Parasitism and chromosome dynamics in protozoan parasites: is there a connection?

MOLECULAR BIOCHEMICAL PARASITOLOGY

ELSEVIER

Molecular

and Biochemical

Parasitology

70 (1995) l-8

Minireview

Parasitism and chromosome dynamics in protozoan parasites: is there a connection? Michael Lanzer a,*, Katja Fischer a, Sylvie M. Le Blancq b a Zentrum fir Infektionsforschung, Riintgenring 11, D-97070 Wiirzburg, Germany b Division of Tropical Medicine, School of Public Health, Columbia University, 630 West 168th Street, New York, NY 10032, USA Received 12 October 1994; accepted 23 January

1995

Abstract Genomic plasticity is a hallmark of many protozoan parasites, including Plasmodium spp, Trypanosoma spp, Leishmania and Giardia Zambfia. Strikingly, there is a common theme regarding the structural basis of this karyotype variability. Chromosomes are compartmentalized into conserved central domains and polymorphic chromosome ends. Since antigenencoding genes frequently reside in telomere-proximal domains, it is tempting to speculate that the genetic flexibility of chromosome ends has been recruited as a tool in immune evasion strategies by some parasitic protozoa. ssp

Keywords:

Genomic

plasticity;

Genome organisation;

Protozoon;

Plasmodiumfalciparum; GiardiaZamblia;Trypanosomabrucei

1. Introduction Many parasitic protozoa exhibit considerable genomic plasticity that often extends from allelic variation of single genes to polymorphisms of entire chromosomes [l-8]. This phenomenon is seen in organisms that share no phylogenetic relationship. This superficial similarity could indicate an association between genomic plasticity and the parasitic mode of life, i.e., immune evasion, either directly, as in the programmed gene rearrangements of the variable surface glycoprotein (VSG) genes in Trypanosoma brucei, or indirectly as a byproduct of a

* Corresponding 571-954.

author. Tel.: (49-931)

572-131;

Fax: (49-931)

0166-6851/95/$09.50 0 1995 Elsevier Science B.V. All rights reserved SSDI 0166-6851(95)00021-6

hyperactive recombination machinery that generates antigen diversity. In this review, we present a summary of our current knowledge of genome organization in Plasmodium falciparum, Giardia lamblia and Trypanosoma brucei, as instructive examples in a discussion of genomic variability. Space constraints preclude consideration of other protozoan parasites.

2. Chromosomal polymorphisms chromosome ends

are located

at

Chromosome organization in several protozoan parasites seems to be guided by the same blueprints. Recent studies suggest a chromosome compartmentalization into conserved central domains and polymorphic chromosome ends. A well-studied example

2

M. Lanzer et al. /Molecular

and Biochemical Parasitology

are chromosomes of the human malaria parasite P. fulciparum. Transmitted to humans by the bite of infected Anopheles mosquitos, P. falciparum propagates by schizogony within hepatocytes and erythrocytes. The life cycle includes obligatory sexual reproduction with meiosis occurring shortly after zygote formation in the mosquito and prior to transmission. The 30-Mb haploid P. fulciparum genome comprises 14 chromosomes, as determined by both pulse-field gel electrophoresis and by counting the number of kinetochores (attachment sites of the spindle apparatus) in electron micrographs [9,10]. Comparison of homologous chromosomes from different geographic isolates has shown that all 14 P. fulciparum chromosomes display extensive clone-dependent size polymorphisms of up to 15% in both field isolates and parasites cultured in vitro [3,4,11,12]. While some genetic variability results from sexual recombination during meiosis, the extensive chromosomal polymorphism observed originates spontaneously during mitotic growth (reviewed in Ref. 13). To address the structural and possibly biological ramification of chromosomal polymorphisms, several P. falciparum chromosomes have been cloned in their entirety as arrays of yeast artificial chromosomes (YAC clones) [14,15]. Analysis of chromosome 2 and 4 contig maps has revealed that polymorphisms are confined to chromosome ends, defining approx. 100 kb at either end of a chromosome as genetically variable [14,15]. The central chromosome domains, by contrast, are conserved. A similar chromosomal compartmentalization is also apparent in G. lumblia (Ref. 16 and Hou, Le Blancq, E, Chu and Lee, data not shown), a parasite of the upper intestinal tract of mammals, including humans, where it causes a broad clinical spectrum of symptoms ranging from diarrhea to malabsorption [17]. The life cycle consists of vegetative trophozoites and infective cysts, and meiosis has not been observed. Each trophozoite contains two transcriptionally active nuclei that contain the same amount of DNA [Ml. Genome size and chromosome number have not been determined unequivocally, with estimates of the haploid genome ranging from 12 to 80 Mb [19,20] and chromosome number ranging from 8 to 50 per trophozoite [l&21]. Consequently, the

70 (1995) l-8

ploidy has not been determined. Recent data suggests polyploidy, perhaps tetraploidy, with some aneuploid components (Refs. 22,23 and Hou, Le Blancq, E, Chu and Lee, data not shown). Clinical and field isolates of G. lamblia show striking karyotype heterogeneity in terms of chromosome number and size [5,6]. Furthermore, chromosome rearrangements occur spontaneously at a rate of approx. 1% per cell per division cycle in vitro during mitotic growth 1251. The chromosomes that have been studied extensively in G. lamblia are those that encode the ribosoma1 RNA (rRNA) genes. The ends of these chromosomes are subject to extensive and frequent polymorphisms, while the central chromosomal domains are conserved (Refs. 16,25 and Hou, Le Blancq, E, Chu and Lee, data not shown). The rRNA genes are present as short arrays close to the telomeres of these chromosomes [26-281, and changes in their copy number contribute to the size polymorphisms (Ref. 16 and Hou, Le Blancq, E, Chu and Lee, data not shown). Although transcription of the telomeric rRNA genes has not been demonstrated directly, the telomeric location of all detectable rRNA genes strongly suggests that at least part of the polymorphic subtelomeric domains are transcriptionally active [26,27]. The transcription of polymorphic telomerit domains, namely the VSG gene expression sites, has been demonstrated directly in T. brucei [29-311 (see below). By contrast, P. falciparum chromosome ends are transcriptionally-inactive as determined by both Northern analysis and transcription mapping of telomerit YAC clones [14,29]. They are organized as arrays of repetitive sequence elements that extend 85 kb, on average, from the telomere [32]. This terminal region, designated the ‘subtelomeric domain’ is common to all wild-type P. falciparum chromosomes. The structural organization of chromosomes into poylmorphic ends and conserved central domains has also been found in yeast, as well as other protozoa such as Leishmania infantum, Plasmodium berghei and T. brucei [1,33-371. T. brucei, the etiologic agent of sleeping sickness in humans and cattle, is a kinetoplastid flagellated protozoa that propagates in the blood stream of its mammalian hosts, and the midgut of the tsetse fly. Sexual reproduction occurs in the fly, but it is not required for transmission. T.

M. Laruer et al. /Molecular

and Biochemical Parasitology

brucei is diploid, with a genome of 70 Mb that comprises more than 100 minichromosomes, ranging in size from 50 to 150 kb, and at least 18 large chromosomes up to 5.7 Mb [38,39]. Homologous chromosomes in T. brucei differ in size by about 20%, but some of the chromosome pairs show a 2-fold size difference [39]. Karyotype differences are seen among different field isolates and clonal populations in the laboratory [1,40]. The subtelomeric regions of T. brucei chromosomes are polymorphic and they contain repetitive elements [1,37].

3. Different phisms

mechanisms

contribute

to polymor-

Several different types of polymorphisms are responsible for the genetic variability of chromosome ends. A major source of polymorphisms are frequent expansion and contraction of repetitive subtelomeric sequence elements (Refs. 11,14,33,34,41 and Hou, Le Blancq, E, Chu and Lee, data not shown). Such rearrangements could be the result of sister chromatid interactions or ectopic recombination between heterologous chromosomes during mitosis, or slippage of the DNA polymerase during replication. The function of subtelomeric sequences is still obscure in protozoan parasites. A potential role in chromosome pairing has been proposed for plasmodial subtelomeric sequences (see below). In yeast the subtelomeric elements, X and Y’, serve as (i> origins of replication and (ii) buffers between telomere repeat sequences and transcribed regions, thereby preventing the silencing of telomere proximal genes [35,42]. Whether subtelomeric repeats have equivalent functions in protozoa is not known, although it is an intriguing possibility. Besides variability of subtelomeric domains, other types of polymorphisms such as truncations are evident in G. lamblia and malaria parasites (Refs. 26,28,43-49 and Hou, Le Blancq, E, Chu and Lee, data not shown). Chromosomal truncations have been extensively studied in P. falciparum since these events occur spontaneously during mitotic propagation, affect all 14 chromosomes, and are frequently observed in parasites cultured in vitro [43-491. They are initiated by double-strand breaks, and while the terminal segment is lost, the residual fragment that

3

70 (1995) l-8

contains the centromere is stabilized by the de novo addition of telomere repeat sequences 1441. A hot spot for double-strand chromosome breakages has recently been mapped to the P. falciparum KAHRP gene on chromosome 2, 90 kb distal from the telomere [50]. The distance between the breakpoints showed a periodicity identical to the P. falciparum nucleosome repeat unit of 156 + 4 bp [50]. Nucleosome core particles are 148 k 5 bp in size, internucleosomal linkers 8 13 bp [50,51]. Comparison of nucleosomal organization with chromosome breakpoints revealed that mitotic double-strand breaks occur in nucleosome linkers of transcriptionally-active domains [50]. Thus, chromosome breakages in P. falciparum may be initiated by an endolytical activity that requires open chromatin for cleavage. Transcriptional activity results in disruption of the nucleosome structure and depletion of histone Hl, which extends beyond the core particle along the DNA into the intemucleosomal linker [52]. Comparison of several chromosomal breakage sites in P. falciparum has revealed a conserved CA dinucleotide immediately preceding the breakpoint, indicating a potential cleavage preference [4446,48,50]. Alternatively, the conserved CA dinucleotide could function in healing broken chromosomes. Since P. falciparum telomere repeats are composed of GGGTTCA as well as GGGTITA [53], a CA dinucleotide could function as a necessary signal for the de novo addition of telomere repeat sequences by a P. falciparum telomerase. This model is consistent with the observation that other eukaryotic telomerases possess a 3’ to 5’ exonuclease activity and are capable of elongating non-telomeric primers [54-561.

4. Chromosomal variation

polymorphism

and

autigenic

Pathogens have acquired effective mechanisms that enable them to evade the host’s immune attack. One of the principal evasion mechanisms is antigenic variation. It has been observed in several protozoan pathogens including T. brucei, P. falciparum and G. lamblia [57-631. T. brucei is a paradigm for the potential of DNA rearrangements to effect changes in gene expression.

4

M. Lancer et al. /Molecular

and Biochemical Parasitology

Antigenic variation in T. brucei involves the serial expression of different VSGs that compose the surface coat of the trypanosome and bear the brunt of the immune attack from the host [37,57,58,64]. Each cell contains a repertoire of more than 1000 different VSG genes, but only one is expressed at a time. The transcriptionally active VSG gene is located at a telomeric expression site, whereas the other VSG genes are situated at silent basic copy sites that are located at chromosome-internal positions, or at inactive telomeres (reviewed in Refs. 37,641. There are several expression sites and the VSG coat is changed by either activating a previously silent telomeric expression site or by replacing the VSG gene in the active expression site. These latter events comprise programmed DNA rearrangements [64]. The regulatory mechanisms that control VSG expression site activation and inactivation are not known. One potential mechanism under investigation is a putative trypanosomal telomere position effect similar to telomere silencing in yeast [65], where the expression of genes placed close to telomere repeats is reversibly repressed [42]. G. lamblia also possesses a repertoire of variantspecific surface protein (VSP) genes, but the mechanism(s) that controls their differential expression is still unclear [22,63,66]. The VSP genes do not appear to be expressed from telomeric expression sites, and DNA rearrangements associated with the VSP genes have not been linked directly to the control of their expression [22,67]. Antigenic variation in P. falciparum involves a class of proteins called PfEMPl that are present on the surface of parasitized erythrocytes [59,60]. Whether DNA rearrangements play a role in the control of PjEMPl gene expression or antigenic variation, respectively, is not yet known. It is clear, however, that chromosomal truncations affect the expression of other antigen encoding genes, such as KUZRP, RESA, HRPZZ, PFll-1 on chromosomes 2, 1, 8 and 10, respectively [43-491. The truncations extend into the coding regions of these genes, thereby abrogating their expression and resulting in mutants with altered antigenic phenotypes. However, these mutations tend to be deleterious, as best exemplified by KAHRP- mutants, and may not play a constructive role in vivo. The KAHRP gene product is a component of

70 (1995) l-8

‘knob’ structures on the surface of parasitized erythrocytes that are implicated in cytoadherence to endothelial cells [68,69]. Chromosomal breakages disrupting the KAHRP gene result in mutants that are knobless and cytoadherence-deficient [43,70]. These mutants possess a growth advantage in cell culture but are cleared by the host’s spleen in vivo [71]. Truncations of other chromosomes are also detrimental to parasite fitness and virulence, for example, a truncation of chromosome 9 abrogates gametogenesis [49]. It is also unlikely that relocation of telomere repeats due to chromosomal rearrangements plays a relevant role in plasmodial gene regulation. Chromosome 2 truncations that result in the juxtaposition of telomeres to the GLMP gene had no obvious effect on the expression of this gene [51]. In the mutant and in the wild-type, the GLARE gene is expressed during the ring stage. A telomere position effect has thus far not been observed in P. falciparum. Thus, there is as yet no evidence to support a general association between chromosomal polymorphisms and antigenic variation per se in parasitic protozoa. It is important to note that organisms that do not exhibit antigenic variation, such as Leishmania ssp., have similarly plastic genomes [36,72,73]. Furthermore, chromosomal polymorphism, indeed genomic plasticity, is not restricted to parasites. Other unicellular eukaryotes such as yeast also demonstrate karyotype variability [74,75], and free-living ciliated protozoa undergo extensive genomic reorganization during their life cycle [76]. Notwithstanding this, the genetic variability observed is indicative of a hyperactive recombination machinery that confers flexibility on the genome. Such flexibility can facilitate survival in the face of environmental challenges. For example, gross DNA rearrangements are observed when protozoan parasites are exposed to drugs. In P. falciparum resistance to mefloquine arises by amplification of the Pfidrl gene locus, since its gene products seems to mediate mefloquine export [77]. Conversely, under chloroquine selection, the copy number of Z’fmdrl genes is reduced [78], suggesting that the Pfmdrl gene product is involved in the import of this antimalarial drug [79]. Resistance to pyrimethamine in P. chabaudi, and methotrexate in Leishmania major results from amplification of the dihydrofolate reduc-

M. Lamer et al. /Molecular

and Biochemical Parasitology 70 (I 995) l-8

tase-thymidylate synthase (DHFR-TS) gene [SO,Sl]. Chromosomal rearrangements may also occur when parasites are subject to nutrient stress. In L. tarentoZae, nutrient shock can induce chromosomal truncations or appearance of an additional chromosome

[a. 5. Subtelomeric domains facilitate ectopic recombination It has been proposed that exchange of VSG genes in T. brucei is mediated by ectopic recombination, i.e., pairing of non-homologous chromosomes followed by duplicative transposition via gene conversion and/ or reciprocal recombination [37,64]. Pairing between homologous, as well as heterologous chromosomes preferentially initiates at the telomere during both meiosis and mitosis [83-851. Telomeres from different T. brucei chromosomes form distinct clusters within the nucleus [86]. The unusually high number of chromosomes, combined with a tight spatial organization within the nucleus, could facilitate ectopic recombination during mitosis. Repetitive sequences at chromosome ends also seem to promote chromosome pairing [85]. The organization of P. falciparum subtelomeric regions as long arrays of repetitive sequences may have evolved to facilitate this process. Genetic data from P. falciparum suggest a high frequency of recombination and crossing-over events close to chromosome ends [87]. Interestingly, many P. falciparum antigens known map to subtelomere-proximal domains [88]. This location would expose these genes to a high frequency of recombination, enhancing the emergence of new antigenic phenotypes. In this context it is worth mentioning that several antigens comprise families, such as HZWI/HZWZZ, ZESA /ZEYA-2, GBP/ GBP-H and Pj332 /Pfll-1 (Refs. 10,48,88 and K. Hinterberger, D. Mattei and A. Scherf, personal communication). The individual members are located in subtelomere-proximal regions of different chromosomes [88]. These gene families may have arisen by duplication of ancestral genes via ectopic recombination. In contrast, housekeeping genes residing in central chromosomal domains would be subject to far fewer recombination, leading to greater conservation.

5

There is further structural and genetic evidence in support of this model. The P. falciparum clone HB3 is aneuploid for a 200-kb segment that contains several genes such as RESA-2 and P@32 [89]. This segment is present on two chromosomes, chromosome 11, its original location and on chromosome 13 as a result of a duplicative translocation. When the HB3 strain was crossed with the wild-type clone Dd2, progeny emerged in which the HB3 chromosome 13 segment was transposed back to its original location on chromosome 11 [89]. Thus, heterologous chromosomes can pair, facilitating illegitimate recombination that may then result in the exchange and possibly duplication of subtelomere-proximal genes. It is here where a hyperactive recombinatory machinery could bestow an advantage on the parasite. Translocations are indeed frequently observed in all malaria species [14,15,901. Once a gene family has emerged, the individual members can diverge independently, thereby expanding the parasite’s antigenie profile.

6. Conclusions

Several features of genome organization are held in common by various protozoan parasites, namely: compartmentalisation of the chromosome into conserved core domains and variable chromosome ends; very high frequencies of chromosomal rearrangements and the contribution of different mechanisms to chromosomal polymorphisms. Whether genomic plasticity contributes to the parasitic mode of life in a generalized manner remains an open question. Specific examples, such as programmed gene rearrangements in T. brucei, certainly demonstrate its potential. The ability to clone and sequence entire chromosomes will facilitate the physical characterization of various protozoan genomes. This and recent successes in transfection, added to other molecular tools, will allow detailed investigation of the mechanisms that control antigenic variation and chromosome rearrangements. Such studies should show whether or not there is any causal relationship between chromosome structure and parasitism.

M. Lanzer et al. /Molecular

6

and Biochemical Parasitology 70 (1995) 1-8

Acknowledgements This work was supported by the terium ftir Forschung und Technologie publik Deutschland (M.L. and K.F.) grant AI26497-06 in the international Infectious Disease Research Program

Bundesminisder Bundesreand by N.I.H. Collaborative to S.L.B.

References [ll Van der Ploeg, L.H.T.,

Cornelissen, A.W.C.A., Michels, P.A.M. and Borst, P. (1984) Chromosome rearrangements in frypanosoma brucei. Cell 39, 213-221. [21 Holmes Giannini, S., Schittini, M., Keithly, J.S., Warburton P.W., Cantor, CR. and Van der Ploeg, L.H.T. (19861 Karyotype analysis of Leishmania species and its use in classification and clinical diagnosis. Science 232, 762-765. [31 Kemp D.J., Corcoran, L.M., Coppel, R.L., Stahl, H.D., Bianco, A.E., Brown, G.V. and Anders, R.F. (1985) Size variations in chromosomes from independent cultured isolates of Plasmodium falciparum. Nature 315, 347-350. T., Ver141 Van der Ploeg, L.H.T., Smits, M., Ponnudurai, meulen, A., Meuwissen, J.H.E.Th. and Iangsley, G. (1985) Chromosome-sized DNA molecules of Plasmodium falciparum. Science 229, 658-661. [51Korman, S.H., Le Blancq, S.M., Deckelbaum, R.J. and Van der Ploeg, L.H.T. (1992) Investigation of human giardiasis by karyotype analysis. J. Clin. Invest. 89, 1725-1733. 161Sarafis, K. and Isaac-Renton, J. (1993) Pulsed-field gel electrophoresis as a method of biotyping of Giardia lamblia. Am. J. Trop. Med. Hyg. 48, 134-144. [71 Van der Ploeg, L.H.T., Schwartz, D.C., Cantor, CR. and Borst, P. (1984) Antigenic variation in Trypanosoma brucei analyzed by electrophoretic separation of chromosome sized DNA molecules. Cell 37, 77-84. [8] Sharkey, A., Langsley, G., Patarapotikul, J., MercereauPuijalon, O., McLean, A.P. and Walliker, D. (1988) Chromosome size variation in the malaria parasites of rodents, Plasmodium chabaudi. Mol. Biochem. Parasitol. 22, 47-54. [9] Prensier, G. and Slomianny, C. (1986) The karyotype of Plasmodium falciparum determined by ultrastructural serial sectioning and 3D reconstruction. J. Parasitol. 72, 731-736. [lo] Wellems T.E., Walliker, D., Smith, CL., do Rosario, V.E., Maloy, W.L., Howard, R.J., Carter, R. and McCutchan, T.F. (1987) A histidine-rich protein gene marks a linkage group favored strongly in a genetic cross of Plasmodium falciparum. Cell 49, 633-642. (111 Corcoran, L.M., Forsyth, K.P., Bianco, A.E., Brown, G.V. and Kemp, D.J. (1986) Chromosome size polymorphisms in Plasmodium faiciparum can involve deletions and are frequent in natural parasite populations. Cell 44, 87-95. [12] Biggs, B.A., Kemp, D.J. and Brown, G.V. (19891 Subtelomerit chromosome deletions in field isolates of Plasmodium falciparum and their relationship to loss of cytoadherence in vitro. Proc. Natl. Acad. Sci. USA 86, 2428-2432.

031 Walliker, D. (1989) Genetic recombination in malaria parasites. Exp. Parasitol. 69, 303-309. [I41 Lanzer, M., De Bruin, D. and Ravetch, J.V. (1993) Transcriptional differences in polymorphic and conserved domains of a complete cloned P. falciparum chromosome. Nature 361, 654-657. [I51 Rubio, J.P., Triglia, T., Kemp, D.J., De Bruin, D., Ravetch, J.V. and Cowman, A.F. (In press) A YAC contig map of Plasmodium falciparum chromosome 4: Transcriptional analysis and characterization of a DNA amplification between two recently separated isolates. Genomics. I161Adam, R.D. (1992) Chromosome-size variation in Giardia lamblia: the role of rDNA repeats. Nucleic Acids Res. 20, 3057-3061. D71 Heyworth, M.F. (1992) Immunology of Giardia and Cryp tosporidium infections. J. Infect. Dis. 199, 465-472. W31Kabnick, KS. and Peattie, D.A. (1990) In situ analyses reveal that the two nuclei of Giardia lamblia are equivalent. J. Cell Sci. 95, 353-360. 1191 Fan, J., Korman, S.H., Cantor, C.R. and Smith, C.L. (1991) Giardia lamblia: haploid genome size determined by pulsed field gel electrophoresis is less than 12 Mb. Nucleic Acids Res. 19, 1905-1908. 1201 Boothroyd, J.C., Wang, A., Campbell, D.A. and Wang, CC. (1987) An unusually compact ribosomal DNA repeat in the protozoan Giardia lamblia. Nucleic Acids Res. 10, 40654084. El1 Adam, R.D., Nash, T.E. and Wellens, T.E. (1988) The Giardia lamblia trophozoite contains sets of closely related chromosomes. Nucleic Acids Res. 16, 4555-4567. t221 Yang, Y. and Adam, R.D. (1994) Allele-specific expression of a variant-specific surface protein (VSP) of Giardia lamblia. Nucleic Acids Res. 22, 2102-2108. [231 Upcroft, J.A., Healey, A. and Upcroft, P. (1993) Chromosoma1 duplication in Giardia duodenalis. Int. J. Parasitol. 23, 609-616. [241 Le Blancq, SM., Korman, H.S. and Van der Ploeg, L.H.T. (1992) Spontaneous chromosome rearrangements in the protozoan Giardia lamblia: estimation of mutation rates. Nucleic Acids Res. 17, 4539-4545. [25] Chen, N., Upcroft, J.A. and Upcroft, P. (1994) Physical map of a 2 Mb chromosome of the intestinal protozoan parasite G. duodenalis. Chromosome Res. 2, 307-313. [26] Le Blancq, S.M., Kase, R.S. and Van der Ploeg, L.H.T. (1991) Analysis of a Giardia lamblia rRNA encoding telomere with [TAGGG], as the telomere repeat. Nucleic Acids Res. 19, 5790. [27] Le Blancq, S.M., Korman, S.H. and Van der Ploeg, L.H.T. (1991) Frequent rearrangements of rRNA-encoding chromosomes in Giardia lamblia. Nucleic Acids Res. 19, 44054412. [28] Adam, R.D., Nash, T.E. and Wellems, T.E. (1991) Telomeric location of Giardia rDNA genes. Mol. Cell. Biol. 11, 33263330. [29] Pays, E., Lheureux, M. and Steinert, M. (1981) The expression-linked copy of surface antigen gene in Trypanosoma is probably the one transcribed. Nature 292, 265-267.

M. Lanzer et al. /Molecular

7

and Biochemical Parasitology 70 (I 995) I-8

[30] Hoeijmakers, J.H.J., Frasch, A.C.C., Bemards, A., Borst, P. and Cross, G.A.M. (1980) Novel expression-linked copies of the genes for variant surface antigens in trypanosomes. Nature 284, 78-80. [31] Kooter, J.M., Van der Spek, H.J., Wagter, R., d’oliviera, C.E., Van der Hoeven, F., Johnson, P.J. and Borst, P. (1987) The anatomy and transcription of a telomeric expression site for variant-specific surface antigens in T. brucei. Cell 51. 261-272. [32] de Bruin, D., Lanzer, M. and Ravetch, J.V. (1994) The polymorphic subtelomeric regions of Plasmodium falciparum chromosomes contain arrays of repetitive sequence elements. Proc. Natl. Acad. Sci. USA 91, 619-623. [33] Ponzi, M., Pace, T., Dore, E., Picci, L. and Frontali, C. (1992) Extensive turnover of telomeric DNA at a Plastnodium berghei chromosomal extremity marked by rare recombinational event. Nucleic Acids Res. 17, 4491-4497. [34] Pace. T., Ponzi, M., Dore, E. and Frontali, C. (1987) Telomerit motifs are present in a highly repetitive element in the Plasmodium berghei genome. Mol. Biochem. Parasitol. 24, 193-202. [35] Biessmann, H. and Mason J.M. (1992) Genetics and molecular biology of telomeres. Adv. Genet. 30, 185-249. 1361 Blaineau, C, Bastian, P.. Rioux, J.-A.. Roizes, G. and Pays, M. (1991) Long-range restriction maps of size-variable homologous chromosomes in Leishmania infantum. Mol. Biochem. Parasitol. 46, 293-302 [37] Van der Ploeg, L.H.T. (1990) Antigenic variation in African trypanosomes: genetic recombination and transcriptional control of VSG genes. In: Frontiers in Molecular Biology: Genome Rearrangements and Amplification (Hames, B.D. and Glover, D.M.. eds.), pp. 51-97. IRL Press, New York, NY. [38] Van der Ploeg, L.H.T., Smith, C.L., Polvere, R.I. and Gottesdiener, K.M. (1989) Improved separation of chromosomesized DNA from Trypanosoma brucei stock 427-60. Nucleic Acids Res. 17, 3217-3227. [39] Gottesdiener, K., Garcia-Anoveros, J., Lee, M.G.-S. and Van der Ploeg, L.H.T. (1990) Chromosome organization of the protozoan Trypanosoma brucei. Mol. Cell. Biol. 10, 60796083. [40] Gibson, W. and Garside, L. (1991) Genetic exchange in Trypanosoma brucei: variable chromosomal location of housekeeping genes in different trypanosome stocks. Mol. Biochem. Parasitol. 39, 9-22. [41] Corcoran, L.M., Thompson, J.K., Walliker, D. and Kemp, D.J. ( 1988) Homologous recombination within subtelomeric repeat sequences generates chromosome size polymorphism in P. falciparum. Cell 53, 807-813. [42] Gottschling, D.E., Aparicio, O.M., Billington, B.I. and Zakian. V.A. (1990) Position effect at S. cereuisiae telomeres: reversible repression of Pol 11transcription. Cell 63, 751-762. [43] Pologe, L.G. and Ravetch, J.V. (1986) A chromosomal rearrangement in a P. falciparum histidine-rich protein gene is associated with the knobless phenotype. Nature 322, 474477. [44] Pologe, L.G. and Ravetch, J.V. (1988) Large deletions result

from breakage [45]

[46]

[47]

[48]

[49]

[50]

[51]

[52]

[53]

[54]

[55]

[56]

[57] [58]

and healing

of P. falciparum chromosomes.

Cell 55, 869-874. Pologe, L.G., De Bruin, D. and Ravetch, J.V. (1990) A and T homopolymeric stretches mediate a DNA inversion in Plasmodium falciparum which results in the loss of gene expression. Mol. Cell. Biol. 10, 3243-3246. Scherf, A. and Mattei, D. (1992) Cloning and characterization of chromosome breakpoints of Plasmodium falciparum: breakage and new telomere formation occurs frequently and randomly in subtelomeric genes. Nucleic Acids Res. 20, 1491-1496. Shirley, M.W., Biggs, B.A., Forsyth, K.P., Brown, H.J., Tompson, J.K., Brown, G.V. and Kemp, D.J. (1990) Chromosome 9 from independent clones and isolates of Plasmodium falciparum undergoes subtelomeric deletions with similar breakpoints in vitro. Mol. Biochem. Parasitol. 40, 137146. Scherf, A., Carter, R., Peterson, C., Alano, P., Nelson, R., Aikawa, M., Mattei, D., Pereira da Silva, L. and Leech, J. (1992) Gene inactivation of PFIl-1 of Plasmodium falciparum by chromosome breakage and healing: identification of a gametocyte-specific protein with a potential role in gametogenesis. EMBO J. 11, 2293-2301. Day, K.P.. Karamalis, F., Thompson, J., Barnes. D.A., Peterson, C., Brown, H., Brown, G.V. and Kemp, D.J. (1993) Genes necessary for expression of a virulence determinant and for transmission of Plasmodium falciparum are located on a 0.3 megabase region of chromosome 9. Proc. Natl. Acad. Sci. USA 90, 8292-8296. Lanzer, M., Wertheimer, S.P., De Bruin, D. and Ravetch, J.V. (1994) Chromatin structure determines the sites of chromosome breakages in Plasmodium falciparum. Nucleic Acids Res. 15, 3099-3103. Lanzer, M., De Bruin, D., Wertheimer, S.P. and Ravetch, J.V. (1994) Transcriptional and nucleosomal characterization of a subtelomeric gene cluster flanking a site of chromosoma1 rearrangements in Plasmodium falciparum. Nucleic Acids Res. 22, 4176-4182. Clark, D.J. and Felsenfeld, G. (1992) A nucleosome is transferred out of the path of a transcribing polymerase. Cell 71, 11-22. Vernick, K.D. and McCutchan, T.F. (1988) Sequence and structure of a Plasmodium falciparum telomere. Mol. Biochem. Parasitol. 28, 85-94. Collins, K. and Greider. C.W. (1993) Tetrahymena telomerase catalyzes nucleolytic cleavage and nonprocessive elongation. Genes Dev. 7. 1364-1376. Harrington, L.A. and Greider, C.W. (1991) Telomerase primer specificity and chromosome healing. Nature 353, 451-454. Morin, G.B. (1991) Recognition of a chromosome truncation site associated with cu-thalassaemia by human telomerase. Nature 353, 454-456. Cross, G.A.M. (1978) Antigenic variation in trypanosomes. Proc. R. Sot. London Ser. B202, 55-72. Vickerman, K. (1978) Antigenic variation in trypanosomes. Nature 273, 613-617.

8

M. Laruer et al. /Molecular

and Biochemical Parasitology 70 (1995) l-8

1591 Roberts, D.J., Craig, A.G., Berendt, A.R., Pinches, R., Nash, G., Marsh, K. and Newbold, C.I. (1992) Rapid switching to multiple antigenic and adhesive phenotypes in malaria. Nature 357, 689-692. [60] Biggs, B.A., Gooze, L., Wycherley, K., Wollish, W., Southwell, B., Leech, J.H. and Brown, G.V. (1991) Antigenic variation in Plasmodium falciparum. Proc. Natl. Acad. Sci. USA 88, 9171-9174. [61] Nash, T.E., Aggarwal, A., Adam, R.D., Conrad, J.T. and Merritt, J.W. (1988) Antigenic variation in .Giardia lamblia. J. Immunol. 141, 636-641. [62] Aggarwal, A. and Nash, T.E. (1988) Antigenic variation of Giardia lamblia in vivo. Infect. Immun. 56, 1420-1423. [63] Nash, T. (1992) Surface antigen variability and variation in Giardia lamblia. Parasitol. Today 7, 229-235. [64] Borst, P. and Greaves, D. (1987) Programmed gene rearrangements altering gene expression. Science 235, 658-667. [65] Van der Werf, A., Van Assel, S., Aerts, D., Steinert, M. and Pays, E. (1990) Telomere interaction may condition the programming of antigen expression in Typanosoma brucei. EMBO J. 9, 1035-1040. [66] Adam, R.D., Yang, Y.M. and Nash, T.E. (1992) The cysteine-rich protein gene family of Giardia lamblia: loss of the CRP170 gene in an antigenic variant. Mol. Cell. Biol. 12, 1194-1201. [67] Adam, R.D., Aggarwal, A., Lal, A.A., De la Cruz, V.F., McCutchan, T. and Nash, T.E. (1988) Antigenic variation of a cysteine-rich protein in Giardia lamblia. J. Exp. Med. 167, 109-118. [68] Aikawa, M., Rabbage, J.R. and Wellde, B.T. (1972) Junctional apparatus in erythrocytes infected with malaria parasites. 2. Zellforsch. Mikrosk. Anat. 124, 722-727. 1691 Raventos-Suarez, C., Kaul, D.K., Macaluso, F. and Nagel, R.L (1985) Membrane knobs are required for the microcirculatory obstruction induced by Plasmodium falciparum infected erythrocytes. Proc. Natl. Acad. Sci. USA 82, 38293833. [70] Udeinya, I.J., Schmidt, J.A., Aikawa, M.A., Miller, L.H. and Green, I. (1981) Falciparum malaria infected erythrocytes specifically bind to cultured human endothelial cells. Science 213,555-557. 1711 Bamwell, J.W., Howard, R.J. and Miller, L.H. (1983) Influence of the spleen on the expression of surface antigens on parasitized erythrocytes. Ciba Foundation Symposium, Vol. 94, pp. 117-132. Pitman Press, London. [72] Bishop, R.P. (1990) Extensive homologies between Leishmania donouani chromosomes of markedly different size. Mol. Biochem. Parasitol. 38, l-11. [73] Cruz, A.K., Titus, R. and Beverley, SM. (1993) Plasticity in chromosome number and testing of essential genes in Leishmania by targeting. Proc. Natl. Acad. Sci. USA 90, 15991603. [74] Gerring, S.L., Connelly, C. and Hieter, P. (1991) Positional mapping of genes by chromosome blotting and chromosome fragmentation. Methods Enzymol. 194, 57-77. [75] Carle, G.F. and Olson, M.V. (1985) An electrophoretic karyotype for yeast. Proc. Natl. Acad. Sci. USA 82, 3756-3760.

[76] Blackbum, E.H. and Karrer, K.M. (1986) Genomic reorganization in ciliated Protozoans. Annu. Rev. Genet. 20, 501-521. [77] Cowman, A.F., Galatis, D. and Thompson, J.K. (19941 Selection of mefloquine resistance in Plasmodium falciparum is linked to amplification of the Pfmdrl gene and cross-resistance to halofantrine and quinine. Proc. Natl. Acad. Sci. USA 91, 1143-1147. [78] Barnes, D.A., Foote, S.J., Galatis, D., Kemp, D.J. and Cowman, A.F. (1992) Selection for high-levels of chloroquine resistance results in deamplification of the pfmdrl gene and increased sensitivity to mefloquine in Plasmodium falciparum. EMBO J. 11,3067-3075. [79] Van Es, H H.G., Karcz, S., Chu, F., Cowman, A.F., Vidal, S. Gros, P. and Schurr, E. (1994) Expression of the plasmodial pfmdrl gene in mammalian cells is associated with increased susceptibility to chloroquine. Mol. Cell. Biol. 14, 2419-2428. [80] Cowman, A.F. and Lew, A.M. (1989) Antifolate drug selection results in duplication and rearrangement of chromosome 7 in Plasmodium chabaudi. Mol. Cell. Biol. 9, 5182-5188. [81] Ellenberger, T.E. and Beverley, SM. (1989) Multiple drug resistance and conservative amplification of the H region in Leishmania major. J. Biol. Chem. 264, 15094-15103. [82] L. Rovai, Tripp, C., Stuart, K. and Simpson, L. (1992) Recurrent polymorphisms in small chromosomes of Leishmania tarentolae after nutrient stress or subcloning. Mol. Biol. Parasitol. 50, 115-126. [83] Zickler, D., Moreau, P.J., Huynh, A.D. and Slezec, A.M. (1992) Correlation between pairing initiation sites, recombination modules and meiotic recombination in Sordasia macrospora. Genetics 132, 135-148. [84] Loidl, J., Scherthan, H., Kaback, D.B. (1994) Physical association between nonhomologous chromosomes preceds distributive disjunction in yeast. Proc. Natl. Acad. Sci. USA 91, 331. [85] Petes, T.D. and Hill, C.W. (1988) Recombination between repeated genes in microorganisms. Annu. Rev. Genet. 22, 147-168. [86] Chung, H.M., Shea, C., Fields, S., Taub, R.N. and Van der Ploeg, L.H.T. (1990) Architectural organization in the interphase nucleus of the protozoan Trypanosoma brucei: location of telomeres and minichromosomes. EMBO J. 8.2611-2619. [87] Vemick, K.D., Walliker, D. and McCutchan, T.F. (1988) Genetic hypervariability of telomere-related sequences is associated with meiosis in Plasmodium falciparum. Nucleic Acids Res. 16, 6973-6985. [88] Triglia, T., Wellems, T.E. and Kemp, D.J. (1992) Towards a high-resolution map of the Plasmodium falciparum genome. Parasitol. Today 8, 225-229. [89] Hinterberg, K., Mattei, D., Wellems, T. E. and Scherf, A. (1994) Interchromosomal exchange of a large subtelomeric segment in a Plasmodium falciparum cross. EMBO J. 13, 4174-4180. [90] Janse, C.J., Ramesar, J. and Mons, B. (1992) Chromosome translocation in Plasmodium berghei. Nucleic Acids Res. 3, 581-586.