Origin of Plasmodium falciparum malaria is traced by mitochondrial DNA

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Molecular and Biochemical Parasitology 111 (2000) 163 – 171 www.parasitology-online.com.

Origin of Plasmodium falciparum malaria is traced by mitochondrial DNA David J. Conway a,*, Caterina Fanello a, Jennifer M. Lloyd a, Ban M.A.-S. Al-Joubori a, Aftab H. Baloch a, Sushela D. Somanath a, Cally Roper a, Ayoade M.J. Oduola b, Bert Mulder c, Marinete M. Povoa d, Balbir Singh e, Alan W. Thomas f a

Department of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, Keppel St, London WC1E 7HT, UK b College of Medicine, Uni6ersity of Ibadan, Ibadan, Nigeria c Malaria Department, OCEAC, Yaounde, Cameroon d Ser6ico de Parasitologia, Instituto E6andro Chagas, Belem PA, Brazil e Uni6ersiti Malaysia Sarawak, Kota Samarahan 94300, Sarawak, Malaysia f Biomedical Primate Research Centre, PO Box 3306, 2288 GH Rijswijk, The Netherlands Received 24 May 2000; received in revised form 27 July 2000; accepted 28 July 2000

Abstract The origin and geographical spread of Plasmodium falciparum is here determined by analysis of mitochondrial DNA sequence polymorphism and divergence from its most closely related species P. reichenowi (a rare parasite of chimpanzees). The complete 6 kb mitochondrial genome was sequenced from the single known isolate of P. reichenowi and from four different cultured isolates of P. falciparum, and aligned with the two previously derived P. falciparum sequences. The extremely low synonymous nucleotide polymorphism in P. falciparum (p= 0.0004) contrasts with the divergence at such sites between the two species (K= 0.1201), and supports a hypothesis that P. falciparum has recently emerged from a single ancestral population. To survey the geographical distribution of mitochondrial haplotypes in P. falciparum, 104 isolates from several endemic areas were typed for each of the identified single nucleotide polymorphisms. The haplotypes show a radiation out of Africa, with unique types in Southeast Asia and South America being related to African types by single nucleotide changes. This indicates that P. falciparum originated in Africa and colonised Southeast Asia and South America separately. © 2000 Elsevier Science B.V. All rights reserved. Keywords: Plasmodium falciparum; Plasmodium reichenowi; Mitochondrial genome; Genetics; Species; Evolution Abbre6iations: mt, mitochondrial; nt, nucleotide. Note: Nucleotide sequences reported in this paper are available in the EMBL, GenBank™ and DDBJ databases under the accession numbers AJ251941 and AJ276844-AJ276847. * Corresponding author. Tel.: + 44-20-79272331; fax: + 44-20-76368739. E-mail address: [email protected] (D.J. Conway). 

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1. Introduction The origins of major infectious diseases of humans are of relevance to understanding evolution of pathogen virulence and natural selection on the human genome [1]. The malaria parasite Plasmodium falciparum has had an unparalleled impact on human gene polymorphisms [2], and remains a major agent of human mortality [3]. Human sickle cell and a-thalassaemia alleles (which confer some protection from P. falciparum malaria) have multiple independent geographical origins in Africa and Asia, and no ancestral haplotype is very widespread, suggesting that selection by malaria has mainly occurred after human settlement of these regions [4]. Emergence of P. falciparum from a small founding population less than 50 000 years ago has been suggested from the very low level of synonymous single nucleotide polymorphism in some housekeeping genes [5]. However, rRNA gene sequence data indicate that it diverged from the most closely related extant species P. reichenowi at approximately the same time as chimpanzees and hominids (5 – 10 million years ago) [6], and some alleles of P. falciparum antigen genes may have even more ancient origins which would predate this split [7 – 10]. The origin of a species, and its geographical spread, can sometimes be resolved by study of mitochondrial (mt) sequence divergence and variation [11]. Malaria parasites have a small (  6 kb) tandemly repeated linear mt genome [12] which is uniparentally inherited [13]. Replication involves recombination within but probably not between mt lineages [14]. A broad phylogeny of distantly related Plasmodium species based on sequences of the mitochondrial cytochrome b gene [15] was consistent with that derived from rRNA gene sequences in chromosomal DNA [6]. Here, the complete mt genome sequence was derived from P. reichenowi and four cultured isolates of P. falciparum, and analysed together with two previously derived P. falciparum sequences. This identifies, and provides a quantitative survey of, intraand inter-specific nucleotide differences. The single nucleotide polymorphisms (SNPs) and their composite haplotypes within P. falciparum were

then determined from 104 field isolates, revealing a stark geographical radiation of mtDNA haplotypes.

2. Materials and methods

2.1. Sequence analysis of P. reichenowi and P. falciparum mitochondrial genomes Complete mt genome sequences were derived from P. reichenowi and P. falciparum from diverse sources (7G8 from Brazil, NF54 imported to the Netherlands from Africa, T9/96 and K1 from Thailand). This was performed by PCR amplification and sequencing of eight overlapping regions covering a complete linear copy of the mt genome. The nucleotide (nt) positions based on the EMBL sequence of the C10 P. falciparum clone (accession no. M76611), and pairs of oligonucleotide PCR primers were: (1) nt 85–741, fwd 5%-CAACACCATCCAATTTGATTGGG-3%, rev 5%-CAGAATAAATGAACATGTATGTCC3%; (2) nt 650–1626, fwd 5%-CCTTACGTACTCTAGCTATGAAC-3% rev 5%-TTGCAAGGCTGCGATGAGACG-3%; (3) nt 1493–2289, fwd 5%AGAACGGTGAGATAATGTGCCG-3%, rev 5%AAATCCTCCGAATAATCCTGGCA-3%; (4) nt 2142–3135, fwd 5%-ATTATCAGTAATA-CTACGTACTGAAT-3%, rev 5%-ACCTAATATAACTCCAGTAGTACC-3%; (5) 3049–3954, fwd 5%TTTGGTATGATACACAGCTCTTCA-3%, rev 5%-TCCAAATTACTGCTACTGGAATAG-3%; (6) nt 3841–4748, fwd 5-TATTTATTGTAACTGCTTTCGTTG, rev 5%-AAGCATCCATCTACAGCTATGGT-3%; (7) nt 4644–5534, fwd 5%-TGACATTTCTGAGTATTGAGCGG-3%, rev 5%-CTTCCTGATCGACTCGTGAGG-3%; (8) nt 5459–(next tandem copy)185, 5%-CTACAAGTTCACTGTCAACTA-3%, rev 5%-CATCTGTTTCTATTCTTATGGGC-3%. Amplification of fragment (5) using the above primers was inefficient for P. reichenowi, and sequences of the flanking overlapping fragments showed that nucleotide differences between the species existed within the template sequences, so another pair of primers was designed for this region of the P. reichenowi

D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171

sequence: fwd 5%-TTTGGTATGATACATAGCTCTTCC-3%, rev 5%-TCCAAATAACTGCTACTGGAATAG-3%. Amplification was performed in a total volume of 100 ml with the following components: 2 mM MgCl2, 1 × Bioline amplification reaction buffer, 200 mM dNTPs, 0.1 mM each oligonucleotide primer, 0.5 U BioTaq™ (Bioline) polymerase, and 2 ml solution of DNA template. A hot start step was performed at 95°C for 10 min, after which samples were placed on ice and BioTaq™ polymerase added. Temperature cycling was as follows; 95°C for 1 min, 60°C for 1 min, 72°C for 1 min for 3 cycles; 95°C for 1 min, 55°C for 1 min, 72°C for 1 min for 42 cycles, and 10 min at 72°C final extension. The amplified products (5 ml) were run on 1.5% agarose gels in TAE buffer to detect the correct band, using EcoR1 + HindIII-digested Lambda DNA (Promega) as molecular mass markers. Amplified products were cloned in pGEM®-T Easy Vector (Promega), with transformation and growth performed in JM 109 Escherichia coli High Efficiency Competent Cells. Inserts in purified recombinant plasmids were sequenced using Thermo Sequenase™ dye terminator cycle sequencing pre-mix kit (Amersham™) using SP6 and T7 universal primers. Samples were run on an ABI -377 DNA sequencer (Perkin – Elmer), and sequences were checked and assembled using Sequence Navigator software. To exclude any error, the P. reichenowi mt genome was sequenced and edited twice, starting from independent PCR amplifications. Any ambiguity, and putative polymorphism among P. falciparum isolates, was checked by additional amplification and sequencing (very important as the actual level of polymorphism within P. falciparum was considerably lower than the PCR misincorporation error rate). Inter-specific nucleotide divergence (K) and intraspecific polymorphism (p) among aligned sequences was calculated with the DnaSP 3.0 software [16].

2.2. P. falciparum populations sampled for mtDNA haplotypes In order that haplotypes could be resolved, 104 P. falciparum isolates were chosen, each of which


had appeared not to contain mixed genotypes at nuclear polymorphic loci. There were 73 from Africa (25 isolates from Ibadan, Nigeria; 30 from Yaounde, Cameroon, 18 from Kwazulu-Natal, South Africa), 11 from Southeast Asia (Malinsau in Sabah, Malaysian Borneo) and 20 from South America (12 from Porto Velho and 8 from Tailandia, Brazil). Genomic DNA was prepared from the original isolates without in vitro culture.

2.3. Genotyping mtDNA in natural populations of P. falciparum The four polymorphic nucleotides which were identified by sequencing (positions 772, 1692, 4179, and 4952) were typed in the field isolates by PCR amplification followed by sequence specific oligonucleotide probing (PCR-SSOP). Four small fragments of the mt genome were amplified, each incorporating one of the polymorphic nucleotide sites. The nucleotide positions of the fragments, and the primer pairs were as follows: (1) nt 651– 880, fwd 5%-CCTTACGTACTCTAGCTATGAAC-3%, rev 5%-ATATATGATACTTCTACCGAATGG; (2) nt 1642–1730, fwd 5%-GTGTATTGTTGCCTTGTACACA, rev 5%-CATCTCAACTCTACAGGTTAAC-3%; (3) nt 4065–4289, fwd 5%-CATTTACATGGTAGCACAAATCC-3%, rev 5%-TGGTAGAAAGTACCATTCAGGTA-3%; (4) nt 4849–5083, fwd 5%-GGAGTTGGCAAGTTAAAGAAGTT-3%, rev 5%-GTCAATCAAACATGAATATAGACG-3%. To increase the yield of fragment (2), nested PCR was performed on a template (nt 1635–1758) which had been prepared by a first round PCR with the following primers; fwd 5%-ATCTAGCGTGTATTGTTGCCTTG-3%, rev 5%-GGCGTAAAATTACCTTTCCGGC-3%. Amplifications were performed in 10 ml volumes in 96-well plates, with the following components; 1.5 mM MgCl2, 1 × Bioline amplification reaction buffer, 200 mM dNTPs, 0.1 mM each oligonucleotide primer, 0.5 U BioTaq™ (Bioline) polymerase and 1 ml solution of DNA template (with variable concentration of DNA in field isolates). Temperature cycling was as follows; 95°C for 1 min, 60°C for 1 min, 72°C for 1 min for 3 cycles; 95° for 1 min, 55°C for 1 min, 72°C for 1 min for 42 cycles.


D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171

Products were denatured, and 1.5 ml dot-blotted onto replicate nylon membranes (MagnaGraph™) in 96-dot arrays. Membranes were blocked (in 4× SSPE, 0.1% Lauroylsarcosine, 1.0% milk powder) at 37°C for 30 min. Oligonucleotide probes specific for the alternative alleles of each of the 4 single nucleotide polymorphisms were, 772T 5%-TAACCAGATTATTTCAAC-3%, 772C 5%-TAACCAGACTATTTCAAC-3%, 1692A 5%-TATACTGAGTATAGAACT-3%, 1692G 5%-TATACTGGGTATAGAACT-3%, 4179T 5%-TTTATTTTTAATACAAAG-3%, 4179C 5%-TTTATTTCTAATACAAAG-3%, 4192T 5%-CTATTTATATTTATCGAT-3%, 4192C 5%-CTATTTATACTTATCGAT-3%. Probes were 3%-labelled with digoxigenin (DIG) (Boehringer Mannheim) and incubated with membranes at a final concentration of 1 nM in separate tubes containing 5 ml TMAC hybridisation solution (3M Tetramethylammonium chloride, 50 mM Tris–HCl pH 8.0, 0.1% SDS, 2mM EDTA pH 8.0), rotating at 53°C for 90 min. Membranes were then washed while agitating for 2 × 10 min in 2× SSPE per 0.1% SDS at room temperature (low-stringency washes) and 2×10 min (high-stringency washes) in TMAC solution at 56°C. Probes were detected using anti-DIG-AP Fab fragment conjugated with alkaline phosphatase and CSPD reagent as a substrate for alkaline phosphatase (Boehringer Mannheim) following the manufacturer’s guidelines. Membranes were exposed to Hyperfilm-ECL for 1 – 3 h and films were developed and scored independently by two investigators to determine the allelic pattern of hybridisation. Any rare discordance or uncertainly in allelic scoring led to a sample being repeated. This procedure robustly discriminates alleles differing by a single nucleotide, as described previously for studies on chromosomal gene loci in P. falciparum [17,18]. Haplotypes were resolved (the isolates had been intentionally chosen not to contain mixed clones), and their frequencies determined in each geographical population sample. The inter-population component of variance in haplotype frequencies, FST, was calculated as the u value [19], and the statistical significance of its deviation from zero was derived by running 10 000 permu-

tations on randomised data, using the FSTAT 1.2 program.

3. Results

3.1. mtDNA sequence polymorphism in P. falciparum and di6ergence from P. reichenowi Full mt genome sequences were derived from the single known isolate of P. reichenowi and four isolates of P. falciparum (7G8, NF54, T9/96 and K1), and aligned together with sequences from an additional two P. falciparum isolates which had previously been reported (C10 of uncertain origin, CAMP from peninsular Malaysia) [12,20]. In the complete alignment of 5965 base pairs, there were 139 nucleotides which differed between P. reichenowi and each of the six P. falciparum isolates, and four nucleotides which differed among any of the P. falciparum isolates (Fig. 1). The divergence and polymorphism at different types of nucleotide sites in the mt genome is summarised in Table 1. The nucleotide differences are most abundant at synonymous sites within protein coding genes, which supports the idea that there are probably strong constraints on changes at other sites in the mt genome [20]. At synonymous nucleotide sites, the divergence between the species (12%) is 300 times greater than the polymorphism (0.04%) within P. falciparum. Thus, the time since the common ancestor of both species is probably at least 2 orders of magnitude longer than the time since the origin of modern P. falciparum populations. If the mitochondrial lineages separated with the species approximately 5–10 million years ago [6], P. falciparum mt sequences probably derive from an origin within the last 50 000 years.

3.2. Geographical distribution and radiation of mtDNA haplotypes in P. falciparum To investigate the geographical origin of P. falciparum, haplotypes of mtDNA were studied by typing the four single nucleotide polymorphisms described above (nucleotides 772, 1692, 4179, and 4952), in 104 isolates from different

D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171

locations. Fig. 2 shows an example of the typing of alleles by PCR-SSOP (illustrated for eight isolates typed at position 772) and all 104 isolates were typed in a similar manner at each of the 4 nucleotide positions. Five haplotypes were found, with frequencies in each of the populations shown


in Table 2. Three haplotypes are seen in Africa (the most frequent being CGCC) and there is significant heterogeneity in frequencies among the three African populations (FST = 0.21, PB 0.001). The populations sampled from other continents have different haplotypes (TGCC in Malaysia,

Fig. 1. Scheme of the P. falciparum/P. reichenowi mitochondrial (mt) genome. Nucleotide positions are numbered as by Feagin et al. [12] (accession no. M76611), putative positions of genes are shown as shaded boxes (grey, protein coding genes; black, LSU rRNA fragments; white, SSU rRNA fragments), with transcription in forward orientation shown above the line and transcription from the opposite strand shown below (the mt genome is poly-cistronically transcribed from each strand [32,33]). Nucleotide positions of these genes were derived from accession no. M76611 (version 12, updated 04-MAR-2000), and it is possible that additional rRNA gene fragments may be described in the future [34]. Positions of the single nucleotide differences between the species and polymorphisms among the six isolates of P. falciparum are shown underneath. The mt genome sequences of P. reichenowi and P. falciparum NF54, 7G8, K1 and T9/96 have the accession numbers AJ251941 and AJ276844 — AJ276847 (P. falciparum C10 and CAMP sequences previously reported have accession numbers M76611 and M99416). The four single nucleotide polymorphisms are at positions 772 (T/C), 1692 (G/A), 4179 (T/C), and 4952 (T/C). The haplotypes are CGCC (T9/96 and CAMP), TGCC (K1 and C10), CGCT (NF54), and CATC (7G8). The P. reichenowi sequence also has a single nucleotide deletion at position 5451, and the reported sequence of CAMP has a single nucleotide deletion at position 5883. Table 1 Mitochondrial DNA divergence between P. falciparum and P. reichenowi and polymorphism within P. falciparum Type of nucleotide site (n =number analysed)

Synonymous (n =817) Non-synonymous (n = 2693) rRNA genes (n =1101) Intergenic (n = 1354)b Total (n = 5965) a

Divergence between P. falciparum and P. reichenowi a

Polymorphism within P. falciparum

Number of fixed differences

Number of polymorphic sites

Divergence (K)

Diversity (p)

93 19 9 18

0.1201 0.0082 0.0082 0.0147

1 1 0 2

0.0004 0.0002 0.0000 0.0005





The six complete mtDNA genome sequences of P. falciparum and one of P. reichenowi are analysed (5965 aligned nucleotide positions, after omission of 2 positions containing deletions). Divergence (K) and diversity (p) indices show the proportion of nucleotides which differ in pairwise comparisons of sequences, for each given type of nucleotide position. The Jukes and Cantor correction is applied, which has a marginal effect for these low levels of sequence differences. b These intergenic positions are not known to code for protein or rRNA (EMBL accession no. M76611, updated 04-MAR-2000) but it is possible that rRNA transcripts will be mapped to some of them [34]. A small number of other positions encoded both rRNA and protein sequences on opposite strands and are categorised here as the latter.


D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171

Fig. 2. Example of typing one of the single nucleotide polymorphisms (nt position 772), using polymerase chain reaction amplification folowed by allele sequence-specific oligonucleotide probing (PCR-SSOP). Results are shown from a corresponding portion of replicate membranes probed with oligonucleotides recognising alleles 772T and 772C (the oligonucleotides differ at the ninth position). The membrane portions each contain PCR products from eight isolates (four isolates from Malaysia on the top row, and 4 from South Africa on the bottom row). It can be clearly seen that the isolates on the top row hybridise to probe 772T and not to 772C, so these parasites have nucleotide T at position 772; conversely the isolates on the bottom row hybridise to probe 772C and not 772T, so these parasites have nucleotide C at this position. Such unequivocal discrimination can be routinely obtained for all single nucleotide polymorphisms (protocol outlined in Section 2). Observed variation in the signal intensity for different isolates is a normal feature of varying parasite DNA abundance between field isolates, which presents no difficulty for scoring relative hybridisation of allelic probes for each individual isolate. Table 2 Mitochondrial DNA haplotype frequencies in geographical samples of P. falciparum field isolates Haplotype (nt positions 772, 1692, 4179, 4952)

Nigeria (n= 25)

Cameroon (n= 30)

South Africa (n = 18)

Malaysia (n = 11)

Brazil (n = 20)


0.60 0.36 0.04 – –

0.50 0.50 – – –

1.00 – – – –

– – – 1.00 –

– 0.10 – – 0.90

and CATC in Brazil) and only two Brazilian isolates have an African haplotype (CACC). Thus, the inter-continental variance in haplotype frequencies is extremely high (FST =0.65, P B 0.001). The mtDNA haplotypes are inter-related by single nucleotide differences, and a simple network of haplotype relationships may be constructed. This shows a radiation out of Africa (Fig. 3). Most African isolates have the haplotypes CACC or CGCC which are at the centre of the network, and there is a related African haplotype CGCT which is rare (seen only in NF54 and one Nigerian isolate). Outside of Africa, most isolates have continent-specific types which are at the termini of the network (CATC in 7G8 and most Brazilian

field isolates; TGCC in the Malaysian field isolates and cultured Thai isolate K1), and the remainder have the expected putative ancestral types (CACC in two Brazilian field isolates, and CGCC which was seen in Southeast Asian cultured isolates CAMP and T9/96).

4. Discussion This study gives strong molecular evidence for a recent African origin of P. falciparum, and subsequent colonisations of Southeast Asia and South America. The nucleotide diversity throughout the mt genome of P. falciparum (0.03% overall, 0.04%

Fig. 3. Geographical distribution of P. falciparum mtDNA haplotypes, shown by their nucleotides at each of the 4 polymorphic sites (positions 772, 1692, 4179, 4952). Solid lines indicate single nucleotide differences between haplotypes. Dashed arrows indicated putative colonisation events of haplotypes from Africa into Southeast Asia and South America. The continental distribution of haplotypes was derived from field isolates (Table 2), and from putative origins of cultured isolates which have been fully sequenced (see text of Section 3).

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D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171

at synonymous positions) is approximately two orders of magnitude lower than the divergence with P. reichenowi (2.41% overall, 12.01% at synonymous positions). The best estimate of the split between P. falciparum and P. reichenowi is 5 – 10 million years ago [6], so it is likely that existing P. falciparum mt DNA sequences derive from a common origin in the order of 50 000 years ago. The precision of this estimation is limited by the very low polymorphism, but it is concordant with an estimate based on low synonymous nucleotide polymorphism in some nuclear genes [5] which allowed for the possibility of an even more recent date. It should be noted that the result obtained here, which compares intra- and inter-species divergence, is robust to any effect of high AT content and biased codon usage in the mitochondrial genome, as this is similar in all Plasmodium species studied [20]. The relationship among mt DNA haplotypes in P. falciparum, and their geographical distribution, shows a clear radiation out of Africa. The two common African types are central in the simple haplotype tree, which strongly suggests that they are ancestral to the other types. The branch termini consist of one type common in Southeast Asia, an unrelated type common in South America, and a rare type in Africa. This indicates that Southeast Asia and South America were colonised separately by P. falciparum, a conclusion which is further supported by the detection of the respective putative ancestral types in these regions (Fig. 3). The data are not incompatible with a hypothesis that P. falciparum first emerged in early agricultural communities in Africa [21], aided by recent evolution in larval ecology of the African mosquito vector species Anopheles gambiae sensu stricto [22]. Several issues present themselves for future research. It remains possible that an intensive sequencing survey in Africa might reveal more mtDNA diversity than that seen among the complete sequences to date, which were mainly from nonAfrican parasites. Accurate dating of colonisation events, or tracing the spread of P. falciparum within each continent, will probably require more sequence diversity than that afforded by the study of the mt genome. It is complex to study ancestral relationships among single nucleotide polymorphism haplotypes in the nuclear genome due to frequent

recombination [17,23], so the 35-kb circular plastid genome [24] or rapidly mutating microsatellite loci [23,25] could be the best markers for such studies. It is surprising that the terminal mtDNA haplotypes should have become so common in South America and Southeast Asia, in comparison to their putative ancestors, and it is unknown if positive selection has operated on these haplotypes. The almost complete partitioning of mtDNA haplotypes between continents is similar to the distribution of alleles of a gamete surface protein gene Pfs48 /45 [26], but contrasts with the more broadly distributed alleles of asexual blood-stage antigen genes surveyed so far [27,28]. This might indicate continent-specific divergent selection on both the mtDNA and Pfs48 / 45, or balancing selection on most asexual bloodstage antigen genes surveyed. In strong contrast with the low overall diversity in P. falciparum mtDNA, and in some nuclear genes [5], there are very divergent and apparently ancient alleles in several antigen genes [7–10]. Thus, it is likely that some of the antigen polymorphisms have been maintained for a long time by balancing selection [29,30], which should encourage investigation of their alleles as potential targets of acquired immune responses [27,31]. Acknowledgements Financial support was provided by The Wellcome Trust (grant ref. 055487), the European Commission, and the University of London Central Research Fund. Assistance in sample collection and transport was provided by Olumide Ogundahunsi, Janet Cox-Singh, Ricardo Machado, and Chris Drakeley. Helpful comments on the manuscript were given by Tim Anderson, David Goldstein and Chung-I Wu. References [1] Ewald PW. Evolution of Infectious Disease. Oxford: Oxford University Press, 1994. [2] Weatherall DJ. From genotype to phenotype: genetics and medical practice in the new millennium. Phil Trans R Soc Lond B 1999;354:1995– 2010. [3] Snow RW, Craig M, Deichmann U, Marsh K. Estimating mortality, morbidity and disability due to malaria among

D.J. Conway et al. / Molecular and Biochemical Parasitology 111 (2000) 163–171















Africa’s non-pregnant population. Bull WHO 1999;77:624– 40. Flint J, Harding RM, Boyce AJ, Clegg JB. The population genetics of the haemoglobinopathies. Bailliere’s Clin Haematol 1993;6:215–62. Rich SM, Licht MC, Hudson RR, Ayala FJ. Malaria’s eve: evidence of a recent population bottleneck throughout the world populations of Plasmodium falciparum. Proc Natl Acad Sci USA 1998;95:4425–30. Escalante AA, Ayala FJ. Phylogeny of the malarial genus Plasmodium, derived from rRNA gene sequences. Proc Natl Acad Sci USA 1994;91:11373–7. Hughes AL. Positive selection and interallelic recombination at the Merozoite Surface Antigen-1 (MSA-1) locus of Plasmodium falciparum. Mol Biol Evol 1992;9:381–93. Dubbeld MA, Kocken CHM, Thomas AW. Merozoite surface protein 2 of Plasmodium reichenowi is a unique mosaic of Plasmodium falciparum allelic forms and speciesspecific elements. Mol Biochem Parasitol 1998;92:187–92. Okenu DMN, Thomas AW, Conway DJ. Allelic lineages of the merozoite surface protein 3 gene in Plasmodium reichenowi and Plasmodium falciparum. Mol Biochem Parasitol 2000;109:185–8. Kochen CHM, Narum DL, Massougbodji A, Ayivi B, Dubbeld MA, van der Wel A, et al. Molecular characterization of Plasmodium reichenoni apical membrane antigen-1 (AMA-1) comparison with P. falciparum AMA-1, and antibody-mediated inhibition of red cell invasion. Mol Biochem Parasitol 2000;109:147–56. Avise JC, et al. Intraspecific phylogeography: the mitochondrial DNA bridge between population genetics and systematics. Ann Rev Ecol Syst 1987;18:489–522. Feagin JE, Werner E, Gardner MJ, Williamson DH, Wilson RJM. Homologies between the contiguous and fragmented rRNAs of the two Plasmodium falciparum extrachromosomal DNAs are limited to core sequences. Nucleic Acids Res 1992;20:879 – 87. Creasey AM, Ranford-Cartwright LC, Moore DJ, Williamson, Wilson RJM, Walliker D, Carter R. Uniparental inheritance of the mitochondrial gene cytochrome b in Plasmodium falciparum. Curr Genet 1993;23:360–4. Preiser PR, Wilson RJM, Moore PW, McCready S, Hajibagheri MAN, Blight KJ, et al. Recombination associated with replication of malarial mitochondrial DNA. EMBO J 1996;15:684–93. Escalante AA, Freeland DE, Collins WE, Lal AA. The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochondrial genome. Proc Natl Acad Sci USA 1998;95:8124–9. Rozas J, Rozas R. DnaSP version 3: an integrated program for molecular population genetics and molecular evolution analysis. Bioinformatics 1999;15:174–5. Conway DJ, Roper C, Oduola AMJ, Arnot DE, Kremsner PG, Grobusch MP, et al. High recombination rate in natural populations of Plasmodium falciparum. Proc Natl Acad Sci USA 1999;96:4506 – 45011.


[18] Alloueche A, Silveira H, Conway DJ, Bojang K, Doherty T, Cohen J, et al. High throughput sequence typing of T-cell epitope polymorphisms in Plasmodium falciparum circumsporozoite protein. Mol Biochem Parasitol 2000;106:273 – 82. [19] Weir BS, Cockerham CC. Estimating F statistics for the analysis of population structure, Evolution 1984; 1358 – 70. [20] McIntosh MT, Srivastava R, Vaidya AB. Divergent evolutionary constraints on mitochondrial and nuclear genomes of malaria parasites. Mol Biochem Parasitol 1998;95:69 – 80. [21] Wiesenfeld SL. Sickle-cell trait in human biological and cultural evolution. Science 1967;157:1135– 40. [22] Coluzzi M. The clay feet of the malaria giant and its African roots: hypotheses and inferences about origin, spread and control of Plasmodium falciparum. Parasitologia 1999;41:277 – 85. [23] Su X-Z, et al. A genetic map and recombination parameters of the human malaria parasite Plasmodium falciparum. Science 1999;286:1351– 3. [24] Wilson RJM, et al. Complete gene map of the plastid-like DNA of the malaria parasite Plasmodium falciparum. J Mol Biol 1996;261:155 – 72. [25] Anderson TJC, Su X-Z, Bockaire M, Lagog M, Day KP. Twelve microsatellite markers for characterisation of Plasmodium falciparum from finger prick blood samples. Parasitology 1999;119:113 – 25. [26] Drakeley CJ, Duraisingh MT, Povoa M, Conway DJ, Targett GAT, Baker DA. Geographical distribution of a variant epitope of Pfs48 /45, a Plasmodium falciparum transmission-blocking vaccine candidate. Mol Biochem Parasitol 1996;81:253 – 7. [27] Conway DJ. Natural selection on polymorphic malaria antigens and the search for a vaccine. Parasitol Today 1997;13:26 – 9. [28] Anderson TJC, Day KP. Geographical structure and sequence evolution as inferred from the Plasmodium falciparum S-antigen locus. Mol Biochem Parasitol 2000;106:321 – 6. [29] Hughes MY, Hughes AL. Natural selection on Plasmodium surface proteins. Mol Biochem Parasitol 1995;71:99 – 113. [30] Escalante AA, Lal AA, Ayala FJ. Genetic polymorphism and natural selection in the malaria parasite Plasmodium falciparum. Genetics 1998;149:189 – 202. [31] Conway DJ, Cavanagh DR, Tanabe K, Roper C, Mikes ZS, Sakihama N, et al. A principal target of human immunity to malaria identified by molecular population genetic and immunological analyses. Nat Med 2000;6:689 – 92. [32] Ji Y, Mericle BL, Rehkopf DH, Anderson JD, Feagin JE. The Plasmodium falciparum 6 kb element is polycistronically transcribed. Mol Biochem Parasitol 1996;81:211 – 23. [33] Rehkopf DH, Gillespie DE, Harrell MI, Feagin JE. Transcriptional mapping and RNA processing of the Plasmodium falciparum mitochondrial mRNAs. Mol Biochem Parasitol 2000;105:91 – 103. [34] Feagin JE, Mericle BL, Werner, Morris M. Identification of additional rRNA fragments encoded by the Plasmodium falciparum 6 kb element, Nucleic Acids Res 1997;438 – 46.

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