Rapid genotyping of loci involved in antifolate drug resistance inPlasmodium falciparum by primer extension

Shalini Naira,*, Alan Brockmanb,c, Lucy Paiphunb, Francois Nostenb,c,d, Tim J.C. Andersona

aSouthwest Foundation for Biomedical Research (SFBR), PO Box 760549, San Antonio, TX 78245, USAbShoklo Malaria Research Unit (SMRU), Mae Sot, Tak, Thailand

cFaculty of Tropical Medicine, Mahidol University, Bangkok, ThailanddNuffield Department of Medicine, John Radcliffe Hospital, Oxford, UK

Received 10 January 2002; received in revised form 28 February 2002; accepted 28 February 2002

Abstract

Current methods used to genotype point mutations in Plasmodium falciparum genes involved in resistance to antifolate drugs include

restriction digestion of PCR products, allele-specific amplification or sequencing. Here we demonstrate that known point mutations in

dihydrofolate reductase and dihydropteroate synthase can be scored quickly and accurately by single-nucleotide primer extension and

detection of florescent products on a capillary sequencer. We use this method to genotype parasites in natural infections from the Thai-

Myanmar border. This approach could greatly simplify large-scale screening of resistance mutations of the type required for evaluating and

updating antimalarial drug treatment policies. The method can be easily adapted to other P. falciparum genes and will greatly simplify

scoring of point mutations in this and other parasitic organisms. q 2002 Australian Society for Parasitology Inc. Published by Elsevier

Science Ltd. All rights reserved.

Keywords: Plasmodium falciparum; Dihydrofolate reductase; Dihydropteroate synthase; Primer extension; Thailand

1. Introduction

Numerous association studies, elegant transfection

experiments, and expression of Plasmodium falciparum

genes in yeast or Escherichia coli provide overwhelming

evidence that specific mutations in dihydrofolate reductase

(dhfr) and dihydropteroate synthase genes (dhps) confer

resistance to antifolate drugs such as sulfadoxine and pyri-

methamine (Cowman et al., 1988; Sirawaraporn et al., 1990;

Foote et al., 1990; Plowe et al., 1997; Khan et al., 1997;

Cortese and Plowe, 1998; Triglia et al., 1998; Diourte et al.,

1999; Doumbo et al., 2000; Basco et al., 2000; Nagesha et

al., 2001; Hankins et al., 2001). Furthermore, these muta-

tions are correlated with both in vivo and in vitro drug

resistance in many studies (Kublin et al., 1998; Basco et

al., 2000; Nzila et al., 2000). As a consequence, molecular

characterisation of these mutations can provide a simple

way to evaluate the efficacy of this class of drugs and

field surveys may be used to evaluate suitable antimalarial

treatment strategies (Plowe et al., 1995; Wang et al., 1997).

A variety of methods are currently used for molecular

characterisation of mutations in dhfr and dhps. These

include sequencing (Basco et al., 2000), allele-specific

amplification (Zolg et al., 1990; Gyang et al., 1992; Wang

et al., 1995), and restriction analysis of polymerase chain

reaction (PCR) products (Eldin de Pecoulas et al., 1995;

Duraisingh et al., 1998). These methods are either expensive

(sequencing) or cumbersome and time consuming (allele-

specific amplification or restriction digestion). For example,

allele-specific amplification requires separate amplification

reactions with oligonucleotides specific for the alternative

bases at each one of the polymorphic bases to be scored,

while restriction digest approaches involves electrophoresis

of multiple digestion products.

A number of new methods are now available for rapid

automated scoring of single nucleotide polymorphisms

(SNPs) (Kwok, 2000; Syvanen, 2001).We apply one of

these – primer extension – to genotype mutations at dhps

and dhfr. In this technique, mutations are detected by fluor-

escent single base extension of primers designed adjacent to

polymorphic sites (Pastinen et al., 1996; Tully et al., 1996).

An attractive feature of this method is that multiple muta-

tions can be genotyped in a single reaction and scored in a

single lane of a gel or a single capillary on an automated

International Journal for Parasitology 32 (2002) 852–858

0020-7519/02/$20.00 q 2002 Australian Society for Parasitology Inc. Published by Elsevier Science Ltd. All rights reserved.

PII: S0020-7519(02)00033-4

www.parasitology-online.com

* Corresponding author. Tel.: 11-210-258-9642; fax: 11-210-670-3344.

E-mail address: snair@darwin.sfbr.org (S. Nair).

sequencer. As a consequence, multiple mutations can be

genotyped in hundreds of samples in a single day resulting

in considerable savings of time and money. We demonstrate

the efficacy of this approach by genotyping P. falciparum

infections from the Thailand-Myanmar border.

2. Materials and methods

2.1. Genotyping strategy

We used a primer extension method to detect mutations in

dhfr and dhps. The essence of this method is very simple.

The segment of DNA containing the polymorphic sites is

first amplified by standard PCR methods. Oligos designed

with their 3 0 base adjacent to the polymorphic base are then

enzymatically extended by a single fluorescent dideoxy

nucleotide (ddNTP). Since each of the four ddNTPs are

labelled with different colours, the extended oligos are

labelled in different colours depending on the base present

at the targeted polymorphic site. The extended oligos can

then be size separated by electrophoresis and the label

colour determined. Multiple mutations can be genotyped

in a single reaction by designing oligos of different lengths.

Detailed protocols are described below.

Genotyping was performed using the ABI PRISM SNaP-

shote Multiplex Kit (Applied Biosystems) and scored on an

ABI 3100 capillary sequencer using GENESCAN and

GENOTYPER software. Following initial amplification of

template DNA (25 ml reaction), unincorporated nucleotides

(dNTPs) and excess primers were removed by digestion (1 h

378C, 15 min 758C) with 5 U Shrimp Alkaline Phosphatase

(Amersham) and 2 U Exonuclease 1 (Amersham). Primers

for the SNaPshot reactions were designed one base short of

the SNP to be typed. In the extension reaction (or SnaPshot

reaction) each primer binds to the template in the presence

of fluorescently labelled ddNTPs and Amplitaq DNA Poly-

merase and the primers are extended by one fluorescent

nucleotide at their 3 0 ends. Following removal of excess

fluorescently labelled ddNTPs by Shrimp Alkaline Phos-

phatase digestion (1 U enzyme in 5 ml reaction, 1 h, 378C;

15 min 758C), the fluorescently labelled products generated

by the SNaPshot reaction (0.05–0.5 ml) were electrophor-

esed in a ABI 3100 capillary sequencer together with 0.5 ml

Genescane-120 LIZe labelled size standards (15–120 bp)

(Applied Biosystems) and 9 ml HiDi Formamide (Applied

Biosystems). More than one mutation can be typed by

designing primers with tails of different lengths. We geno-

typed five point mutations encoding five amino acid changes

in dhfr (codons 16, 51, 59, 108 and 164) and six point

mutations encoding five amino acid changes in dhps

(codon 436 containing two point mutations, codons 437,

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858 853

Table 1

Primers and amplification conditions for genotyping dhfr and dhps

Primer ID Sequence (5 0 to 3 0)

dhfr primers (template)

dhfr-f ACGTTTTCGATATTTATGC

dhfr-r TCACATTCATATGTACTATTTATTC

dhfr primers (SNaPshot)

dhfr51f AGGAGTATTACCATGGAAATGTA

dhfr16r ctCTCATTTTTGCTTTCAACCTTACAACAT

dhfr108f ctgactACAAAATGTTGTAGTTATGGGAAGAACAA

dhfr164r gactgactgactAATTCTTGATAAACAACGGAACCTCCTA

dhfr59r gactgactgactgactTGATTCATTCACATATGTTGTAACTGCAC

dhps primers (template)

dhps-f GTTGAACCTAAACGTGCTGT

dhps-r TTCATCATGTAATTTTTGTTGTG

dhps primers (SNaPshot)

dhps613r TTGATCATTCATGCAATGGG

dhps540f gactGAGGAAATCCACATACAATGGAT

dhps581r TAAGAGTTTAATAGATTGATCATGTTTCTTC

dhps436(1)f gactgactAGTGTTATAGATATAGGTGGAGAATCC

dhps436(2)r gactgactgactgactTGGATTAGGTATAACAAAAAGGAICA

dhps437r gactgactgactgactgactTTTTTGGATTAGGTATAACAAAAGGA

PCR conditions for amplification of both dhfr and dhps were identical. The reaction mix (25 ml) contained 2 ng genomic DNA, 200 mM each dNTP, 1.5

mM Mg21, 1 £ PCR buffer, 0.25 mM each primer, 0.6 U Taq (Takara Shuzo Co.) while cycling conditions were as follows: (948C, 2 min, (948 C, 20 s; 458C, 20

s; 658C, 30 s) £ 30 cycles. The SNaPshot reaction for dhfr (5 ml) contained 2.5 ml reaction mix, 1.5 ml amplified template from the initial PCR reaction, 0.5 ml

primer pool at a concentration of 0.2 mM for each primer, and 0.5 ml dH20. SNaPshot reaction for dhps was the same except that one primer dhps 437r was at a

concentration of 0.4 mM in the primer pool. Reaction conditions for the SNaPshot reaction were as shown in the ABI PRISM SNaPshote Multiplex Kit (968 C,

10 s; 508C, 5 s, 608C, 30 s) £ 25. (gact)n tails added to the SnaPshot primers are shown in lower case, while Inosine (I) bases are incorporated in primers where

the complimentary DNA sequence may be polymorphic.

540, 581 and 613). PCR amplification conditions and primer

sequences are shown in Table 1, while the positions and

orientation of the oligos used in the extension reaction are

shown in Fig. 1.

2.2. Field samples

We collected P. falciparum-infected blood samples with

.0.5% parasitaemia from patients visiting the malaria

clinic at Mawker-Thai on the Thai-Myanmar border

between December 1998 and August 2001. The collection

protocols were approved by the Ethical Committee of the

Faculty of Tropical Medicine, Mahidol University, Bang-

kok, and by the Institutional Review board at the University

of Texas Health Science Center at San Antonio. Parasite

DNA was prepared by Phenol/Chloroform extraction of

whole blood, following removal of buffy coats. Two nano-

grams of DNA were used in each PCR reaction.

2.3. Sequencing

To investigate the reliability of genotyping by primer

extension we sequenced 10 dhfr and eight dhps alleles

from single clone infections from Mawker-Thai in Thailand

for comparison with data generated by primer extension.

These single clone infections were identified by analysis

of multiple microsatellite loci (data not shown). The

template primers shown in Table 1 were used to amplify

the fragments of interest and these were sequenced directly

using the ABI PRISM BigDyee Terminator Cycle sequen-

cing Ready Reaction kit (Applied Biosystems). As addi-

tional controls, we genotyped a variety of laboratory

parasite lines (3D7, HB3, Dd2, VS/1, FCR3) obtained

from MR4/ATCC (Manassus, VA, USA) with known dhfr

and dhps sequence.

2.4. Analysis

Alternate alleles at each polymorphic site are labelled

with different dyes and migrate at a slightly different posi-

tion. The differing migration pattern of different dye labels

also allows samples containing different dhfr or dhps alleles

to be scored with the GENOTYPER software. We compared

the haplotype frequencies generated in this study with those

from other studies of antifolate resistance in Thailand using

F-statistics (Cockerham and Weir, 1984). These were calcu-

lated using the program FSTAT (Goudet, J. 2000. FSTAT, a

program to estimate and test gene diversities and fixation

indices (version 2.9.1). Available from http://www.unil.ch/

izea/softwares/fstat.html) and significance of measures of

differentiation were estimated by permutation.

3. Results and discussion

In the first section we describe and discuss the results of

the field survey. In the second we highlight the advantages

and disadvantages of genotyping by primer extension.

3.1. Field survey

We genotyped 174 malaria-infections for dhfr and 168 for

dhps. Examples of peak profiles for both field samples and

controls are shown in Fig. 2, interpretation of peak colours

and product lengths is described in Table 2, and a summary

of amino-acid haplotypes is shown in Table 3. Nineteen

infections contained multiple clones. In these samples,

peaks of different colours were observed for one or more

base positions at either of the two loci (Fig. 2). Of these 19

samples, three samples contained multiple alleles at both

loci examined. The total number of samples for which

unambiguous haplotypes could be established was 163 for

dhfr and 157 for dhps. We conducted this work primarily to

demonstrate the effectiveness of primer extension based

genotyping. However, three features of the data generated

are of particular interest and are discussed below:

† All dhfr and dhps alleles examined contained mutations

at one or more of the amino-acid position examined. At

dhfr we found four different alleles, containing between

two and four amino-acid changes considered to confer

resistance (Table 3), with the commonest allele contain-

ing Ala-16/Ile-51/Arg-59/Asn-108/Leu-164. Similarly,

at dhps, we observed five different alleles containing

between two and three amino-acid changes considered

to confer resistance. The commonest dhps allele was

Ser-436/Gly-437/Glu-540/Gly-581/Ala-613. Parasites in

Thailand are known to show high levels of resistance to

sulfadoxine-pyrimethamine (White, 1992). However, the

amino-acids profiles from Mawker-Thai suggest a higher

level of resistance than reported elsewhere in Southeast

Asia or in other countries. The high prevalence of muta-

tions thought to confer resistance is particularly interest-

ing, since sulfadoxine/pyrimethamine was abandoned in

this part of Thailand in the early 1980s. Hence, for over

20 years resistant alleles of dhfr and dhps have been

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858854

Fig. 1. Schematic diagram showing the position and orientation of SNaP-

shot primers for: (a) dhfr; and (b) dhps. The 5 0 ends of the primers are

upturned to indicate (gact)n tails (Table 1).

maintained in the apparent absence of antifolate drug

selection. This may indicate that parasites bearing resis-

tant alleles suffer little or no fitness reduction relative to

wild-type parasites in the absence of selection, or that

additional compensatory mutations have restored fitness.

Both these possibilities are troubling, since they suggest

that reversion to susceptibility is unlikely to occur.

However, it is possible that sulfadoxine-pyrimethamine

is still used in Myanmar, and Fansimef (a combination of

sulfadoxine-pyrimethamine and mefloquine) which was

officially abandoned in 1994, is sometimes still used in

Thailand. One further possible explanation is that wide-

spread treatment with the antibiotic trimethoprim-sulfa-

methoxazole helps maintain high levels of resistance to

sulfadoxine-pyrimethamine (Iyer et al., 2001).

† Plowe et al. (1997) examined dhfr and dhps haplotypes in

parasites from Mali, Kenya, Malawi and Bolivia. At

dhps, they rarely observed Ala-436 and Gly-437 in the

same haplotype, and never observed Ala-436 with either

the Glu-540 or Gly-581 mutations. They therefore

suggested that such combinations may have deleterious

effects on enzyme function. In this study the Ala-436/

Gly-437 combination was common, occurring in 35

infections (22%) examined. Furthermore, Ala-436 was

combined with Glu-540 in 32 (20%) of samples. It there-

fore seems unlikely that Plowe et al. (1997) explanation

can explain the absence of these haplotypes in Africa and

Bolivia. Ala-436 was never observed with Gly-581,

supporting their observations. However, this combina-

tion was found in 1/10 Thai samples sequenced by Triglia

et al. (1997), demonstrating that parasites bearing Ala-

436/Gly-581 are viable.

† We compared our results with those of Biswas et al.

(2000) who examined 50 malaria infections from Thai-

land (from Trad, Tak, Kanchanaburi and Nongkai). This

mixed sample showed greater diversity of haplotypes

than in this study (seven dhfr genotypes as opposed to

four in this study; 10 dhps haplotypes as opposed to five

in this study). To examine differentiation between these

two population samples we used F-statistics. We

observed dramatic differentiation between the two popu-

lation samples (dhfr, FST ¼ 0:34, P , 0:0002; dhps:

FST ¼ 0:456, P , 0:0002). This high level of differentia-

tion is likely to have resulted from the recent evolution of

drug resistance and differing levels of antifolate drug

selection in different regions. On a practical note, such

dramatic differentiation cautions against designing

national health policies on the basis of small regional

samples of parasites. Rational drug treatment policies

may best be designed at the local level by monitoring

prevalence of resistance mutations. The rapid genotyping

methods described here may help make such surveillance

possible in African countries where antifolate resistance

is emerging, since many thousands of parasites collected

from different locations can be genotyped with relative

ease.

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858 855

Fig. 2. Electropherograms generated from the Plot View window of the

GENOTYPER program showing peak profiles generated for dhfr and dhps.

Arrows indicate the positions of the peaks amplified for each of the muta-

tions scored. Horizontal bars indicate the positions of alternate alleles at a

particular codon. These have only been shown for bases that are poly-

morphic in this figure (codons 51 and 164 for dhfr and 436(1), 540 and

581 for dhps). The green, blue, red and black peaks indicate the ddNTP

added during primer extension. Interpretation of the peak profiles in terms

of codon and amino acid mutations is explained in Table 2. Panel (a) dhfr

genotyping: FS1-3 are field isolates from Mawker–Thai, while K1 is a

laboratory clone. (b) dhps genotyping: FS1-2 are field samples. FS1

contains a single dhps allele, while FS2 contains at least two clones with

polymorphism at codon positions 540, 581 and 436. HB3 is a laboratory

clone.

3.2. Pros and cons of primer extension methods

3.2.1. Accuracy

Direct sequencing of 10 single clone infections from

Thailand for dhfr and eight clones for dhps revealed two

polymorphic codons in dhfr (51, 164) and three in dhps (436

(first nucleotide position), 540, 581). No additional poly-

morphic sites were observed in these sequences. As an addi-

tional control we examined haplotype profiles from

laboratory isolates with known dhfr (3D7, FCR3 and VS/

1) and dhps (3D7, HB3, Dd2) sequences. This was done to

examine the reliability of base calling for mutations for sites

that were not polymorphic in the field samples (codons 16,

59 and 108 in dhfr, and codon 436 (second nucleotide posi-

tion), 437 and codons 613 in dhps). This combination of

sequenced field samples and laboratory clones provided

controls for polymorphism at all sites assayed. In all

cases, SNaPshot haplotype profiles of both sequenced

laboratory clones and laboratory isolates (Table 2) were

identical to predictions. This demonstrates that the peaks

generated result from SNaPshot primers annealing in the

correct position in the sequence, and gives us confidence

in the data generated.

3.2.2. Efficiency

Restriction digestion and allele specific amplification

methods have the virtue of simplicity. These methods do

not require complicated equipment and therefore are ideal

for remote field laboratories. The primer extension methods

described here require the use of an automated sequencer

and are not ideally suited to such remote field laboratories.

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858856

Table 2

Interpretation of SNaPshot data for dhfr and dhps

(a) dhfr

Amino acid residue 51 16 108 164 59

Actual fragment length (bp) 24 31 36 41 46

Observed fragment length (bp)a 27–28 31–32 38 43–44 47–48

Amino acid change Asn ! Ile Ala ! Val Ser ! Asn/Thr Ile ! Leu Cys ! Arg

Nucleotide change AAT ! ATT GCA ! GTA AGC ! AAC/ACC ATA ! TTA TGT ! CGT

Colour changeb Green ! red Blue ! green Blue ! green/black Red ! green Green ! blue

Predicted peak colors (controls)

3D7 Green Blue Blue Red Green

FCR3 Green Green Black Red Green

VS/1 Red Blue Green Green Blue

(b) dhps

Amino acid residue 613 540 581 436(1)/436(2) 437

Actual fragment length (bp) 21 28 32 36/41 46

Observed fragment length (bp)a 25–26 29–30 33–34 38–39/44–45 49–50

Amino acid change Ala ! Thr/Ser Lys ! Glu Ala ! Gly Ser ! Ala/Phe Ala ! Gly

1) TCT ! GCT

Nucleotide change GCC ! ACC/TCC AAA ! GAA GCG ! GGG 2) TCT ! TTT GCG ! GGG

1) red ! blue

Colour changeb Black ! red/green Green ! blue Blue ! black 2) blue ! green Blue ! black

Predicted peak colours (controls)

3D7 Black Green Blue Red/blue Black

Hb3 Black Green Blue Red/blue Blue

Dd2 Green Green Blue Red/green Black

a Black, green, blue and red peaks result from primer extension with ddCTP-dTAMRAe, ddATP-dR6G, ddGTP-dR110, and ddTTP-dROXe, respectively.b Observed length is measured with reference to labelled size standard. Observed length may differ from actual length of fragments since fluorescent labelled

ddNTPs alter migration patterns.

Table 3

Haplotypes observed from Mawker–Thaia

(a) dhfr (n ¼ 163)

16 51 59 108 164 Haplotype frequency

Ala Ile Arg Asn Leu 0.72

Ala Asn Arg Asn Leu 0.11

Ala Ile Arg Asn Ile 0.11

Ala Asn Arg Asn Ile 0.06

(b) dhps (n ¼ 157)

436 437 540 581 613 Haplotype frequency

Ser Gly Glu Gly Ala 0.68

Ala Gly Glu Ala Ala 0.20

Ser Gly Lys Gly Ala 0.08

Ala Gly Lys Ala Ala 0.02

Ser Gly Glu Ala Ala 0.01

a Polymorphic amino-acid residue numbers are shown along the top of

each table, while amino acids conferring resistance are shown in bold.

Haplotype frequencies are calculated only from infections containing single

alleles.

However, the speed at which data can be generated makes

such methods ideal for use in central laboratories, and for

use in large survey projects. The dhps and dhfr genotypes

generated in this paper were genotyped in four 96-well

microtitre plates in less than 2 days work with an

ABI3100 capillary sequencer. There is no reason why

several thousand infections could not be genotyped within

a week using 384 well plates. In comparison, generation of

data on this scale using conventional restriction digest or

allele specific amplification methods would take consider-

ably longer. Are large-scale surveys necessary? For asses-

sing drug resistance mutations in a local area, quite small

numbers of samples may be sufficient. However, the

evidence presented here of strong local differentiation in

dhfr and dhps allele frequencies suggest that quite fine-

scale regional sampling may be necessary to formulate

sensible treatment policies. Furthermore, the rapid emer-

gence of antifolate resistance (Doumbo et al., 2000; Nzila

et al., 2000) means that parasite populations may need to be

sampled continually to monitor the spread of resistance.

3.2.3. Cost

The SNaPshot kit containing premixed fluorescent

ddNTPs and Taq currently costs ,USD1.05 per reaction

(for 5 ml reactions, as used here). The additional costs of

Shrimp Alkaline Phosphatase, Exonuclease 1, microtitre

plates, and the running costs of sequencer bring the total

cost per gene to approximately USD2.00, or USD4.00 for

analysing both genes from a single sample (USD0.36 per

mutation). The method is not as cheap as restriction diges-

tion or allele-specific amplification in material terms.

However, the speed and efficiency may result in consider-

able savings in labour costs and time.

3.2.4. Template quality

In this study we used parasite infections with .0.5%

parasitaemia, and high quality DNA was prepared by

Phenol/Chloroform extraction. In many field studies

finger-prick blood samples are collected and parasitaemias

may be very low, resulting in considerable variation in the

quality and concentration of P. falciparum DNA present in

the initial PCR reaction. In this case, increasing the number

of cycles to 40 in the initial PCR reaction generates suffi-

cient template for the SNaPshot reaction (Nair, unpublished

data). Alternatively, for samples with very small amounts of

parasite DNA, or badly sheared DNA a semi-nested ampli-

fication strategy (Anderson et al., 1999) may be used to

generate sufficient template for the primer extension reac-

tion.

3.2.5. Multiple infections

Multiple clones of P. falciparum are frequently observed

within infected blood samples, and this complicates scoring

of polymorphisms and assessment of population allele

frequencies (Hill and Babiker, 1995). Multiple infections

can be fairly easily characterised by primer extension,

since oligonucleotides containing ddNTPs labelled with

different dyes show different migration properties. Hence,

in a mixed infection, polymorphic bases are visible as two

peaks that are slightly offset (Fig. 2). We found 19 infections

with more than one peak at any one of the variable bases in

either dhps or dhfr. Infections showing multiple peaks at

sites in dhfr are more likely to be polymorphic for sites in

dhps than expected by chance (x2c ¼ 5:53, df ¼ 1,

P , 0:025). This feature of these data gives us considerable

confidence in our ability to score multiple infections.

One caveat should be mentioned for those working in

areas with high levels of multiple infections. Primers

extended with different bases, and therefore different dyes,

may not necessarily have identical peak height. As a conse-

quence, peak heights may not provide an unbiased way to

identify the predominant clone within a blood sample, and

scoring predominant peaks may result in biased estimates of

population allele frequencies (see Anderson et al. (1999) for

a discussion of related problems for microsatellite data).

Nevertheless, primer extension does provide a simple

method to assess the prevalence of drug resistance muta-

tions within the infected human population, which is prob-

ably the most important parameter for those interested in

optimising efficacy of drug treatment. The efficiency with

which this method can detect and quantify the composition

of alleles in multiple infections could be addressed by

making artificial mixtures of two parasites with known

alleles. We have not conducted these experiments.

Other modern SNP typing methods could also be used for

genotyping mutations in dhps and dhfr. For example, real-

time PCR methods have been described for scoring the

Ser ! Asn mutation at codon 108 in dhfr (Durand et al.,

2000). The main advantage of primer extension methods

is that multiple mutations can be scored concurrently.

These SNP genotyping methods have many other applica-

tions for work with P. falciparum and other parasitic organ-

isms. Single nucleotide polymorphisms occur at a rate for

approximately 1 per kb in the P. falciparum genome and a

SNP map of Chr3 has now been constructed (Mu et al.,

unpublished). We expect that rapid SNP genotyping meth-

ods such as those utilised here will soon become routine

tools for mapping, population genetics and molecular epide-

miology studies in P. falciparum and other parasites.

Acknowledgements

We thank the staff of the SMRU for assistance with field

collection of blood samples and the patients visiting the field

clinic at Mawker-Thai. The SMRU is part of the Wellcome-

Trust Mahidol University-Oxford Tropical Medicine

Research Program supported by the Wellcome Trust of

Great Britain. FN is a Wellcome Trust Senior Clinical

Fellow. This work was funded by NIH grant AI48071 to

TJCA.

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858 857

References

Anderson, T.J., Su, X.Z., Bockarie, M., Lagog, M., Day, K.P., 1999.

Twelve microsatellite markers for characterisation of Plasmodium

falciparum from finger-prick blood samples. Parasitology 119 (Pt 2),

113–25.

Basco, L.K., Tahar, R., Keundjian, A., Ringwald, P., 2000. Sequence varia-

tions in the genes encoding dihydropteroate synthase and dihydrofolate

reductase and clinical response to sulfadoxine-pyrimethamine in

patients with acute uncomplicated falciparum malaria. J. Infect. Dis.

182, 624–8.

Biswas, S., Escalante, A., Chaiyaroj, S., Angkasekwinai, P., Lal, A.A.,

2000. Prevalence of point mutations in the dihydrofolate reductase

and dihydropteroate synthetase genes of Plasmodium falciparum

isolates from India and Thailand: a molecular epidemiologic study.

Trop. Med. Int. Health 5, 737–43.

Cockerham, C.C., Weir, B.S., 1984. Estimating F-statistics for the analysis

of population structure. Evolution 38, 1358–70.

Cortese, J.F., Plowe, C.V., 1998. Antifolate resistance due to new and

known Plasmodium falciparum dihydrofolate reductase mutations

expressed in yeast. Mol. Biochem. Parasitol. 94, 205–14.

Cowman, A.F., Morry, M.J., Biggs, B.A., Cross, G.A., Foote, S.J., 1988.

Amino acid changes linked to pyrimethamine resistance in the dihydro-

folate reductase-thymidylate synthase gene of Plasmodium falciparum.

Proc. Natl. Acad. Sci. USA 85, 9109–13.

Diourte, Y., Djimde, A., Doumbo, O.K., Sagara, I., Coulibaly, Y., Dicko,

A., Diallo, M., Diakite, M., Cortese, J.F., Plowe, C.V., 1999. Pyrimetha-

mine-sulfadoxine efficacy and selection for mutations in Plasmodium

falciparum dihydrofolate reductase and dihydropteroate synthase in

Mali. Am. J. Trop. Med. Hyg. 60, 475–8.

Doumbo, O.K., Kayentao, K., Djimde, A., Cortese, J.F., Diourte, Y.,

Konare, A., Kublin, J.G., Plowe, C.V., 2000. Rapid selection of Plas-

modium falciparum dihydrofolate reductase mutants by pyrimethamine

prophylaxis. J. Infect. Dis. 182, 993–6.

Duraisingh, M.T., Curtis, J., Warhurst, D.C., 1998. Plasmodium falci-

parum: detection of polymorphisms in the dihydrofolate reductase

and dihydropteroate synthetase genes by PCR and restriction digestion.

Exp. Parasitol. 89, 1–8.

Durand, R., Eslahpazire, J., Jafari, S., Delabre, J.F., Marmorat-Khuong, A.,

di Piazza, J.P., le Bras, J., 2000. Use of molecular beacons to detect an

antifolate resistance-associated mutation in Plasmodium falciparum.

Antimicrob. Agents Chemother. 44, 3461–4.

Eldin de Pecoulas, P., Basco, L.K., Abdallah, B., Dje, M.K., le Bras, J.,

Mazabraud, A., 1995. Plasmodium falciparum: detection of antifolate

resistance by mutation-specific restriction enzyme digestion. Exp. Para-

sitol. 80, 483–7.

Foote, S.J., Galatis, D., Cowman, A.F., 1990. Amino acids in the dihydro-

folate reductase-thymidylate synthase gene of Plasmodium falciparum

involved in cycloguanil resistance differ from those involved in pyri-

methamine resistance. Proc. Natl. Acad. Sci. USA 87, 3014–7.

Gyang, F.N., Peterson, D.S., Wellems, T.E., 1992. Plasmodium falciparum:

rapid detection of dihydrofolate reductase mutations that confer resis-

tance to cycloguanil and pyrimethamine. Exp. Parasitol. 74, 470–2.

Hankins, E.G., Warhurst, D.C., Sibley, C.H., 2001. Novel alleles of the

Plasmodium falciparum dhfr highly resistant to pyrimethamine and

chlorcycloguanil, but not WR99210. Mol. Biochem. Parasitol. 117,

91–102.

Hill, W.G., Babiker, H.A., 1995. Estimation of numbers of malaria clones

in blood samples. Proc. R. Soc. Lond. B Biol. Sci. 262, 249–57.

Iyer, J.K., Milhous, W.K., Cortese, J.F., Kublin, J.G., Plowe, C.D., 2001.

Plasmodium falciparum cross-resistance between trimethoprim and

pyrimethamine. Lancet 358, 1066–7.

Khan, B., Omar, S., Kanyara, J.N., Warren-Perry, M., Nyalwidhe, J., Peter-

son, D.S., Wellems, T., Kaniaru, S., Gitonga, J., Mulaa, F.J., Koech,

D.K., 1997. Antifolate drug resistance and point mutations in Plasmo-

dium falciparum in Kenya. Trans. R. Soc. Trop. Med. Hyg. 91, 456–60.

Kublin, J.G., Witzig, R.S., Shankar, A.H., Zurita, J.Q., Gilman, R.H.,

Guarda, J.A., Cortese, J.F., Plowe, C.V., 1998. Molecular assays for

surveillance of antifolate-resistant malaria. Lancet 351, 1629–30.

Kwok, P.Y., 2000. High-throughput genotyping assay approaches. Pharma-

cogenomics 1, 95–100.

Nagesha, H.S., Din, S., Casey, G.J., Susanti, A.I., Fryauff, D.J., Reeder, J.C.,

Cowman, A.F., 2001. Mutations in the pfmdr1, dhfr and dhps genes of

Plasmodium falciparum are associated with in-vivo drug resistance in

West Papua, Indonesia. Trans. R. Soc. Trop. Med. Hyg. 95, 43–49.

Nzila, A.M., Mberu, E.K., Sulo, J., Dayo, H., Winstanley, P.A., Sibley,

C.H., Watkins, W.M., 2000. Towards an understanding of the mechan-

ism of pyrimethamine-sulfadoxine resistance in Plasmodium falci-

parum: genotyping of dihydrofolate reductase and dihydropteroate

synthase of Kenyan parasites. Antimicrob. Agents Chemother. 44,

991–6.

Pastinen, T., Partanen, J., Syvanen, A.C., 1996. Multiplex, fluorescent,

solid-phase minisequencing for efficient screening of DNA sequence

variation. Clin. Chem. 42, 1391–7.

Plowe, C.V., Djimde, A., Bouare, M., Doumbo, O., Wellems, T.E., 1995.

Pyrimethamine and proguanil resistance-conferring mutations in Plas-

modium falciparum dihydrofolate reductase: polymerase chain reaction

methods for surveillance in Africa. Am. J. Trop. Med. Hyg. 52, 565–8.

Plowe, C.V., Cortese, J.F., Djimde, A., Nwanyanwu, O.C., Watkins, W.M.,

Winstanley, P.A., Estrada-Franco, J.G., Mollinedo, R.E., Avila, J.C.,

Cespedes, J.L., Carter, D., Doumbo, O.K., 1997. Mutations in Plasmo-

dium falciparum dihydrofolate reductase and dihydropteroate synthase

and epidemiologic patterns of pyrimethamine-sulfadoxine use and

resistance. J. Infect. Dis. 176, 1590–6.

Sirawaraporn, W., Sirawaraporn, R., Cowman, A.F., Yuthavong, Y., Santi,

D.V., 1990. Heterologous expression of active thymidylate synthase-

dihydrofolate reductase from Plasmodium falciparum. Biochemistry

29, 10779–85.

Syvanen, A.C., 2001. Accessing genetic variation: genotyping single

nucleotide polymorphisms. Nat. Rev. Genet. 2, 930–42.

Triglia, T., Menting, J.G., Wilson, C., Cowman, A.F., 1997. Mutations in

dihydropteroate synthase are responsible for sulfone and sulfonamide

resistance in Plasmodium falciparum. Proc. Natl. Acad. Sci. USA 94,

13944–9.

Triglia, T., Wang, P., Sims, P.F., Hyde, J.E., Cowman, A.F., 1998. Allelic

exchange at the endogenous genomic locus in Plasmodium falciparum

proves the role of dihydropteroate synthase in sulfadoxine-resistant

malaria. EMBO J. 17, 3807–15.

Tully, G., Sullivan, K.M., Nixon, P., Stones, R.E., Gill, P., 1996. Rapid

detection of mitochondrial sequence polymorphisms using multiplex

solid-phase fluorescent minisequencing. Genomics 34, 107–13.

Wang, P., Brooks, D.R., Sims, P.F., Hyde, J.E., 1995. A mutation-specific

PCR system to detect sequence variation in the dihydropteroate synthe-

tase gene of Plasmodium falciparum. Mol. Biochem. Parasitol. 71, 115–

25.

Wang, P., Lee, C.S., Bayoumi, R., Djimde, A., Doumbo, O., Swedberg, G.,

Dao, L.D., Mshinda, H., Tanner, M., Watkins, W.M., Sims, P.F., Hyde,

J.E., 1997. Resistance to antifolates in Plasmodium falciparum moni-

tored by sequence analysis of dihydropteroate synthetase and dihydro-

folate reductase alleles in a large number of field samples of diverse

origins. Mol. Biochem. Parasitol. 89, 161–77.

White, N.J., 1992. Antimalarial drug resistance: the pace quickens. J. Anti-

microb. Chemother. 30, 571–85.

Zolg, J.W., Chen, G.X., Plitt, J.R., 1990. Detection of pyrimethamine resis-

tance in Plasmodium falciparum by mutation-specific polymerase chain

reaction. Mol. Biochem. Parasitol. 39, 257–65.

S. Nair et al. / International Journal for Parasitology 32 (2002) 852–858858