Polymorphisms in the dihydrofolate reductase (DHFR) and ...

Tropical Medicine and International Health volume 3 no 8 pp 605–609 august 1998

Polymorphisms in the dihydrofolate reductase (DHFR) and dihydropteroate synthetase (DHPS) genes of Plasmodium falciparum and in vivo resistance to sulphadoxine/pyrimethamine in isolates from Tanzania T. Jelinek1,2,A. M. Rønn3, M. M. Lemnge4, J. Curtis1, J. Mhina4, M.T. Duraisingh1, I. C. Bygbjerg3 and D. C.Warhurst1 1 London School of Hygiene and Tropical Medicine, Department of Medical Parasitology, London, UK 2 Department of Infectious Diseases and Tropical Medicine, University of Munich, Germany 3 Centre of Medical Parasitology, Unit of Tropical Medicine, Department of Infectious Diseases, State University Hospital, Rigshospitalet, Copenhagen, Denmark 4 National Institute for Medical Research, Amani Research Centre, Amani, Tanzania

Summary

The efficacy of sulphadoxine/pyrimethamine (S/P) in treatment of uncomplicated falciparum malaria in Africa is increasingly compromised by development of resistance. The occurrence of mutations associated with the active site sequence in the Plasmodium falciparum genes coding for dihydrofolate reductase (DHFR) and dihydropteroate synthetase (DHPS) is associated with in vitro resistance to pyrimethamine and sulphadoxine. This study investigates the occurrence of these mutations in infected blood samples taken from Tanzanian children before treatment with S/P and their relationship to parasite breakthrough by day 7. The results show that alleles of DHPS (436-alanine, 437-alanine and 540-lysine) were significantly reduced in prevalence on day 7 after S/P treatment. In this area, a DHPS with 436-serine, 437-glycine and 540-glutamate appears to play a major role in resistance to S/P in vivo. Evidence for the influence of mutations in the DHFR gene in this investigation is not clear, probably because of the high prevalence of ‘resistance-related’ mutations at day 0 in the local parasite population. For apparently the same reason, it was not possible to show a statistical association between S/P resistance and the presence of particular polymorphisms in the DHFR and DHPS genes before treatment.

keywords malaria, drug resistance, Plasmodium falciparum, Tanzania correspondence Dr Tomas Jelinek University of Munich, Department of Infectious Diseases and Tropical Medicine, Leopoldstr. 5, 80802 Munich, Germany. Fax: 149 89 33 61 12; E-mail: [email protected]

Introduction The spread of chloroquine-resistant Plasmodium falciparum malaria has led to an increasing use of the antifolate combination of sulphadoxine/pyrimethamine (S/P) in Africa. Clinical resistance to S/P has arisen rapidly in other areas of the world, after it replaced chloroquine. Monitoring and, if possible, delaying the spread of S/P-resistance is therefore a major public health objective in affected African countries. All efforts to control resistance will require methods that make it possible to map S/P-resistant malaria quickly and accurately in epidemiological studies. Currently used in vitro and in vivo methods for the measurement of drug resistance in P. falciparum are not well-suited for fast epidemiological

© 1998 Blackwell Science Ltd

surveillance. A technique is needed that allows the screening of a large number of samples without unduly straining scarce financial resources. Drug resistance of Plasmodium falciparum to pyrimethamine has been associated with point mutations in the dihydrofolate reductase (DHFR) gene. This gene encodes the enzyme dihydrofolate reductase, which is selectively inhibited by pyrimethamine and cycloguanil (the active form of proguanil). Serine in position 108 of DHFR is found in strains sensitive to both drugs, whereas a mutation to asparagine at that position is associated with resistance to pyrimethamine and a moderate loss of response to cycloguanil. A mutation to threonine at position 108, together with an alanine to valine change at position 16, 605

Tropical Medicine and International Health

volume 3 no 8 pp 605–609 august 1998

T. Jelinek et al. Polymorphisms in P. falciparum genes and in vivo resistance

appears to confer resistance to cycloguanil with only slight loss of response to pyrimethamine. A modulation towards higher levels of pyrimethamine resistance appears to be facilitated by mutations of asparagine to isoleucine at position 51 and of cysteine to arginine at position 59 when associated with the asparagine-108 mutation. A mutation from isoleucine to leucine at position 164 in combination with the asparagine-108 and one or both of the isoleucine-51 or arginine-59 mutations has been found in P. falciparum strains that are highly resistant to both drugs (Foote et al. 1990; Peterson et al. 1990). Polymorphisms at 5 highly conserved positions within the dihydropteroate synthetase (DHPS) gene have been reported in sulphadoxine-resistant isolates of P. falciparum: codons 436, 437, 581, 613 (Brooks et al. 1994; Triglia & Cowman 1994), and also 540 (Wang et al. 1997). However, unlike DHFR, no single polymorphism has been associated with all resistant strains and only limited data are available on the global distribution of these polymorphisms (Wang et al. 1995). The occurrence of resistance of P. falciparum to pyrimethamine was first reported from north-east Tanzania in 1954 (Clyde 1954). In 1957, resistance to pyrimethamine was observed after 5 months of prophylactic use of weekly doses (Clyde & Shute 1957). The combination of sulphadoxine-pyrimethamine has been effective in this area, despite occasional reports of resistance in expatriates (Timmermanns et al. 1982; Vleugels et al. 1982) and an increasing number of RI resistance in expatriates during the 1980s (Shapira et al. 1986). In 1994, however, a therapeutic study among children in Tanga region, some of whom had received prophylactic doses of pyrimethamine-dapsone for one year, showed that 73.7% failed to clear the parasites from their blood within 7 days (Rønn et al. 1996). In order to monitor the development of resistance against S/P and to assess possible associations between in vivo test results and point mutations of the antifolate target genes, a second and third therapeutic study were undertaken in 1995 and 1996 in the same area and within the same population as in 1994.

Patients, materials and methods In vivo testing Forty-five children in 1995 and 27 in 1996 aged 1–9 years were recruited for the studies in the village of Magoda, Tanga Region, Tanzania. Inclusion criteria were: monoinfection with P. falciparum, an asexual parasitaemia of $1000/ml blood, no intake of antimalarials during the 2 weeks prior to presentation, no intake of sulphadoxine-pyrimethamine during the previous 2 weeks, no complicated malaria and informed consent of the parents. Most of the children participating in the survey did not suffer from clinical signs of 606

malaria. All enrolled children were treated with a single dose of sulphadoxine-pyrimethamine tablets (Fansidar™), following the dosage advice of the manufacturer (Rønn et al. 1996). Blood was drawn from all children on days 0, 1, 2, 3, and 7 and examined by microscopists for malaria parasites using Giemsa stained thin and thick blood films. The number of parasites per ml in P. falciparum infections was assessed on thick films according to published recommendations by counting the number of asexual stages of P. falciparum per 200 leucocytes and multiplying the result by 40, assuming a mean leucocyte count of 8 3 109/l (Anonymous 1991). The heparinized blood that was drawn at day 0 and thick blood films taken at day 7 were stored appropriately and transported to the London School of Hygiene and Tropical Medicine for further examination. Preparation and amplification of DNA DNA was purified from whole blood by phenol-chloroformextraction. Following a modified procedure used for DNA extraction from material derived from Egyptian mummies (Pääbo 1990), blood slides taken at day 7 were incubated in Tris-HCl, EDTA, SDS, Dithiothreitol and Proteinase K for 16 h. Subsequently, the dissolved material was submitted to a phenol-chloroform-extraction. A nested PCR method was used in all samples. Two ml of purified DNA were added to a final reaction volume of 50 ml. In the first round, a 648basepair (bp) portion of the DHFR gene was amplified by use of the primers M1 (59-TTT-ATG-ATG-GAA-CAA-GTCTGC-39) and M5 (59-AGT-ATA-TAC-ATC-GCT-AAC-AGA39). Similarly, a 710 bp portion of the DHPS gene was amplified by use of R2 (59-AAC-CTA-AAC-GTC-CTG-TTCAA-39) and R/(59-AAT-TGT-GTG-ATT-TGT-CCA-CAA-39). For the second round of the DHFR gene, 2 ml of amplified DNA from the M1-M5 primer pair were added to each of 2 PCR mixtures: F (59-GAA-ATG-TAA-TTC-CCT-AGA-TATGgA-ATA-TT-39) and M4 (59-TTA-ATT-TCC-CAA-GTAAAA-CTA-TTA-GAg-CTT-C-39) to detect 59-arginine, 108serine, and 108-threonine, or M3 (59-TTT-ATG-ATG-GAACAA-GaC-TGg-GAC-GTT-39) and F/(59-AAA-TTC-TTGATA-AAC-AAC-GGA-ACC-Ttt-TA-39) to amplify the fragments containing 16-alanine, 51-asparagine, 108asparagine, and 164-leucine. Similarly, for the DHPS gene, DNA amplified with the R2-R/primer pair was added to each of 2 PCR mixtures: K (59-TGC-TAG-TGT-TAT-AGA-TATAGC-atG-AGc-ATC-39) and K/(59-CTA-TAA-CGA-GGTATT-gCA-TTT-AAT-gCA-AGA-A-39) to detect 436-serine, 436-alanine, 437-glycine, 437-alanine, 540-glutamic acid, or L (59-ATA-GGA-TAC-TAT-TTG-ATA-TTG-GAc-cAG-GGATTc-G-39) and L/(59-TAT-TAC-AAC-ATT-TTG-ATC-ATTCgc-GCA-Acc-GG-39) for 581-alanine, 581-glycine, 613threonine, 613-serine and 613-alanine. The primer pair K/and





J (59-TGC-TAG-TGT-TAT-AGA-TAT-AGG-TGG-AGA-AagC-39) was used to detect 436-phenylalanine. Positive and negative controls were included in all amplification procedures. Restriction fragment length polymorphisms (RFLP) Depending on the quality of the bands in the control gel, 2–5 ml of PCR product were incubated with restriction enzymes according to the manufacturers instructions (New England Biolabs, Beverly, MA, USA). The 326 bp product of the primer pair F-M4 was cut by AluI at codon 108 (serine). In the same PCR product, BstNI detected the 108-threonine mutation and XmnI detected 59-arginine. The amplified product of the M3-F/primer pair (522 bp) was cut by DraI to detect 164-leucine and by BsrI to detect 108-asparagine. A restriction site for NlaIII in this PCR product was destroyed by the mutation from 16-alanine to 16-valine and the restriction site for Tsp509I at 51-asparagine was destroyed by the mutation to 51-isoleucine. For the DHPS gene, the 438 bp product of K-K/was cut by MnlI to identify 436serine, MSPAI to detect 436-alanine, AvaII for 437-glycine, MwoI for 437-alanine, and FokI for 540-glutamic acid. The 161 bp product of L-L/was cut by BstUI to detect 581-alanine, Bsl1 for 581-glycine, MwoI for 613-alanine, and AgeI for 613-threonine or BsaWI for 613-threonine and serine. All digested products were separated by electrophoresis in an 1% SeaKem™ plus 1% NuSieve™ (both FMC BioProducts, Rockland, ME, USA) gel. DNA from established laboratory strains of P. falciparum served as controls of PCR and enzyme digests. In particular, the strains K1 (Thailand), FC27 (Papua New Guinea), FCR3 (The Gambia), W2 (Indochina), V1/S (Vietnam) and 7G8 (Brazil) were used.

Results In 1995, 21 (46.6%) of the 45 recruited children still had microscopically detectable parasitaemia 7 days after treatment with sulphadoxine/pyrimethamine (S/P). The median parasitaemia at day 7 in patients harbouring resistant strains was 80/ml (range: 40–7,720/ml), as opposed to 2,320/ml at day 0 (range: 1000–50,240/ml). In contrast, 26 (96.3%) of the 27 children recruited in 1996 had microscopically detectable parasitaemia at day 7 after treatment (P , 0.00006). In this group, the median parasitaemia was 5,880/ml (range: 1,800–62,400/ml) at day 0, and 640/ml (range: 40–66,240/ml) at day 7. P. falciparum isolates from all 45 children who were recruited in 1995 (DHFR results have already been published: Jelinek et al. 1997) showed polymorphisms in at least one codon of the DHFR or DHPS genes. Variations (more than


one allele detected) were restricted to 51asparagine/isoleucine, 59-cysteine/arginine, and 108serine/asparagine on the DHFR gene and to 436alanine/serine, 437-alanine/glycine, 540-lysine/glutamic acid on the DHPS gene (Table 1). No polymorphisms were detected at codons 16-alanine and 164-isoleucine of DHFR nor 581- and 613-alanine of DHPS. No significant association between polymorphisms on either the DHFR or the DHPS gene (or a combination of these) and clinical S/P resistance could be detected in the 1995 isolates (Table 1). The DNA from thick blood smears from day 7 after S/P treatment in 1996 were examined for genetic polymorphisms along with the blood samples from day 0. Twenty-six of 27 infections in 1996 were classified as resistant. Compared to all isolates from 1995, the 1996 increase of in vivo resistance against S/P went along with a disappearance of the variant 51-asparagine which had been present in 31.3% of the 1995 isolates (P 5 0.003; Yates) (Table 2). Eighteen of the 27 isolates collected in 1996 showed variation at day 0. However, at day 7 no mixed allelic infections were seen (see Table 2). Variations were apparent at 59-arginine/cysteine of DHFR, and in DHPS at 437-alanine/glycine and 540-lysine/glutamic acid. Comparison of the 1996 isolates collected at day 0 with those collected at day 7 showed a significant decrease of the alleles 436-alanine (P 5 0.05), 437-alanine (P 5 0.0005), and 540-lysine (P 5 0.001) of DHPS. Table 1 Polymorphisms in codons of the DHFR- and DHPS-genes and their association to in vivo resistance in patients from north-east Tanzania during 1995

Gene

Codon

Resistant (%) Sensitive (%) All (%) (n 5 21) (n 5 24) (n 5 45)

DHFR 16-alanine 51-asparagine 51-isoleucine 59-arginine 59-cysteine 108-serine 108-asparagine 108-threonine 164-isoleucine

21 (100%) 07 (33.3%) 14 (66.7%) 19 (90.5%) 13 (61.9%) 04 (19%) 21 (100%) 00 21 (100%)

24 (100%) 07 (29.2%) 20 (83.3%) 22 (91.7%) 12 (50%) 04 (16.7%) 23 (95.8%) 00 24 (100%)

45 (100%) 14 (31.1%) 34 (75.6%) 41 (91.1%) 25 (55.6%) 08 (17.8%) 44 (97.8%) 00 45 (100%)

DHPS

21 (100%) 02 (9.5%) 00 12 (57.1%) 18 (85.7%) 11 (52.4%) 17 (81%) 21 (100%) 00 21 (100%)

24 (100%) 07 (29.2%) 00 14 (58.3%) 19 (79.2%) 14 (58.3%) 19 (79.2%) 24 (100%) 00 24 (100%)

45 (100%) 09 (20%) 00 26 (57.8%) 37 (82.2%) 25 (55.6%) 36 (80%) 45 (100%) 00 45 (100%)

436-serine 436-alanine 436-phenylalanine 437-alanine 437-glycine 540-lysine 540-glutamic acid 581-alanine 581-glycine 613-alanine

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Table 2 Polymorphisms in codons of the DHFR- and DHPS-genes: comparison between samples collected in 1995 and 1996 at day 0 and follow-up samples from day 7 collected in 1996

Gene

Codon

1995 day 0 (n 5 45)

1996 day 0 (n 5 27)

1996 day 7 (n 5 26)

DHFR

16-alanine 51-asparagine 51-isoleucine 59-arginine 59-cysteine 108-serine 108-asparagine 108-threonine 164-isoleucine

45 (100%) 14 (31.3%) 34 (75.6%) 41 (91.1%) 25 (55.6%) 08 (17.8%) 44 (97.8%) 00 45 (100%)

27 (100%) 00 27 (100%) 25 (92.9%) 09 (33.3%) 01 (3.7%) 27 (100%) 00 27 (100%)

26 (100%) 00 26 (100%) 21 (80.8%) 05 (19.2%) 00 26 (100%) 00 26 (100%)

DHPS

436-serine 436-alanine 436-phenylalanine 437-alanine 437-glycine 540-lysine 540-glutamic acid 581-alanine 581-glycine 613-alanine

45 (100%) 09 (20%) 00 26 (57.8%) 37 (82.2%) 25 (55.6%) 36 (80%) 45 (100%) 00 45 (100%)

27 (100%) 05 (18.5%) 00 18 (66.7%) 23 (85.2%) 17 (63%) 23 (85.2%) 27 (100%) 00 27 (100%)

26 (100%) 00 00 04 (15.4%) 22 (84.6%) 04 (15.4%) 22 (84.6%) 26 (100%) 00 26 (100%)

Discussion The combination of sulphadoxine/pyrimethamine (S/P) is widely used in Africa and has been promoted by WHO as an alternative to chloroquine for the rapid treatment of uncomplicated malaria. Under the pressure of changing drug resistance patterns in P. falciparum strains and diminishing public health resources in Africa, the development of fast, reliable, and affordable methods for the determination of drug resistance is paramount. A technique that is able to determine developing, but not yet clinically apparent drug resistance would be desirable. Therefore various studies concerning the determination of genetic changes as the basis of clinical resistance have been conducted. Polymorphisms in various codons of the DHFR and DHPS genes have been associated with in vitro resistance to antifolate drug combinations in various laboratory clones and occasional field isolates (Foote et al. 1990; Peterson et al. 1990; Brooks et al. 1994; Triglia & Cowman 1994; Basco et al. 1995; Reeder et al. 1996). While resistance to pyrimethamine alone is shown by parasites carrying only the 108-asparagine mutation on DHFR, S/P resistance could occur in two ways. First, further mutation in the DHFR gene might confer sufficient pyrimethamine resistance to render the synergistic S/P combination ineffective, or second, and more likely, the 608

emergence of sufficient resistance to sulphadoxine might yield the same result. Analysis of both genes is therefore necessary. The two field studies presented here were conducted in 1995 and 1996 with participation of 72 children from north-east Tanzania, who presented with asymptomatic P. falciparum parasitaemia. In 1995, 21 (46.6%) of 45 children were still parasitaemic at day 7 after treatment with sulphadoxine/pyrimethamine. In 1996, the rate of resistant samples had risen to 96.3%. The reasons for this increase of in vivo resistance in the studied population remains unclear. A variety of S/P preparations are on sale in the markets of the Tanga Region, but, to our knowledge, patterns of distribution did not change from 1995 to 1996. A comparison of the polymorphisms present at day 0 in 1995 in a population with a resistance rate of 46.6% and the polymorphisms that were detected in 1996 in a situation with 96.3% resistance to S/P revealed a significant change in one codon (Table 2): although present in 31.3% of the isolates in 1995, the allele 51-asparagine was not detected in 1996. Selection towards a certain genotype under S/P pressure became obvious when results from isolates from day 7 were compared with those obtained before treatment (Table 2). A significant selection against the alleles 436-alanine, 437alanine and 540-lysine on DHPS was found in 1996. Resistance against combinations of antifolates probably requires at least one resistant-related polymorphism in both the DHPS and the DHFR genes. Isolated polymorphisms in one of these genes will not necessarily lead to resistance to a drug combination. The findings of this study, taking into account changes in consecutive years and the influence of treatment on DHFR and DHPS polymorphisms, suggest that alleles 51-isoleucine on DHFR and 436-serine, 437-glycine and 540-glutamic acid on DHPS play a major role in the development of in vivo resistance in P. falciparum strains against S/P (Table 2). This confirms previous observations on laboratory strains (Foote et al. 1990; Peterson et al. 1990; Brooks et al. 1994; Triglia & Cowman 1994). As an epidemiological approach, the determination of polymorphisms of the DHFR gene has been used previously to estimate the amount of parasite resistance against pyrimethamine in West Africa and this method has been proposed for surveillance purposes (Plowe et al. 1995). This approach is valid provided the polymorphisms examined are confirmed to be the cause of the resistance in vivo. For example, Curtis et al. 1996 demonstrated selection for 108asparagine of DHFR after treatment with S/P or chlorproguanil/dapsone in Tanzanian children with asymptomatic infections. In the present study, no statistically significant association between polymorphisms observed at day 0 on either or both genes and in vivo S/p resistance was detected in 1995 or 1996. This is probably because of the high





prevalence of ‘resistance-related’ mutations before treatment in the local parasite population. These observations indicate that further studies on in vivo resistance and gene polymorphisms are advisable in different geographical areas, before screening for drug resistancerelated mutations as an epidemiological and drug-policy planning tool can be recommended. Such studies should be supplemented with in vivo studies on how parasites as well as clinical symptoms are cleared (Anonymous 1996).

Acknowledgements This work was supported financially by DANIDA, Wellcome Trust, Medical Research Council, Friedrich Baur Trust, and PHLS. We thank all villagers and village health workers for their assistance. Technical assistance by Mr. J. Raphael and Mrs. M. Muniss during the in vivo studies is greatly appreciated. Permission to publish was granted by Professor W. L. Kilamo, Director General, National Institute for Medical Research, Tanzania.

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