma cruzi (AAO72729), Leishmania major (AAC163344), Trypanosoma brucei (AAD51713), Leishmania donovani (AAQ96331), Cynops pyrrhogaster (BAA78377), Danio rerio (NP_998371), Xenopus laevis (BAA07501), Pan troglodytes (XP_522884), Cricetulus griseus (P70099), Homo sapiens (AAV38511), Canis familiaris (BAB91246), Gallus gallus (P37383), Mus musculus (NP_035364), Oryctolagus cuniculus (AAC28561), Anopheles gambiae (EAA05962), Bombyx mori (AAB53330), Oryza sativa (BAB85492), Schizosaccharomyces pombe (P36601), Candida albicans (EAK94361), Zea mays (Q67EU8), Encephalitozoon cuniculi (CAD25992), Dictyostelium discoideum (AAO52375), Kluyveromyces lactis (AAP13463), Lycopersicon esculentum (Q40134), Saccharomyces cerevisiae (NP_011021), Ashbya

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100 bp of sequence homology and is severely affected by base mismatches [130]. The functions that the microhomology dependent recombination pathway provide in T. brucei are not known, as are the factors which catalyze or regulate the reaction, whether or not it is used also by other trypanosomatids and, crucially, if it can contribute to VSG switching. The VSG switching assay used to examine the function of RAD51 (and the putative KU70/80 heterodimer) in antigenic variation does not allow for analysis of mosaic VSG formation, so it is not yet possible to say what type of recombination reaction this may be. However, the mosaic VSG genes whose formation have been described arose using recombination based on short (around 85 bp) stretches of DNA sequence sharing limited (around 70%) identity [131]. It is therefore tempting to speculate that this recombination cannot be catalyzed by RAD51, and has substrate requirements more compatible with microhomologous recombination. The second reason for suggesting that VSG switching may not be a canonical form of RAD51dependent recombination arises from examination of the roles of further T. brucei homologues of genes that are known to be important contributors to homologous recombination in other organisms. Surprisingly, some such genes do not detectably influence VSG switching. Five T. brucei orthologues of eukaryotic MMR proteins have been described to date, and at least two of them (MSH2 and MLH1) function in repairing replication errors and chemical damage [132]. MMR has also been shown to be a significant constraint upon T. brucei RAD51-dependent homologous recombination [130], as has been described in other organisms. Nevertheless, MMR has no noticeable role in regulating T. brucei antigenic variation [130], despite the heavily reliance of the process upon homologous recombination. Similarly, T. brucei MRE11 has been shown to be important in catalysing the recombination of transformed DNA constructs in the parasite genome, compatible with its suggested recombination functions in other organisms [133], but has no

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influence on VSG switching [134]. These potentially conflicting data may be explained by inadequacies in the available assays for VSG switching. Alternatively, they may point to VSG switching being predominantly an unusual form of RAD51-dependent homologous recombination, bypassing many of the factors that would be expected to exert an influence on it. However, it may equally be that each gene or repair complex, such as MMR, has novelties in T. brucei relative to other organisms which means that their functions cannot be extrapolated to VSG switching. Rad51 remains the only T. brucei gene to date that has been shown to directly catalyze or regulate VSG switching, either by recombinational or transcriptional means [135]. This demonstrates that, despite extensive molecular descriptions of how T. brucei VSG switching is executed, our understanding of the genetic components of the process, and its biochemical details, are in their infancy. Some studies suggested that VSG point mutations may be associated with the gene conversion form of VSG switching [136–138], and perhaps also with mosaic VSG formation [57]. The former has been interpreted as implying that a ‘mutator’ DNA polymerase, or an RNA intermediate, may be involved [136,139], adding another potential layer of genetic variability which may contribute to immune evasion. However, the involvement of a reverse transcriptase step in VSG gene conversion seems incompatible with the role of RAD51 and homologous recombination in the reaction, and it has been questioned whether small scale changes in VSG epitopes can provide evasion from the host specific immune response [140]. No evidence has been provided that differences in MMR efficiency occur in different T. brucei sub-species or strains (or indeed different species of African Trypanosome), analogous to the findings in T. cruzi (above). In fact, preliminary comparison of MSH2 alleles from different T. brucei isolates does not reveal any functional polymorphisms (R. Barnes, R. McCulloch, unpublished). Nevertheless, considerable

gossypii (AAS54591), Tetrahymena thermophila (AAC39117), Candida glabrata (CAG60427), Arabidopsis thaliana (NP_568402), Entamoeba histolytica (AAP35107), Physcomitrella patens (CAC82996), Drosophila melanogaster (Q27297), Plasmodium berghei (CAH95058), Plasmodium yoelii yoelii (EAA15553), Plasmodium falciparum (AAN76809), Chlamydia trachomatis (P48287), Rhodospirillum rubrum (ZP_00270420), Lactobacillus paraplantarum (CAC44773), Escherichia coli (P03017), Salmonella ente´rica (AAV78544), Vibrio cholerae (CAH57124), Haemophilus influenzae (NP_438757).

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variation in the size of T. brucei chromosome homologues, both within the same cell and between strains and sub-species, has been described [50]. The basis of this genetic variability, and its potential relationship with antigenic variation, are beginning to be understood [141,142], but its consequences in terms of genetic exchange between T. brucei strains and sub-species, and potential phenotypic changes, remain to be determined.

6. Conclusions In both T. cruzi and T. brucei, we are beginning to understand the profound influence that genes involved in DNA metabolism, or at least specifically DNA repair and recombination, have on growth and infectivity. Nevertheless, the picture is incomplete and it is clear that the roles of several other genes involved in aspects of DNA metabolism relevant to the generation of sequence variability need to be investigated. Among those, the newly characterized family of DNA polymerases that present a low fidelity of DNA replication and are likely associated with mutator phenotypes in cancer cells and during immunoglobulin class switch might be investigated [143–147]. If T. cruzi and T. brucei have equivalents of these Y-family DNA polymerases, they might be also contribute to generating the antigenic diversity that allows the parasite to escape the immune system. Beyond this, it will be valuable to explore if the rates and profiles of genetic variability are uniform around the genomes of these parasites. As in bacteria, it may suit parasites to have so-called ‘contingency’ loci [13], which are able to mutate at higher than background rates. With the completion of the ‘Tri-Tryp’ genome project this, and other, interesting questions can soon be addressed.

Acknowledgments We are indebted to Dr. Carlos Menck (University of Sa˜o Paulo) for critical reading of the manuscript and valuable suggestions. Work in Carlos Renato Machado and Santuza M.R. Teixeira laboratories is supported by grants from Conselho Nacional de Pesquisas (CNPq), Comissa˜o de Aperfeic¸oameno de

Pessoal de Nı´vel Superior (CAPES/COFECUB), World Health Organization and Fundac¸a˜o de amparo a pesquisa de Minas Gerais (FAPEMIG). Luiz Augusto-Pinto received a doctoral fellowship from CNPq. Richard McCulloch is a Royal Society University Research Fellow, and work in his lab has been supported by the Wellcome Trust and the Medical Research Council. Figs. 1 and 2 are adapted from Trends in Parasitology, Vol. 20, Issue 3, Richard McCulloch, Antigenic variation in African Trypanosomes: monitoring progress, Pages 117–121, Copyright (2004), with permission from Elsevier.

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