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Antimicrobial Agents and Chemotherapy, November 2008, p. 3883-3888, Vol. 52, No. 11
0066-4804/08/$08.00+0     doi:10.1128/AAC.00431-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Plasmodium falciparum Strains Harboring Dihydrofolate Reductase with the I164L Mutation Are Absent in Malawi and Zambia Even under Antifolate Drug Pressure

Edwin Ochong,¹ David J. Bell,¹ David J. Johnson,¹ Umberto D'Alessandro,³ Modest Mulenga,⁴ Sant Muangnoicharoen,¹ Jean-Pierre Van Geertruyden,³ Peter A. Winstanley,² Patrick G. Bray,² Stephen A. Ward,¹ and Andrew Owen^2*

Liverpool School of Tropical Medicine, Pembroke Place, Liverpool, L35QA,¹ Department of Pharmacology and Therapeutics, 70 Pembroke Place, University of Liverpool, Liverpool, L69 3GF, United Kingdom,² Department of Parasitology, Prince Leopold Institute of Tropical Medicine, Antwerp, Belgium,³ Tropical Diseases Research Center (TDRC), Ndola, Zambia⁴

Received 1 April 2008/ Returned for modification 26 June 2008/ Accepted 29 July 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The Plasmodium falciparum dihydrofolate reductase (PfDHFR) enzyme is the target of pyrimethamine, a component of the antimalarial pyrimethamine-sulfadoxine. Resistance to this drug is associated primarily with mutations in the Pfdhfr gene. The I164L mutant allele is of particular interest, because strains possessing this mutation are highly resistant to pyrimethamine and to chlorproguanil, a component of chlorproguanil-dapsone. A recent study from Malawi reported this mutation at a prevalence of 4.7% in parasites from human immunodeficiency virus-positive pregnant women by using a real-time PCR method. These observations have huge implications for the use of pyrimethamine-sulfadoxine, chlorproguanil-dapsone, and future antifolate-artemisinin combinations in Africa. It was imperative that this finding be rigorously tested. We identified a number of critical limitations in the original genotyping strategy. Using a refined and validated real-time PCR strategy, we report here that this mutation was absent in 158 isolates from Malawi and 42 isolates from Zambia collected between 2003 and 2005.

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In response to the emergence of chloroquine-resistant Plasmodium falciparum malaria in Africa, drugs targeting parasite folate biosynthesis (antifolates) were deployed as the first-line treatment for uncomplicated malaria. The most commonly used antifolate is pyrimethamine-sulfadoxine (PYR-SDX). PYR targets P. falciparum dihydrofolate reductase (PfDHFR), a key enzyme involved in the NADPH-dependent reduction of dihydrofolate to tetrahydrofolate.

The greatest disadvantage of the use of PYR-SDX has been the rapid evolution of drug-resistant parasites. This is mediated by the sequential acquisition of point mutations in the Pfdhfr and Pfdhps genes (8, 25). With respect to PYR, the single Pfdhfr mutation S108N is responsible for low-level resistance and is followed by an N51I or C59R mutation (resulting in a double mutant). A combination of all three mutations gives rise to the highly PYR resistant triple mutant. A fourth point mutation in Pfdhfr, I164L, is found extensively in Southeast Asia and South America, but there is debate over its existence in Africa (5, 11, 12, 17, 29). The presence of the I164L quadruple mutant confers high-level resistance to PYR, and in Africa this mutation would severely compromise the continued use of PYR-SDX. This mutation would also compromise the use of Lapdap, a combination of chlorproguanil (CPG) and dapsone (DDS) (30), and CPG-DDS-artesunate in Africa. The development and use of these compounds have now been halted due to toxicity in children with glucose-6-phosphate dehydrogenase deficiency.

Many studies have looked for the presence of the I164L mutant allele in Africa; Malawi was the first African country to switch from chloroquine to PYR-SDX in 1993. Using conventional PCR, most studies have not detected this mutation (2, 3, 7, 15, 18, 19, 22-24, 26, 27), but it has been reported at low prevalences in five different African countries.

Of particular interest is the study by Alker et al. (1), who used real-time PCR with fluorescent probes specific for the mutation and reported a 4.7% prevalence in parasites collected from human immunodeficiency virus (HIV)-positive pregnant women in Malawi between 2001 and 2003, a finding that they validated more recently using a heteroduplex tracking assay (9). However, on biological grounds, if the quadruple-mutant alleles were present in reasonable proportions, it is hard to imagine that they would not have been selected to high levels by this time. This is the highest prevalence reported, and it is a high priority for the public health that this finding be further evaluated.

The aims of this study were to confirm and validate the sensitivity, specificity, and reproducibility of the assay reported by Alker et al. (1) and to confirm the presence of the mutant 164 allele in parasites collected from the same location in Malawi. We also tested the hypothesis that sustained antifolate use would have resulted in an increased prevalence of the I164L mutant in subsequent years. Finally, we wanted to determine if treatment failure after treatment with PYR-SDX resulted in the selection of this mutation. The prevalence of the I164L allele was also investigated in clinical isolates from Zambia, a neighboring country with a shorter history of PYR-SDX deployment, and in clinical isolates from the Thailand-Myanmar border in Southeast Asia, an area known to have a high prevalence of the I164L mutation (5, 10).

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Samples. Five Plasmodium falciparum reference strains (K1, 3D7, HB3, Dd2, and V1/S) were selected for optimization and validation of the Pfdhfr I164L mutation-specific assays. K1, 3D7, HB3, and Dd2 are wild type at position 164 (isoleucine), while V1/S is mutant (leucine). The genotypic and phenotypic statuses of the strains were established through PCR-restriction fragment length polymorphism, real-time PCR, and in vitro determination of 50% inhibitory concentrations of PYR, as previously described (13). Further confirmation was performed by DNA sequencing of the region of Pfdhfr encompassing the I164L allele.

Field isolates were obtained from studies conducted in Malawi between 2003 and 2005 (n = 210), in Zambia in 2005 (n = 55), and around the Thailand-Myanmar border in 2005 (n = 50). The Malawian samples were from children less than 5 years old presenting with uncomplicated malaria and were treated as part of a study with PYR-SDX alone or PYR-SDX plus either chloroquine, artesunate, or amodiaquine. The study took place at a government health center 10 km outside of Blantyre, Malawi, where malaria transmission occurs all year round. Parasite isolates were collected prior to treatment and from children who had recurrent parasitemia after treatment. For 57 children, both pre- and posttreatment isolates were available. Genotypic analysis of the msp2 gene showed that approximately one-third of these recurrent parasitemias were due to reinfection and the remainder were due to recrudescence. The pretreatment prevalence of Pfdhfr triple-mutant parasites was 96%, compared to a prevalence of 80% in the parasites described by Alker et al. (1). Details of the Malawian study, which includes Pfdhfr 164 genotyping using a less-sensitive methodology (PCR and allele-specific restriction analysis) have been published elsewhere (4).

The Zambian samples were collected from adults with uncomplicated malaria before treatment as part of a randomized clinical trial with either PYR-SDX or artemether-lumefantrine. The samples were a fair representation of the population, and inclusion/exclusion criteria have been published previously (14, 28). The Thai samples were from adults with P. falciparum malaria before treatment.

Whole blood from the patients in these studies was spotted onto Whatman 3MM filter paper, air dried at room temperature, and stored in individual plastic bags with a desiccant. In addition, for some children in the Malawi study, venous blood was collected in EDTA tubes and stored at –80°C. All the studies contributing samples to this work were conducted under clinical protocols approved by the corresponding institutional review boards.

Isolation and extraction of total DNA. Total genomic DNAs (host and parasite) were extracted either from EDTA-treated whole blood or from blood-spotted filter papers with the QIAamp DNA blood minikit (Qiagen) according to the manufacturer's instructions.

Whole-genome amplification. A high degree of variability in parasitemia and a low recovery rate of parasite genomic DNA from filter paper were observed. Therefore, when the total-parasite DNA concentration was below 10,000 copies per µl, whole-genome amplification by improved primer extension PCR was conducted. This procedure was performed as previously described (6) in order to increase the quantity of DNA and maximize the number of experiments that could be conducted on each sample.

Real-time PCR-based discrimination of Pfdhfr I164L alleles by the method of Alker. In order to assess the sensitivity of the assay, real-time PCR was conducted using the methodology of Alker et al. (1) using a PTC-200 Peltier thermal cycler with a Chromo 4 continuous fluorescence detector (Bio-Rad). This PCR was conducted on reference strains as well as on samples obtained from Malawi that had not undergone whole-genome amplification. The primer and probe sequences are identical to those published previously (1) and were sourced from Applied Biosystems, Cheshire, United Kingdom.

Normalization of Plasmodium DNA by quantification of EF1-. Owing to the high degree of variability in parasite DNA content, it was necessary to normalize the samples according to the parasite DNA copy number. This was achieved by quantification of elongation factor 1 alpha (EF1-) as a marker for the parasite genome copy number. Primers and fluorescent probes specific for the EF1- gene were designed so as to avoid introns (Table 1).

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TABLE 1. All primer and probe sequences utilized in this study

In order to construct a standard curve, DNAs from five reference strains (K1, Dd2, HB3, 3D7, and V1/S) were normalized to 20 ng/µl in a 25-µl final reaction volume with 1x Absolute QPCR mix (ABgene, Surrey, United Kingdom), 0.9 µM EF1- forward and reverse primers, and 0.25 µM EF1- probe. The samples were amplified in a PTC-200 Peltier thermal cycler with a Chromo 4 continuous fluorescence detector. The program consisted of an initial activation at 95°C for 10 min, followed by 45 cycles of denaturation at 92°C for 14 s and annealing/extension at 60°C for 60 s.

Amplicons were checked for the presence of nonspecific products by electrophoresis on a 1% agarose gel. Each amplicon was gel purified by using the Promega Gel Wizard prep kit according to the manufacturer's instructions. The copy number of amplicons was then quantified as described previously (20) and diluted to 1, 10, 100, 1,000, 10,000 and 100,000 copies per µl. EF1- was then quantified alongside clinical samples as described above. Following amplification, the cycle threshold (C_T) was determined for each sample and standard, the data plotted, and the equation of the linear regression used to determine the numbers of copies of parasite DNA in the clinical samples. Parasite DNA was then normalized to 100, 250, 500, 1,000, and 10,000 copies per µl for validation of the assay.

Validation of real-time PCR-based discrimination of Pfdhfr I164L alleles. For quantification of the Pfdhfr I164L DNA, mutant (V1/S) and wild-type (K1) genomes were combined in fixed ratios of 100:0, 99:1, 95:5, 90:10, 75:25, 50:50, 25:75, and 0:100. These standards were then amplified, and the C_T values for the mutant probe were divided by the corresponding C_T values for the wild-type probe. These values were then plotted against log-transformed percentages as a standard curve.

All reactions were carried out in duplicate in a total volume of 25 µl. Each reaction mixture contained 1x Absolute QPCR mix (ABgene), 200 nM each probe, 288 nM forward primer, and 490 nM reverse primer (Table 1). The program consisted of an initial activation at 95°C for 10 min, followed by 45 cycles of denaturation at 92°C for 14 s and annealing/extension at 60°C for 60 s.

Initial experiments were conducted with 100, 250, 500, 1,000, and 10,000 copies per µl of parasite DNA in order to determine the optimum concentration. The intrarun precision of the analyses, expressed as a percentage, was calculated as 100 – [(standard deviation/mean) x 100]. The interrun precision, also expressed as a percentage, was calculated as (calculated log copy number)/(nominal log copy number added) x 100. Determinations were performed using the same amplicon stock solutions. Intrarun precision and accuracy were assessed on six replicates of standards containing 10% and 75% concentrations of the mutant genome. Similarly, interrun precision and accuracy were assessed on six separate runs of standards containing 10% and 75% concentrations of the mutant genome. The limit of detection was defined as the percentage at which the mutant allele could be reliably differentiated from the wild-type allele, and the limit of quantification was defined as values between 90 and 110% for both intra- and interrun accuracy and precision.

For quantification of the percentage of an individual's total parasite population containing the mutant allele, normalized DNAs were amplified as described above. A standard curve was coamplified on each plate, and both standards and samples were assessed at least in duplicate.

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Limitations of the original methodology. In transferring the methodology to our laboratory, a number of problems were encountered. First, a large intersample variability in amplification was observed (Fig. 1A). Second, the wild-type probe was found to bind nonspecifically to the mutant sequence (Fig. 1B), and vice versa (Fig. 1C). This phenomenon could lead to isolates being interpreted as mixed alleles, particularly in Africa, where most infections are polyclonal. The combination of intersample variability and the suboptimal specificity of the probes led us to conclude that the assay was likely to call samples with high parasite DNA concentrations mutant, even if they were actually wild type. A strategy was therefore developed to normalize parasite DNA concentrations (irrespective of total DNA) and to subsequently validate a method for detection of the Pfdhfr I164L mutant by capitalizing on the higher specificity of the mutant probe for the mutant sequence and of the wild-type probe for the wild-type sequence. In order to achieve this, an optimized standard curve was included within every run.

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FIG. 1. Problems experienced with real-time PCR prior to validation. (A) Real-time PCR trace for the mutant allele with Malawian isolates illustrating the high degree of intersample variability. (B) Real-time PCR trace for wild-type and mutant alleles in an isolate (V1S) known to be entirely mutant for this mutation, illustrating the lack of specificity of the wild-type probe. (C) Real-time PCR trace for wild-type and mutant alleles in an isolate (K1) known to be entirely wild type for this mutation, illustrating the lack of specificity of the mutant probe.

Standardization of parasite DNA copy number. The amount of parasite DNA within the total DNA of an experimental sample (containing parasite and host DNAs) was quantified by real-time PCR amplification of EF1- (Fig. 2A). A standard curve generated from purified, quantified prerun amplicons was constructed alongside samples (Fig. 2B). Using this methodology, the median copy number per µl of parasite DNA isolated from Malawian samples was 27,776 (range, 4 to 948,664,062). The median level of parasitemia in the Malawian patients was 66,585 parasites per µl (range, 39 to 644,840 parasites per µl). For log-transformed data, a significant correlation was observed between parasitemia and the copy number of isolated parasite DNA (R² = 0.13; P < 0.0001).

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FIG. 2. Normalization of parasite DNA in order to resolve intersample variability. (A) Representative real-time PCR trace using specific EF1--directed primers and probes. For the template, EF1- amplicons were gel extracted, quantified, and diluted appropriately prior to the real-time PCR assay, as described in Materials and Methods. (B) Representative standard curve for quantification of EF1-. Samples of DNA from clinical isolates underwent quantification of EF1-, and CT values were read from this curve in order to normalize for the copy number of the parasite genome. (C) Real-time PCR trace for the mutant allele within Malawian isolates following normalization to 10,000 copies of EF1- per reaction. Comparison with Fig. 1A reveals the effective normalization of parasite DNA.

Validation of real-time PCR genotyping methodology. The assay was initially tested at parasite DNA copy numbers of 100, 250, 500, 1,000, and 10,000. At 10,000 copies, efficient discrimination of mutant and wild-type alleles was possible at concentrations between 5 and 100% (Fig. 3A and B). Insufficient amplification occurred at parasite DNA concentrations lower than this (data not shown). For the standard curve of mixed mutant (V1S) and wild-type (K1) DNAs (where wild-type and mutant DNAs were mixed so as to contain 0% to 100% mutant DNA), the C_T values obtained for the mutant probe (labeled with 6-carboxyfluorescein [FAM]) were then divided by those for the wild-type probe (labeled with VIC) and plotted against the log-transformed mutant DNA concentration (i.e., the percentage of total DNA that was mutant) (Fig. 3C).

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FIG. 3. Quantification of Pfdhfr I164L mutant alleles by real-time PCR. (A) Representative real-time PCR trace for the mutant probe conducted on mixtures of mutant and wild-type alleles. Parasite DNAs from a mutant strain (V1S) and a wild-type strain (K1) were quantified by normalization to EF1-. The DNA of the mutant allele was then diluted appropriately with that of the wild-type allele so as to yield wild-type DNA containing 0% to 100% mutant DNA. (B) Representative real-time PCR trace for the wild-type probe conducted on mixtures of mutant and wild-type alleles (reciprocal of panel A). (C) Representative standard curve used for quantification of the mutant allele in clinical isolates. The CT values obtained from panel A were divided by those obtained from panel B and plotted against the log-transformed percentage of mutant DNA. See the text for the limits of detection, limits of quantification, and inter- and intrarun accuracy and precision.

The limits of detection and of quantification were shown to be 5% and 10%, respectively. The intrarun accuracy and interrun accuracy were 98.4% and 106.7% at high mutant concentrations (75%) and 89.7% and 96.9% at low mutant concentrations (10%), respectively. The intrarun precision and interrun precision were 92.1% and 95.1% at high mutant concentrations (75%) and 92.2% and 96.8% at low mutant concentrations (10%).

Assessment of Pfdhfr I164L frequency in Malawian, Zambian, and Thai cohorts. In samples where the total-parasite DNA concentration was below 10,000 copies per µl, whole-genome PCR was conducted prior to genotypic analysis. Subsequently, sufficient DNA was obtained from 158 of the 210 Malawian isolates (94 pretreatment isolates and 64 isolates from recurrent parasitemia after treatment), 42 of 55 Zambian isolates, and 38 of 50 Thai isolates. Using the optimized real-time PCR methodology, the frequency of the I164L mutation in these isolates was monitored (Table 2). For the Thai isolates, the I164L mutation was present in 36 out of 38 samples tested. The two remaining samples showed indications that if the mutation was present, then it was below the 5% confidence level of the assay. Conversely, the I164L mutation was below the confidence level in all Malawian and Zambian isolates tested. Furthermore, there was no evidence of the selection of this I164L mutation in any of the 64 Malawian isolates appearing within 42 days of treatment with PYR-SDX (Table 3).

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TABLE 2. Frequency of the Pfdhfr I164L alleles in isolates from Thailand, Zambia, and Malawi

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TABLE 3. Frequency of different Pfdhfr I164L alleles in Malawian isolates from patients naïve to therapy and from patients having received therapy

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
We observed a large variability in the degree of amplification from field isolates. This was partly explained by variability in parasitemia, but the correlation coefficient was low (0.13), indicating that other factors contribute to this variability (e.g., DNA extraction efficiency). Furthermore, only partial selectivity of the mutant probe for the mutant sequence and of the wild-type probe for the wild-type sequence was observed. These discrepancies will result in high concentrations of wild-type sequences being incorrectly identified as partially mutant, which would cast doubt on the real-time PCR-based data reported by Alker et al. (1). However, one patient whose entire parasite population contained the mutant allele was reported, and this finding was confirmed by sequencing. Clearly, this cannot be explained by the lack of specificity of the real-time PCR assay. However, binding of minor groove binder probes prohibits the sequencing of the resultant amplicon, and the article does not provide an explanation for how this problem was circumvented.

Our approach was found to be both accurate and precise for the detection of the I164L mutation and was able to detect the mutation at a level of 5% of the total parasite population. Furthermore, the assay was able to accurately identify the presence of this mutation in Thai isolates at a frequency comparable to that observed in previous reports (5, 16).

The assay presented here allows improved sensitivity over conventional methodologies, but owing to the incomplete specificity of the probes, it is not possible to quantify the mutation if it is present at less than 5% of the total parasite population within an individual. However, of the Malawian isolates studied here, 64 were parasitological failures after treatment with PYR-SDX, and 57 of these had matched pretreatment isolates (Table 3). Antifolate therapy with PYR-SDX would be expected to select for the I164L mutation if it existed even at low levels in our patient population, yet it was not detected in any of the samples tested. The Malawian samples used in this study were collected from young children who self-presented to a health center situated 10 km away from the hospital where the samples for the Alker study were collected. It is unlikely that the parasites were under different drug pressures. This suggests that the I164L mutation was not preexistent within the samples tested in this study.

Our samples were collected between 2003 and 2005, whereas the samples in the previous study were collected between 2001 and 2003. Therefore, one would expect that in this time an expansion in the mutant population would have occurred under continued selection by high-level PYR-SDX as the first-line treatment in Malawi. Finally, the previous study utilized isolates from pregnant mothers who were HIV positive (1). It is possible that the immunosuppression associated with HIV infection or exposure to cotrimoxazole prophylaxis may have influenced the development of the Pfdhfr I164L mutation or that other, as yet unknown selection pressures were present in this patient group and not in children. It should be emphasized that each sample assayed in this paper could have had less than 5% of the mutant allele, and it would not have been detected. Nonetheless, the rarity of this allele in Africa despite more than a decade of use of PYR-SDX as frontline antimalarial therapy is an intriguing phenomenon, particularly given its rapid selection in Southeast Asia under similar circumstances (5, 16) and the fact that it is easily selected for in in vitro studies (21).

Using a fully validated methodology, we could not identify I164L mutants within clinical isolates from Malawi and Zambia, even in posttreatment failure parasites from Malawi. These data are reassuring, because even though CPG-DDS and CPG-DDS-artesunate have now been withdrawn, PYR-SDX is still used extensively for treatment in Africa and plays a major role in intermittent presumptive therapy programs in pregnancy. Our results are in agreement with the majority of previous reports, and coupled with the need for specialized equipment and the cost associated with real-time PCR, there appears to be no urgent need for field application of this method. The failure of antifolate chemotherapy to select I164L mutant parasites in Africa compared to Southeast Asia is an important phenomenon that clearly requires further investigation of the underlying mechanisms.

 
S.A.W. and A.O. designed the study. E.O., D.J.J., and A.O. analyzed the data. E.O., D.J.B., D.J.J., P.A.W., P.G.B., and S.A.W. contributed to the writing of the paper. U.D. provided the Zambian P. falciparum samples. D.J.B. conducted the trial in Malawi and provided the P. falciparum samples. U.D. and M.M. conducted the trial in Zambia, and S.M. conducted the trial in Thailand. E.O. extracted DNA from all samples used in the study and performed the real-time PCR.

This study was funded by support from Gates Malaria Programme Ph.D. studentships to E.O. S.A.W. and P.G.B. are supported by the MRC, BBSRC, and Wellcome Trust. The clinical trial in Malawi was funded by a Wellcome Trust Training Fellowship (066681) awarded to D.J.B. A.O. is supported by the United Kingdom MRC, European Commission, and United Kingdom National Institute of Health Research.

P.A.W. is the chairman, and S.A.W. is an unpaid member, of the MMV development team for the drug "CDA" (Dacart; GSK).

 
* Corresponding author. Mailing address: Department of Pharmacology and Therapeutics, University of Liverpool, 70 Pembroke Place, Liverpool, L69 3GF, United Kingdom. Phone: 44 (0) 151 794 5919. Fax: 44 (0) 151 794 5656. E-mail: aowen{at}liv.ac.uk

Published ahead of print on 25 August 2008.

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1
1. Alker, A. P., V. Mwapasa, A. Purfield, S. J. Rogerson, M. E. Molyneux, D. D. Kamwendo, E. Tadesse, E. Chaluluka, and S. R. Meshnick. 2005. Mutations associated with sulfadoxine-pyrimethamine and chlorproguanil resistance in Plasmodium falciparum isolates from Blantyre, Malawi. Antimicrob. Agents Chemother. 49:3919-3921.[Abstract/Free Full Text]
2
2. Anderson, T. J., S. Nair, C. Jacobzone, A. Zavai, and S. Balkan. 2003. Molecular assessment of drug resistance in Plasmodium falciparum from Bahr El Gazal province, Sudan. Trop. Med. Int. Health 8:1068-1073.[CrossRef][Medline]
3
3. Bates, S. J., P. A. Winstanley, W. M. Watkins, A. Alloueche, J. Bwika, T. C. Happi, P. G. Kremsner, J. G. Kublin, Z. Premji, and C. H. Sibley. 2004. Rare, highly pyrimethamine-resistant alleles of the Plasmodium falciparum dihydrofolate reductase gene from 5 African sites. J. Infect. Dis. 190:1783-1792.[CrossRef][Medline]
4
4. Bell, D. J., S. K. Nyirongo, M. Mukaka, E. E. Zijlstra, C. V. Plowe, M. E. Molyneux, S. A. Ward, and P. A. Winstanley. 2008. Sulfadoxine-pyrimethamine-based combinations for malaria: a randomised blinded trial to compare efficacy, safety and selection of resistance in Malawi. PLoS ONE 3:e1578.[CrossRef]
5
5. Biswas, S., A. Escalante, S. Chaiyaroj, P. Angkasekwinai, and A. A. Lal. 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 5:737-743.[CrossRef][Medline]
6
6. Dietmaier, W., A. Hartmann, S. Wallinger, E. Heinmoller, T. Kerner, E. Endl, K. W. Jauch, F. Hofstadter, and J. Ruschoff. 1999. Multiple mutation analyses in single tumor cells with improved whole genome amplification. Am. J. Pathol. 154:83-95.[Abstract/Free Full Text]
7
7. Djaman, J. A., A. Mazabraud, and L. Basco. 2007. Sulfadoxine-pyrimethamine susceptibilities and analysis of the dihydrofolate reductase and dihydropteroate synthase of Plasmodium falciparum isolates from Cote d'Ivoire. Ann. Trop. Med. Parasitol. 101:103-112.[CrossRef][Medline]
8
8. Hyde, J. E. 1990. The dihydrofolate reductase-thymidylate synthetase gene in the drug resistance of malaria parasites. Pharmacol. Ther. 48:45-59.[CrossRef][Medline]
9
9. Juliano, J. J., P. Trottman, V. Mwapasa, and S. R. Meshnick. 2008. Detection of the dihydrofolate reductase-164L mutation in Plasmodium falciparum infections from Malawi by heteroduplex tracking assay. Am. J. Trop. Med. Hyg. 78:892-894.[Abstract/Free Full Text]
10
10. Khim, N., C. Bouchier, M. T. Ekala, S. Incardona, P. Lim, E. Legrand, R. Jambou, S. Doung, O. M. Puijalon, and T. Fandeur. 2005. Countrywide survey shows very high prevalence of Plasmodium falciparum multilocus resistance genotypes in Cambodia. Antimicrob. Agents Chemother. 49:3147-3152.[Abstract/Free Full Text]
11
11. Krudsood, S., M. Imwong, P. Wilairatana, S. Pukrittayakamee, A. Nonprasert, G. Snounou, N. J. White, and S. Looareesuwan. 2005. Artesunate-dapsone-proguanil treatment of falciparum malaria: genotypic determinants of therapeutic response. Trans. R. Soc. Trop. Med. Hyg. 99:142-149.[CrossRef][Medline]
12
12. Masimirembwa, C. M., N. Phuong-Dung, B. Q. Phuc, L. Duc-Dao, N. D. Sy, O. Skold, and G. Swedberg. 1999. Molecular epidemiology of Plasmodium falciparum antifolate resistance in Vietnam: genotyping for resistance variants of dihydropteroate synthase and dihydrofolate reductase. Int. J. Antimicrob. Agents 12:203-211.[CrossRef][Medline]
13
13. Mberu, E. K., M. K. Mosobo, A. M. Nzila, G. O. Kokwaro, C. H. Sibley, and W. M. Watkins. 2000. The changing in vitro susceptibility pattern to pyrimethamine/sulfadoxine in Plasmodium falciparum field isolates from Kilifi, Kenya. Am. J. Trop. Med. Hyg. 62:396-401.[Abstract]
14
14. Mulenga, M., J. P. Van Geertruyden, L. Mwananyanda, V. Chalwe, F. Moerman, R. Chilengi, C. Van Overmeir, J. C. Dujardin, and U. D'Alessandro. 2006. Safety and efficacy of lumefantrine-artemether (Coartem) for the treatment of uncomplicated Plasmodium falciparum malaria in Zambian adults. Malar. J. 5:73.[CrossRef][Medline]
15
15. Mutabingwa, T., A. Nzila, E. Mberu, E. Nduati, P. Winstanley, E. Hills, and W. Watkins. 2001. Chlorproguanil-dapsone for treatment of drug-resistant falciparum malaria in Tanzania. Lancet 358:1218-1223.[CrossRef][Medline]
16
16. Nair, S., J. T. Williams, A. Brockman, L. Paiphun, M. Mayxay, P. N. Newton, J. P. Guthmann, F. M. Smithuis, T. T. Hien, N. J. White, F. Nosten, and T. J. Anderson. 2003. A selective sweep driven by pyrimethamine treatment in Southeast Asian malaria parasites. Mol. Biol. Evol. 20:1526-1536.[Abstract/Free Full Text]
17
17. Nzila, A., E. Ochong, E. Nduati, K. Gilbert, P. Winstanley, S. Ward, and K. Marsh. 2005. Why has the dihydrofolate reductase 164 mutation not consistently been found in Africa yet? Trans. R. Soc. Trop. Med. Hyg. 99:341-346.[CrossRef][Medline]
18
18. Nzila, A. M., E. K. Mberu, J. Sulo, H. Dayo, P. A. Winstanley, C. H. Sibley, and W. M. Watkins. 2000. Towards an understanding of the mechanism of pyrimethamine-sulfadoxine resistance in Plasmodium falciparum: genotyping of dihydrofolate reductase and dihydropteroate synthase of Kenyan parasites. Antimicrob. Agents Chemother. 44:991-996.[Abstract/Free Full Text]
19
19. Ochong, E., A. Nzila, S. Kimani, G. Kokwaro, T. Mutabingwa, W. Watkins, and K. Marsh. 2003. Molecular monitoring of the Leu-164 mutation of dihydrofolate reductase in a highly sulfadoxine/pyrimethamine-resistant area in Africa. Malar. J. 2:46.[CrossRef][Medline]
20
20. Owen, A., C. Goldring, P. Morgan, D. Chadwick, B. K. Park, and M. Pirmohamed. 2005. Relationship between the C3435T and G2677T(A) polymorphisms in the ABCB1 gene and P-glycoprotein expression in human liver. Br. J. Clin. Pharmacol. 59:365-370.[CrossRef][Medline]
21
21. Paget-McNicol, S., and A. Saul. 2001. Mutation rates in the dihydrofolate reductase gene of Plasmodium falciparum. Parasitology 122:497-505.[Medline]
22
22. Parzy, D., C. Doerig, B. Pradines, A. Rico, T. Fusai, and J. C. Doury. 1997. Proguanil resistance in Plasmodium falciparum African isolates: assessment by mutation-specific polymerase chain reaction and in vitro susceptibility testing. Am. J. Trop. Med. Hyg. 57:646-650.[Abstract/Free Full Text]
23
23. Plowe, C. V., J. F. Cortese, A. Djimde, O. C. Nwanyanwu, W. M. Watkins, P. A. Winstanley, J. G. Estrada-Franco, R. E. Mollinedo, J. C. Avila, J. L. Cespedes, D. Carter, and O. K. Doumbo. 1997. Mutations in Plasmodium falciparum dihydrofolate reductase and dihydropteroate synthase and epidemiologic patterns of pyrimethamine-sulfadoxine use and resistance. J. Infect. Dis. 176:1590-1596.[Medline]
24
24. Plowe, C. V., A. Djimde, M. Bouare, O. Doumbo, and T. E. Wellems. 1995. Pyrimethamine and proguanil resistance-conferring mutations in Plasmodium falciparum dihydrofolate reductase: polymerase chain reaction methods for surveillance in Africa. Am. J. Trop. Med. Hyg. 52:565-568.[Abstract/Free Full Text]
25
25. Plowe, C. V., J. G. Kublin, and O. K. Doumbo. 1998. P. falciparum dihydrofolate reductase and dihydropteroate synthase mutations: epidemiology and role in clinical resistance to antifolates. Drug Resist. Updat. 1:389-396.[CrossRef][Medline]
26
26. Schönfeld, M., I. Barreto Miranda, M. Schunk, I. Maduhu, L. Maboko, M. Hoelscher, N. Berens-Riha, A. Kitua, and T. Loscher. 2007. Molecular surveillance of drug-resistance associated mutations of Plasmodium falciparum in south-west Tanzania. Malar. J. 6:2.[CrossRef][Medline]
27
27. Tahar, R., and L. K. Basco. 2006. Molecular epidemiology of malaria in Cameroon. XXII. Geographic mapping and distribution of Plasmodium falciparum dihydrofolate reductase (dhfr) mutant alleles. Am. J. Trop. Med. Hyg. 75:396-401.[Abstract/Free Full Text]
28
28. Van Geertruyden, J. P., M. Mulenga, L. Mwananyanda, V. Chalwe, F. Moerman, R. Chilengi, W. Kasongo, C. Van Overmeir, J. C. Dujardin, R. Colebunders, L. Kestens, and U. D'Alessandro. 2006. HIV-1 immune suppression and antimalarial treatment outcome in Zambian adults with uncomplicated malaria. J. Infect. Dis. 194:917-925.[CrossRef][Medline]
29
29. Vasconcelos, K. F., C. V. Plowe, C. J. Fontes, D. Kyle, D. F. Wirth, L. H. Pereira da Silva, and M. G. Zalis. 2000. Mutations in Plasmodium falciparum dihydrofolate reductase and dihydropteroate synthase of isolates from the Amazon region of Brazil. Mem. Inst. Oswaldo Cruz 95:721-728.[Medline]
30
30. Wilairatana, P., D. E. Kyle, S. Looareesuwan, K. Chinwongprom, S. Amradee, N. J. White, and W. M. Watkins. 1997. Poor efficacy of antimalarial biguanide-dapsone combinations in the treatment of acute, uncomplicated, falciparum malaria in Thailand. Ann. Trop. Med. Parasitol. 91:125-132.[CrossRef][Medline]

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Antimicrobial Agents and Chemotherapy, November 2008, p. 3883-3888, Vol. 52, No. 11
0066-4804/08/$08.00+0     doi:10.1128/AAC.00431-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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• Alker, A. P., Juliano, J. J., Meshnick, S. R., Owen, A., Ochong, E., Bell, D. J., Johnson, D. J., d'Alessandro, U., Mulenga, M., Muangnoicharoen, S., Van Geertruyden, J. P., Winstanley, P. A., Bray, P. G., Ward, S. A. (2009). Plasmodium falciparum and Dihydrofolate Reductase I164L Mutations in Africa. Antimicrob. Agents Chemother. 53: 1722-1723 [Full Text]  

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