• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Apr 25, 2006; 103(17): 6706–6711.
Published online Apr 14, 2006. doi:  10.1073/pnas.0510363103
PMCID: PMC1458945

Evidence on the chromosomal location of centromeric DNA in Plasmodium falciparum from etoposide-mediated topoisomerase-II cleavage


Centromeres are the chromosomal loci that facilitate segregation, and, in most eukaryotes, they encompass extensive regions of genomic DNA. Topoisomerase-II has been identified as a crucial regulator of segregation in a wide range of organisms and exhibits premitotic accumulation at centromeres. Consistent with this property, treatment of cells with the topoisomerase-II inhibitor etoposide promotes chromosomal cleavage at sites within centromeric DNA. In the case of the human malaria parasite Plasmodium falciparum, despite a completed genome sequence, there are no experimental data on the nature of centromeres. To address this issue, we have used etoposide-mediated topoisomerase-II cleavage as a biochemical marker to map centromeric DNA on all 14 parasite chromosomes. We find that topoisomerase-II activity is concentrated at single chromosomal loci and that cleavage sites extend over ≈10 kb. A shared feature of these topoisomerase-II cleavage sites is the presence of an extremely AT-rich (≈97%) domain with a strictly defined size limit of 2.3–2.5 kb. Repetitive arrays identified within the domains do not display interchromosomal conservation in terms of length, copy number, or sequence. These unusual properties suggest that P. falciparum chromosomes contain a class of “regional” centromere distinct from those described in other eukaryotes, including the human host.

Keywords: chromosome, malaria, segregation

Chromosome segregation ensures faithful inheritance of the genome by daughter cells. In most eukaryotes, centromeres are the chromosomal loci that facilitate this process. They are the site of assembly of the kinetochore, the nucleoprotein complex that acts as the anchor for attachment of the microtubule spindles that separate chromosome pairs and mediate their movement to the daughter nuclei. Most centromeres are “regional” entities and encompass large sections of chromosomal DNA, ranging from 0.3 to 15 Mb in species as diverse as plants, insects, and mammals (13). Even in microorganisms, centromeres can be extensive; in Schizosaccharomyces pombe, for example, they vary from 35 to 110 kb (4, 5). Unusually, in the budding yeast Saccharomyces cerevisiae, centromeres are restricted to specific 125-bp regions termed “point” centromeres that are both sufficient and necessary for spindle attachment (6, 7). In higher eukaryotes, centromeric DNA is typically comprised of arrays of short, highly repeated sequences (often 150–180 bp), interrupted by transposable elements (8, 9). In other cases, such as S. pombe, centromeres are characterized by the presence of multiple arrays of longer repeats (10). Although many organizational features of centromeric DNA are widespread, there is little sequence conservation, even between closely related species (11).

The genome sequence of Plasmodium falciparum, the parasite that causes the most serious form of malaria, has been completed (12). The sequence is 23 Mb, with 14 chromosomes ranging from 0.7 to 3.4 Mb. Transposable elements appear to be absent from the genome, and regions with properties typical of centromeric DNA have not been identified. However, it was noted that P. falciparum chromosomes contain a small, extremely AT-rich region of ≈2 kb. This feature was suggested as a candidate centromere (13), although there has been no experimental verification. Progress in this area has been hampered by the small size of Plasmodium chromosomes, the fact that chromatin does not condense during mitosis (14), difficulties in cloning large segments of AT-rich P. falciparum DNA, and the limited flexibility of parasite genetic manipulation techniques (15). The P. falciparum genome is also unusual in that, with the exception of a putative orthologue of CenH3 (PF13_0185, www.plasmodb.org) (16), there is a lack of obvious equivalents of most of the widely conserved “core” proteins that display a constitutive centromere location and have a central role in kinetochore assembly (3). However, structures analogous to kinetochores have been confirmed in P. falciparum. The first rigorous determination that the parasite had 14 chromosomes was established by counting the number of kinetochores, after three-dimensional reconstruction of the mitotic spindle in schizonts (17).

One protein that has been widely implicated in centromere function (18, 19) and is expressed in P. falciparum is topoisomerase-II (PF14_0316, www.plasmodb.org). During metaphase, topoisomerase-II accumulates specifically at active centromeres, where it is required for proper kinetochore/centromere structure and to decatenate sister chromatids before segregation (1923). Decatenation is a three-step process that involves double-stranded DNA cleavage, passage of one strand of the duplex through the break, and religation to repair the lesion. The topoisomerase-II inhibitor etoposide acts by stabilizing the enzyme–DNA intermediate, blocking the religation step and resulting in DNA cleavage at sites specified by topoisomerase-II binding (24). On this basis, etoposide has been used as a biochemical marker for active centromeres and to further assess the key role of topoisomerase-II in centromere function (23, 25–27). Here, we describe data obtained by using a biochemical mapping approach based on topoisomerase-II activity that has allowed us to produce experimental evidence on the location and nature of centromeric DNA in P. falciparum.


To identify the sites of accumulation of topoisomerase-II on chromosomes of P. falciparum, cultures of infected human red blood cells (6–10% parasitemia) were incubated with 100 μM etoposide for various times as indicated (Materials and Methods and legends to Fig. 11 4). All experiments were performed by using the haploid asexual stages of P. falciparum 3D7, the genome project strain. Parasite chromosomal DNA was isolated and fractionated by pulsed-field gel electrophoresis (PFGE). After Southern blotting and hybridization with cloned probes, we observed that chromosome cleavage had occurred and were able to determine the sizes of the resulting fragments. The data from chromosomes 1 (0.70 Mb) and 5 (1.4 Mb) illustrate the type of results obtained (Figs. 11 4). They show that topoisomerase-II activity is concentrated at a single locus on each chromosome and that the sizes of the cleavage products can be used to infer the location of centromeric DNA.

Fig. 1.
Mapping of etoposide-mediated topoisomerase-II cleavage sites in P. falciparum chromosome 1 by using PFGE analysis and Southern hybridization. Cultures of P. falciparum (strain 3D7)-infected red blood cells were treated with 100 μM etoposide (Sigma) ...
Fig. 2.
Fine-mapping of etoposide-mediated topoisomerase-II cleavage sites in P. falciparum chromosome 1. Genomic DNA from etoposide-treated (20-min incubation at concentrations of 0–100 μM) and nontreated parasites was restriction-digested and ...
Fig. 3.
Mapping of etoposide-mediated topoisomerase-II cleavage sites in chromosome 5. Chromosomal DNA was isolated from treated P. falciparum and fractionated by PFGE as outlined in the legend to Fig. 1. Parasites were treated with 100 μM etoposide for ...
Fig. 4.
Fine-mapping of etoposide-mediated topoisomerase-II cleavage sites in chromosome 5. (Top) Genomic DNA from etoposide-treated (100 μM etoposide for 0–120 min) and nontreated parasites was restriction-digested and analyzed by Southern hybridization ...

With the chromosome 1-specific probes PfGCAP and C1-B, we identified major products of 480 and 220 kb, respectively (Fig. 1), tentatively mapping the cleavage site(s) to a 9-kb “gene-free” region of the chromosome that contains a prominent 2.4-kb AT-rich domain similar to the type of element previously suggested as a candidate centromere (12, 13). Using a probe (C1-A) derived from sequences immediately adjacent to this element, we found that both of these fragments could be detected (Fig. 1). This hybridization pattern indicates that, although etoposide-mediated topoisomerase-II cleavage is restricted to a specific locus, the susceptible area is sufficiently broad that it covers each side of the C1-A sequence. As a result, the C1-A probe hybridizes to both the 480- and the 220-kb products. By implication, therefore, topoisomerase-II cleavage in P. falciparum must be a regional phenomenon.

To assess this finding further, and to determine the extent of the topisomerase-II activity, we restriction-digested genomic DNA from etoposide-treated parasites and examined the cleavage pattern in the vicinity of this putative centromeric region (Fig. 2), revealing the presence of several major bands, indicative of a region of cleavage that stretched over 8–10 kb, including the AT-rich domain and the gene corresponding to the C1-A probe. The observation that topoisomerase-II activity is not restricted to the immediate confines of the AT-rich domain implies that the skewed base content of this region does not, by itself, confer susceptibility to cleavage. When these blots were rehybridized with PfGCAP, a gene situated 50 kb downstream of the putative centromere, no cleavage products were detected (Fig. 2), confirming that topoisomerase-II activity is restricted to sites immediately adjacent to the AT-rich domain. The locations of topoisomerase-II cleavage sites within mammalian centromeres have been mapped to a resolution of ≈50–150 kb (2527).

For chromosome 5, we used probes C5-A and C5-B, which were derived from DNA sequences located on either side of a similar AT-rich domain, and probe C5-C, which was derived from an ORF ≈150 kb from this element. On PFGE blots, we detected products of 500 and 1,000 kb, respectively, with probes C5-A and C5-C, delineating the cleavage sites to the region between these two sequences (Fig. 3). With probe C5-B, we detected both fragments, implying that topisomerase-II cleavage had occurred on each side of the corresponding genomic DNA sequence. Consistent with this finding, when restriction-digested genomic DNA was analyzed (Fig. 4), we observed that topoisomerase-II activity was spread over a 10- to 12-kb region encompassing the C5-B sequence and the 2.3-kb AT-rich domain, with at least four major cleavage sites.

The cleavage patterns produced by all 14 P. falciparum chromosomes after etoposide treatment were determined by Southern analysis of PFGE gels (available from the author upon request). The probes used and the proposed locations of centromeric DNA for each chromosome are summarized in Fig. 5. Topoisomerase-II activity could be mapped to the vicinity of single AT-rich domains with a sharply defined size limit of 2.3–2.5 kb (Fig. 6and data available from the author upon request). These domains exhibit a minimal GC content (<3%), are present in only one copy per chromosome, and are located in gene-free regions that vary from 6 to 12 kb. We could find no common themes to centromere-proximal genes in terms of their orientation, temporal expression, or putative function (www.plasmodb.org). Despite their high AT content, the domains are distinct from introns and other untranslated regions of the P. falciparum genome. They lack the characteristic long stretches of polyA, polyT, and A/T dinucleotide or trinucleotide repeats. We also examined each of the AT-rich domains for other repetitive arrays or for conserved elements (28). A series of repeat motifs of various lengths, copy number, and sequence were identified (Fig. 6). However, apart from the nucleotide content and tightly restricted size range of the domains, we could detect no features that were extensively conserved between chromosomes. In the case of chromosome 10, the data indicate that the topoisomerase-II cleavage site is 1.1 Mb from the left-hand telomere (Fig. 5 and information available from the author upon request), although there is not an apparent AT-rich domain at this site or elsewhere on the chromosome (www.plasmodb.org). Whether this exception represents a functional variant or reflects a gap in sequence assembly remains to be determined.

Fig. 5.
Location of etoposide-mediated topoisomerase-II cleavage sites (black dots) on all 14 P. falciparum chromosomes. The chromosomal locations of the probes used are shown as red bars, and the relevant data are available from the author upon request. The ...
Fig. 6.
Schematic of the AT-rich domains and the corresponding gene-free regions coincident with topisomerase-II cleavage sites on each P. falciparum chromosome. The locations of the proximal ORFs (blue) and their PlasmoDB (www.plasmodb.org) identifiers are shown. ...


We have shown here that treatment of asexual stage P. falciparum parasites with the topoisomerase-II inhibitor etoposide results in chromosomal cleavage at single loci. With the availability of a completed genome sequence (12) and the fact that parasite chromosomes are of a size range that is amenable to analysis using PFGE-based approaches, we have been able to accurately map these lesions. The resulting data identify the major sites where catalytically active topisomerase-II accumulates on P. falciparum chromosomes. Topoisomerase-II plays a central role in chromosome biology and has been implicated in the segregation process in organisms ranging from yeast (18, 29, 30) to vertebrates (3133). After replication, sister chromatids remain attached, partly through strand catenation at centromeres (22), as cells enter mitosis. The premitotic accumulation of topoisomerase-II at functional centromeres is a major regulator of sister-chromatid cohesion and is essential for ordered segregation (19). Evidence for this centromeric sequestration of enzyme activity includes the observation that etoposide-mediated cleavage sites in human chromosomes can be mapped to within the α-satellite arrays (23, 25–27). It has even been suggested that these topoisomerase-II-binding sites could function as epigenetic markers for kinetochore assembly (3). Using etoposide, we have also found (J.M.K., unpublished observation) that topoisomerase-II activity in chromosomes of the parasitic protozoan Trypanosoma cruzi maps specifically to regions that we had shown to be essential for mitotic stability (34). Intriguingly, the cleavage sites in P. falciparum that we have mapped here encompass the extremely AT-rich elements of 2.3–2.5 kb, which are present once per chromosome and which have been proposed as candidate centromeres (12, 13).

These results, based on sites of topoisomerase-II accumulation, provide experimental evidence on the location of centromeres in P. falciparum. The findings suggest the presence of a class of compact centromeric DNA that differs, in terms of size and organization, from both the “regional” and point centromeres described previously in other organisms. These unusual properties may also be a conserved feature of centromeres in the Plasmodium genus. In Plasmodium vivax, although the genome assembly is not yet complete, sharply defined AT-rich domains similar to, and syntenic with, those in P. falciparum can be detected on contigs from chromosomes 4, 6, and 14 (www.plasmodb.org). The precise role of the centromeric DNA sequence in chromosome segregation remains controversial and is a focus of considerable research (3, 9, 35). In Plasmodium species, the lack of experimental data on the location and organization of centromeric DNA has limited any meaningful progress in this area. For instance, it remains to be established how centromeres are recognized by the segregation machinery, which proteins are involved in mediating the process, and whether centromeres are even associated with areas of heterochromatin. In the case of the AT-rich domains, we propose that the DNA sequence per se probably does not play direct a role in segregation, because there is a lack of evident interchromosomal conservation. Rather, we suggest that features or properties associated with the skewed nucleotide content and the sharply defined size range, which are conserved between chromosomes, are more likely to be determinants of centromere function. These AT-rich domains could be involved, directly or indirectly in interaction with specific proteins or complexes, or be implicated in some other aspect of higher-order chromatin structure.

The demonstration that topisomerase-II accumulates on P. falciparum chromosomes at sites adjacent to the AT-rich domains now provides a more robust framework for identifying and functionally characterizing other determinants of centromere function. This finding could have important implications, because the unusual nature of centromeric DNA in P. falciparum, combined with the apparent absence of genes for most of the conventional core centromeric proteins, suggests that some aspects of the segregation process may be distinct from those in the human host. In addition, there could also be differences in the factors that mediate topisomerase-II accumulation at centromeres or that regulate enzyme activity. For example, in Xenopus, conjugation of the ubiqutin-related peptide SUMO-2 to topoisomerase-II by the SUMO ligase PIASy is thought to be a requirement for proper centromeric function and chromosome segregation (33). There is no obvious equivalent of PIASy in the P. falciparum genome database. Finally, our results suggest that the biochemical mapping approach, based on etoposide-mediated topoisomerase-II cleavage, can be applied to other protozoa. This is an extensive group of organisms, frequently of medical importance, where information on the location and organization of centromeric DNA is almost universally lacking.

Materials and Methods

Parasite Culturing.

P. falciparum strain 3D7 was maintained under standard conditions (36). Contaminating trophozoites and schizonts were removed by treatment with 5% sorbitol (37). Synchronous ring-stage cultures were then split into flasks at 6–10% parasitemia and 5% hematocrit. Twenty-six hours after synchronization, when parasites had developed into mature trophozoites/early schizonts, etoposide (Sigma) solubilized in DMSO was added. An equivalent volume of DMSO was added to the control flasks.

Isolation and Analysis of DNA.

To isolate parasite DNA, pellets of the infected red cell culture were resuspended in 10 ml of 50 mM Tris, pH 8.0/5 mM EDTA/100 mM NaCl (TSE), and saponin was added to a final concentration of 0.1%. After red cell lysis was observed, the sample was centrifuged at 2,500 × g for 10 min at 20°C and the parasite pellet washed twice in RPMI medium 1640. For preparation of chromosomal-sized DNA, the pellet was resuspended in TSE and an equal volume of 1.6% low-melting-point agarose (Bio-Rad) and then aliquoted into block molds. Chromosome blocks were incubated in 10 mM Tris/1 mM EDTA, pH 8.0 (TE) supplemented with 1% sodium N-lauroyl sarcosine and 2 mg·ml−1 proteinase K (Sigma) for 72 h before analysis by PFGE. For genomic DNA isolation, the pellet was proteinase K-digested and purified as described in ref. 38. Chromosomal DNA was fractionated by using a Bio-Rad CHEF Mapper system with autoalgorithms set to the appropriate molecular-mass range. Gels were blotted onto nylon membranes by using standard techniques. DNA fragments used for hybridization were generated by PCR, cloned by using the pGEM-T Easy Vector System 1 (Promega) and repurified before radiolabeling.

Table 1
Consensus sequences of the two highest scoring repeat motifs for each chromosome


We thank Richard Pearce for assistance with analysis of repeat sequences; Quinton Fivelman for discussions on the function of centromere-proximal genes; colleagues at the London School of Hygiene and Tropical Medicine for various gene probes; and Hans Dessens, David Horn, Sam Obado, Martin Taylor, Spencer Polley, and Brendan Wren for critical comments on the manuscript. We acknowledge the work of colleagues at PlasmoDB and the sequencing centers (The Institute for Genomic Research, the Wellcome Trust Sanger Institute, and Stanford University). This work was supported by Biotechnology and Biological Sciences Research Council Grant C501292 (to J.M.K.) and Wellcome Trust Grant 066742 (to D.A.B.).


pulsed-field gel electrophoresis.


Conflict of interest statement: No conflicts declared.

This paper was submitted directly (Track II) to the PNAS office.


1. Sun X., Wahlstrom J., Karpen G. Cell. 1997;91:1007–1019. [PMC free article] [PubMed]
2. Cleveland D. W., Mao Y., Sullivan K. F. Cell. 2003;112:407–421. [PubMed]
3. Fukagawa T. Chromosome Res. 2004;12:557–567. [PubMed]
4. Steiner N. C., Hahnenberger K. M., Clarke L. Mol. Cell. Biol. 1993;13:865–874.
5. Wood V., Gwilliam R., Rajandream M.-A., Lyne M., Lyne R., Stewart A., Sgouros J., Peat N., Hayles J., Baker S., et al. Nature. 2002;415:871–880. [PubMed]
6. Cottarel G., Shero J. H., Hieter P., Hegemann J. H. Mol. Cell. Biol. 1989;9:3342–3349. [PMC free article] [PubMed]
7. Pidoux A. L., Allshire R. A. Chromosome Res. 2004;12:521–534. [PubMed]
8. Schueler M. G., Higgins A. W., Rudd M. K., Gustashaw K., Willard H. F. Science. 2001;294:109–115. [PubMed]
9. Wong L. H., Choo K. H. Trends Genet. 2004;20:611–616. [PubMed]
10. Clarke L., Baum M. P. Mol. Cell. Biol. 1990;10:1863–1872. [PMC free article] [PubMed]
11. Henikoff S., Ahmad K., Malik H. S. Science. 2001;293:1098–1102. [PubMed]
12. Gardner M. J., Hall N., Fung E., White O., Berriman M., Hyman R. W., Carlton J. M., Pain A., Nelson K. E., Bowman S., et al. Nature. 2002;419:498–511. [PMC free article] [PubMed]
13. Bowman S., Lawson D., Basham D., Brown D., Chillingworth T., Churcher C. M., Craig A., Davies R. M., Devlin K., Feltwell T., et al. Nature. 1999;400:532–538. [PubMed]
14. Arnot D. E., Gull K. Ann. Trop. Med. Parasitol. 1998;92:361–365. [PubMed]
15. Carvalho T. G., Menard R. Curr. Issues Mol. Biol. 2005;7:39–55. [PubMed]
16. Malik H. S., Henikoff S. Nat. Struct. Biol. 2003;10:882–891. [PubMed]
17. Prensier G., Slomianny C. J. Parasitol. 1986;72:731–736. [PubMed]
18. Bachant J., Alcasabas A., Blat Y., Kleckner N., Elledge S. J. Mol. Cell. 2002;9:1169–1182. [PubMed]
19. Carpenter A. J., Porter A. C. G. Mol. Biol. Cell. 2004;15:5700–5711. [PMC free article] [PubMed]
20. Rattner J. B., Hendzel M. J., Furbee C. S., Muller M. T., Bazett-Jones D. P. J. Cell Biol. 1996;134:1097–1109. [PMC free article] [PubMed]
21. Andersen C. L., Wandall A., Kjeldsen E., Mielke C., Koch J. Chromosome Res. 2002;10:305–312. [PubMed]
22. Haering C. H., Nasmyth K. BioEssays. 2003;25:1178–1191. [PubMed]
23. Porter A. C. G., Farr C. J. Chromosome Res. 2004;12:569–583. [PubMed]
24. Chen G. L., Yang L., Rowe T. C., Halligan B. D., Tewey K. M., Liu L. F. J. Biol. Chem. 2004;259:13560–13566. [PubMed]
25. Floridia G., Zatterale A., Zuffardi O., Tyler-Smith C. EMBO Rep. 2000;1:489–493. [PMC free article] [PubMed]
26. Spence J. M., Critcher R., Ebersole T. A., Valdivia M. M., Earnshaw W. C., Fukagawa T., Farr C. J. EMBO J. 2002;21:5269–5280. [PMC free article] [PubMed]
27. Spence J. M., Fournier R. E., Oshimura M., Regnier V., Farr C. J. Chromosome Res. 2005;13:637–648. [PubMed]
28. Benson G. Nucleic Acids Res. 1999;27:573–580. [PMC free article] [PubMed]
29. Holm C., Goto T., Wang J. C., Botstein D. Cell. 1985;41:553–563. [PubMed]
30. Uemura T., Ohkura H., Adachi Y., Morino K., Shiozaki K., Yanagida M. Cell. 1987;50:917–925. [PubMed]
31. Downes C. S., Mullinger A. M., Johnson R. T. Proc. Natl. Acad. Sci. USA. 1991;88:8895–8899. [PMC free article] [PubMed]
32. Shamu C. E., Murray A. W. J. Cell Biol. 1992;117:921–934. [PMC free article] [PubMed]
33. Azuma Y., Arnaoutov A., Anan T., Dasso M. EMBO J. 2005;24:2172–2182. [PMC free article] [PubMed]
34. Obado S. O., Taylor M. C., Wilkinson S. R., Bromley E. V., Kelly J. M. Genome Res. 2005;15:36–43. [PMC free article] [PubMed]
35. Schueler M. G., Dunn J. M., Bird C. P., Ross M. T., Viggiano L., Rocchi M., Willard H. F., Green E. D. Proc. Natl. Acad. Sci. USA. 2005;102:10563–10568. [PMC free article] [PubMed]
36. Trager W., Jensen J. B. Science. 1976;193:673–675. [PubMed]
37. Lambros C., Vanderberg S. P. J. Parasitol. 1979;65:418–420. [PubMed]
38. Kelly J.M. In: Protocols in Molecular Parasitology. Hyde J. E., Walker J. M., editors. Totowa,NJ: Humana; 1993. pp. 312–321. Methods in Molecular Biology, series ed.

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...