https://orcid.org/0009-0005-4127-1709
Figures
Abstract
Background
Plasmodium vivax has become the predominant species in the border regions of Thailand. The emergence and spread of antimalarial drug resistance in P. vivax is one of the significant challenges for malaria control. Continuous surveillance of drug resistance is therefore necessary for monitoring the development of drug resistance in the region. This study aims to investigate the prevalence of the mutation in the P. vivax multidrug resistant 1 (Pvmdr1), dihydrofolate reductase (Pvdhfr), and dihydropteroate synthetase (Pvdhps) genes conferred resistance to chloroquine (CQ), pyrimethamine (P) and sulfadoxine (S), respectively.
Method
100 P. vivax isolates were obtained between January to May 2023 from a Kanchanaburi province, western Thailand. Nucleotide sequences of Pvmdr1, Pvdhfr, and Pvdhps genes were amplified and sequenced. The frequency of single nucleotide polymorphisms (SNPs)-haplotypes of drug-resistant alleles was assessed. The linkage disequilibrium (LD) tests were also analyzed.
Results
In Pvmdr1, T958M, Y976F, and F1076L, mutations were detected in 100%, 21%, and 23% of the isolates, respectively. In Pvdhfr, the quadruple mutant allele (I57R58M61T117) prevailed in 84% of the samples, followed by (L57R58M61T117) in 11%. For Pvdhps, the double mutant allele (G383G553) was detected (48%), followed by the triple mutant allele (G383M512G553) (47%) of the isolates. The most prevalent combination of Pvdhfr (I57R58M61T117) and Pvdhps (G383G553) alleles was sextuple mutated haplotypes (48%). For LD analysis, the association in the SNPs pairs was found between the intragenic and intergenic regions of the Pvdhfr and Pvdhps genes.
Conclusion
The study has recently updated the high prevalence of three gene mutations associated with CQ and SP resistance. Genetic monitoring is therefore important to intensify in the regions to further assess the spread of drug resistant. Our data also provide evidence on the distribution of drug resistance for the early warning system, thereby threatening P. vivax malaria treatment policy decisions at the national level.
Citation: Sridapan T, Rattanakoch P, Kijprasong K, Srisutham S (2024) Drug resistance markers in Plasmodium vivax isolates from a Kanchanaburi province, Thailand between January to May 2023. PLoS ONE 19(7): e0304337. https://doi.org/10.1371/journal.pone.0304337
Editor: Kristan Alexander Schneider, University of Applied Sciences Hochschule Mittweida, GERMANY
Received: August 27, 2023; Accepted: May 10, 2024; Published: July 5, 2024
Copyright: © 2024 Sridapan et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This research is supported by Ratchadapisek Somphot Fund for Postdoctoral Fellowship, Chulalongkorn University, Thailand Science research and Innovation Fund Chulalongkorn University (HEA663700102), Research funds of Faculty of Allied Health Sciences, Chulalongkorn University. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Malaria remains a significant cause of parasitic public health problems in many tropical and sub-tropical countries. In 2021, 247 million cases and 619,000 malaria deaths occurred, according to a recent report by the World Health Organization (WHO) [1]. Malaria cases and deaths in Thailand continuously decreased by 90% from 2000 to 2019 [2, 3]. However, persistent transmission has remained a major public health challenge and ongoing importation, particularly in vulnerable forests and forest fringes along the international borders of Thailand [2, 3]. Plasmodium vivax (92.19%) is the most widespread species, accounting for most diagnosed human malaria infections in Thailand after Plasmodium falciparum (4.87%) [4]. Even though vivax infections are rarely mortal, it can cause severe illness in humans [5, 6].
The Thai Government has endorsed plans to achieve malaria elimination by 2024 [7]. However, widespread resistance to antimalarial drugs still poses serious challenges to accomplishing this goal [8]. The prevalence of Plasmodium parasite resistance toward antimalarial drugs remains high in border areas due to population migration across the borders. Three-day chloroquine (CQ) co‐administered with fourteen-day primaquine (PQ) remains the first-line treatment for all P. vivax cases for several decades in Thailand, while piperaquine (PPQ) or artemisinin combination therapies (ACTs) plus PQ were the second line treatment in case of CQ resistance (CQR) [9, 10]. There are increasing reports of treatment failure after the administration of CQ in different regions worldwide [11]. In Thailand, most P. vivax isolates are still sensitive and effectively respond well to the CQ standard regimen [12] even though there was a clinical report of CQR to P. vivax in a Thai pregnant woman [13]. This evidence indicates that CQR was already present in Thailand. Meanwhile, sulfadoxine‐pyrimethamine (SP), which is a combination anti-malarial drug was used in mixed infection cases or at the areas with suspected CQR in falciparum malaria in Thailand during 1972–1982. SP was also one of the partner drugs for the treatment of uncomplicated falciparum malaria and intermittent presumptive treatment (IPT) for infants, children, and pregnant women [14, 15]. In Thailand, SP has never been used as a first‐line treatment for vivax malaria [10]. During intensive SP use for the treatment of P. falciparum through coinfection with P. vivax in endemic areas [16, 17], together with the practice of malaria treatment without microscopic confirmation, this evidence leads to P. vivax being exposed to SP selective drug pressure inadvertently, resulting in the emergence of drug resistance to SP in P. vivax populations [18]. Resistance of the P. vivax to SP was highly reported along the Thai–Myanmar and Thai–Cambodia borders [19–21]. Self-treatment of fever or proper use of SP, including co-infection with P. vivax and P. falciparum in those endemic areas, leads to highly continue SP selection pressure onto P. vivax [16, 21].
Several molecular markers that show an association with drug resistance in P. vivax isolates have been identified [22]. P. vivax multidrug resistance (Pvmdr1), homologous to P. falciparum orthologs (Pfmdr1), has been reported to be correlated with CQ resistance [23, 24]. Pvmdr1 gene encodes the digestive vacuole (DV) transmembrane proteins, which function as a CQ transporter [25]. When Pvmdr1 was mutated, CQ diffusion into the DV of the parasite was reduced, conferring CQR [22]. However, the role of Pvmdr1 in CQR remains unclear [26]. The Y976F mutation in Pvmdr1 is associated with decreased in vitro sensitivity to CQ [23, 27]. A point mutation in Y976F and F1076L has been reported as an association with the clinical response of vivax malaria to CQR [13]. These two mutations were also used as molecular markers for monitoring the occurrence and spread of P. vivax CQR from many malaria-endemic regions [20, 28–30]. Meanwhile, dihydropteroate synthase (DHPS) and dihydrofolate reductase (DHFR) are enzymes in the folate biosynthesis pathway of the malarial parasite that are drug targeted of S and P, respectively [31]. SP resistance in P. vivax is caused by point mutations in antifolate resistance genes Pvdhps and Pvdhfr [22]. Mutation in these two genes causes alterations of amino acid residues in the enzyme-binding pockets leading to reduce the affinity of the enzyme for the drug [32]. Molecular studies have identified specific point mutations in the Pvdhfr gene in codons F57I/L, S58R, T61M, and S117T/N, while three mutations in codons S382A/C, A383G, and A553G/C have been detected in the Pvdhps gene [16, 24, 33, 34]. Point mutations of these genes are widespread in P. vivax populations worldwide [19–21, 35–39], resulting in an altered clinical response to SP [40]. The presence of mutations associated with CQ and SP resistance in P. vivax was previously reported in the Thai–Myanmar border regions [19–21, 29]. In response to this situation, continuous monitoring of the emergence and spread of antimalarial drug resistance is required to investigate the current status of the CQ and SP drug resistance in P. vivax cases in this region. The data provided is important for Thailand’s existing national malaria control strategy.
The aim of this study was to investigate the prevalence of genes potentially linked to drug resistance, including Pvmdr1, Pvdhfr, and Pvdhps, in P. vivax parasites isolated from Kanchanaburi, a province in western Thailand bordering Myanmar between January and May 2023.
Materials and methods
Study site
The study was conducted at the Satan Prabaramee Hospital, Naung Proe, Kanchanaburi province, Thailand, during January and May 2023. Kanchanaburi province (14° 1’-10" N; 99° 31’-52" E) is located in the west of Thailand. The province is mountainous, and its climate is defined as tropical, with a monsoon season from May to October and a dry season from November to April. The temperature typically varies from 20 to 37°C. This region was selected as the study site because it was a malarial endemic area where highly mobile populations can contribute to cross-border transmission along the Thai-Myanmar border frequently [41]. Kanchanaburi is one of the top three provinces for malaria cases in Thailand, according to reports from the Department of Disease Control, Ministry of Public Health, Thailand [42]. The inclusion criterion was adults (aged 20 years and over) presenting malaria infections, whereas the exclusion criterion was non-consent.
Ethical considerations
The study protocol was reviewed and approved by the Institutional Review Board for human research of Chulalongkorn University (Permit number: COA 010/66), Bangkok, Thailand. All adult participants were given a detailed explanation of study protocols and procedures. All methods were carried out in accordance with the Declaration of Helsinki and regulations prescribed by the above organizations. Written or thumbprint informed consent was obtained for the use of remnant blood samples from study participants.
Samples
The sample size was calculated based on a previous prevalence of P. vivax infection in western Thailand of 3.8% [43] at a confidence interval (CI) of 99% and a margin of error at 5%, using a single population proportion formula: n = [(Zα/2)2p(1-p)] / d2 = [(2.576)20.038(1–0.038)] / 0.052 = 98, where n = sample size, (Zα/2)2 = a confidence interval (CI) of 99%, p = prevalence of malaria and d = margin of error (5%). This gives a sample size of 98. Thus, a total of 100 participants were recruited for this study. The samples were obtained from 100 confirmed P. vivax-infected patients from January to May 2023. All patients presenting with malaria-related signs were diagnosed and reported by the Satan Prabaramee Hospital in a Kanchanaburi province. After venous blood collection was performed, thin blood films were prepared at the time of sample, stained with Giemsa’s staining and examined under the light microscope by two independents experienced microscopists in the identification of malaria parasites. A blood smear was interpreted as negative results if no parasites were observed under at least 100 high-power fields. The parasitemia was calculated as the formula: [(counted parasites / counted leucocytes) × the assumed number of leucocytes per microliter of blood (8 × 103) [44].
DNA extraction
Genomic DNA was extracted from 200 μl of the remainder of the whole blood samples using QIAamp DNA Blood Kits (Qiagen, Germany) according to the manufacturer’s instructions. The DNA elution volume was 200 μl in elution buffer, as provided by kits. The extracted DNA was stored at -20°C until used.
Nested PCR
A nested PCR previously described [45–47] for amplifying the 18S ribosomal RNA (rRNA) gene of Plasmodium spp. was used to confirm the P. vivax parasite, with some modifications: (i) using 1.5 mM of MgCl2, and (ii) changing the annealing step to 30 cycles at 64°C. The primer sequences and reaction conditions were performed in the Table 1 [45]. The reactions were carried out in a Mastercycler EP Gradient S (Eppendorf, Germany) in a total volume of 20 μl containing 1× PCR buffer, 1.5 mM MgCl2, 1 U Taq DNA polymerase (Vivantis, Malaysia), 0.2 mM dNTPs (Biotechrabbit, Germany), 0.25 μM each of forward and reward primers (Bio Basic Inc., Canada), and 2 μl of DNA template. One microliter of the nest-1 products was used as a template for the second round of PCR (nest-2) using the P. vivax species-specific primers (Table 1) [47]. Nest-2 PCR products were electrophoresed separately on a 1.5% agarose gel (SERVA Electrophoresis GmbH, Germany), stained with SERVA DNA Stain G (SERVA Electrophoresis GmbH, Germany), and visualized under UV light using a UV transilluminator (Bio-Rad, USA) to visualize and size the bands. Reaction with no DNA template having sterile distilled water was used as blank control.
PCR amplification and sequencing
Partial regions flanking major mutations of drug resistance marker genes, including Pvmdr1, Pvdhfr, and Pvdhps were amplified by nested PCR using primer sets from previous studies (Table 1) [48–50]. The reactions were performed in a total volume of 20 μl containing 1× PCR buffer, 1.5 mM MgCl2, 1 U Taq DNA polymerase (Vivantis, Malaysia), 0.2 mM dNTPs (Biotechrabbit, Germany), 0.2 μM each of forward and reward primers (Bio Basic Inc., Canada), and 2 μl of DNA template. The PCR conditions were carried out in Table 1. The nest-2 PCR products were analyzed, as described above. The PCR products were then purified using kits (Bio-Helix, Taiwan) and sent for Sanger DNA sequencing to Macrogen using the ABI 3730 XL DNA Analyzer (South Korea).
Sequence polymorphism and statistical analyses
Sanger sequences generated in this study (n = 100) were analyzed using the free software Bioedit version 7.2.5. Low-quality scores from sequencing errors on nearby upstream bases were cropped.
A lot of noise or background along the bottom of the trace was re-analyzed. All overlapping peaks in electropherograms (heterozygous) were also considered. Sanger sequences whose electropherograms showed single peaks were involved in the analysis. The sequenced fragments were validated by BLASTN searches (https://blast.ncbi.nlm.nih.gov/Blast.cgi). Consensus sequences of all three drug resistance marker genes were then aligned using the clustalW algorithm, as viewed in Bioedit software to identify polymorphisms compared with the following reference sequences of P. vivax; Pvmdr1 (accession no. XM001613678), Pvdhfr (accession no. XM001615032), and Pvdhps (accession no. XM001617159). Each antimalarial drug resistance gene’s nucleotide and deduced amino acid sequences were analyzed and compared to reported resistance-associated mutations. Allele frequencies were calculated as percentages (number of mutants / total number of tested samples). The comparison of haplotype frequency in each gene population was also calculated. Chi square and 2-tailed Fisher’s exact tests were used to determine the difference in prevalence of mutant alleles of each SNP between years using SPSS (version 29.0.0.0 SPSS Inc., IL, USA) for window. Statistical significance was defined as a P- value < 0.001. In addition, both intergenic and intragenic linkage disequilibrium (LD) tests were performed by calculating the r2 values to determine the association between the SNPs of the Pvdhfr and Pvdhps genes using Haploview software [51]. The strength of the statistical significance of LD between the SNPs was represented by the darkness of the boxes.
Results
Analysis of SNPs in P. vivax genetic markers
A total of 100 samples infected by a single population of P. vivax were obtained in the analysis. All samples were diagnosed by microscopy. The parasitemia ranges of the participants was 800–6,640 parasites / μl. 100 microscopy-positive malarial isolates were then confirmed by nested PCR, which showed P. vivax mono-infections without a mixed infection. Genetic markers Pvmdr1, Pvdhfr, and Pvdhps were successfully amplified and resulted in 100 sequences (S1 File and S1 Table). A summary of the observed allelic distribution of P. vivax markers is given in Tables 2–4 and shown in Fig 1.
Pie charts show haplotype frequencies as percentages among 100 P. vivax isolates. Mutant amino acids are shown in bold with underlined letters.
Pvmdr1.
Three mutations at codons T958M, Y976F, and F1076L were observed in the Pvmdr1gene. The T958M mutation was found in all sequenced isolates (100%), while at codons Y976F and F1076L were detected with frequencies of 21% and 23%, respectively (Table 2). This gave rise to four haplotypes comprised by a single-mutant allele (M958) being the predominant (75%), double-mutant allele (a) (M958F976) (2%), double-mutant allele (b) (M958L1076) (4%), and triple-mutant allele (M958F976L1076) (19%) (Fig 1 and Table 3).
Pvdhfr.
Mutations in the Pvdhfr gene at codons 57, 58, 61, and 117 were observed in 95%, 96%, 95%, and 96%, respectively (Table 2). Analysis of Pvdhfr haplotype revealed four distinct allelic forms, including the WT haplotype (F57S58T61S117), double-mutations (R58N117), quadruple-mutations (a) (I57R58M61T117), and quadruple mutations (b) (L57R58M61T117) in 4%, 1%, 84%, and 11%, respectively (Fig 1 and Table 3). Besides those non-synonymous polymorphisms, synonymous mutations were detected at codon Y69 (TTT→TAC) in the WT allele (S1 Table). In addition, two tandem repeat variations between amino acid positions 88 and 103 of Pvdhfr (Fig 4A) were detected in all isolates (S1 Table). Type 1 or wild type, i.e. three repeated sets of four amino acids (5′-GGDN-3′) was most common in isolates (99%), while type 2, which represents six deleted amino acids at codons 98–103 (5′-THGGDN-3′) were found in one isolate (1%). Type 2 isolate samples also carried double mutations at codons 58 and 117 (R58N117) (S1 Table).
Pvdhps.
Mutations at codon 383, 512, and 553 in the Pvdhps gene were detected in 95%, 47%, and 95%, respectively (Table 2). Haplotype analysis of Pvdhps revealed three distinct allelic forms, including the WT haplotype (A383K512A553), double mutations (G383G553), and triple mutations (G383M512G553) in 5%, 48%, and 47%, respectively (Fig 1 and Table 3). There were no mutations observed at the codon S382 in all isolates.
Frequency of combined Pvdhfr and Pvdhps haplotypes.
The correlations between genotypes of the Pvdhfr and Pvdhps alleles were determined among P. vivax isolates (Fig 1 and Table 4). Four Pvdhfr and three Pvdhps alleles were combined and identified as six distinct haplotypes. The most prevalent combination was a quadruple mutant (a) (I57R58M61T117) of Pvdhfr, combined with a double mutant allele (G383G553) of Pvdhps–forming sextuple mutated haplotypes (48%) of isolates, followed by a triple mutant allele (G383M512G553) of Pvdhps–forming septuple (a) mutated haplotypes (35%). A triple mutant allele (G383M512G553) of Pvdhps combined with a quadruple mutant (b) (L57R58M61T117) allele of Pvdhfr–forming septuple (b) mutated haplotypes was found in 11%, followed by a double mutant allele (R58N117) of Pvdhfr–forming quintuple mutated haplotypes in one isolate. Four of the isolates carried wild-type alleles of both genes (Fig 1 and Table 4).
Linkage disequilibrium (LD) analysis.
To evaluate the LD associations of Pvdhfr and Pvdhps haplotypes, the LD pattern for each SNP in the Pvdhfr and Pvdhps genes was assessed (Fig 2). For Pvdhfr, substitution mutations of T169A, C171A, C174G, C182T and G350C indicated the mutations at codons I57, L57, R58, M61 and T117, respectively. Meanwhile, the C1148G, A1535T and C1658G related to the mutations at codons G383, M512 and G553 of Pvdhps, respectively. Several statistically significant intragenic and intergenic associations were detected among the SNPs pairs in both the Pvdhfr and Pvdhps genes.
For Pvdhfr, substitution mutations of T169A, C171A, C174G, C182T and G350C indicated the mutations at codons I57, L57, R58, M61 and T117, respectively, Meanwhile, the C1148G, A1535T and C1658G related to the mutations at codons G383, M512 and G553 of Pvdhps, respectively. The magnitude of the statistical significance of LD between the SNPs was represented by the extent of darkness in the boxes. Dark gray to black boxes correspond to pairs of SNPs with strong evidence of LD, while light gray to white indicate a lower degree of correlation.
Discussion
Border malaria, particularly along the Thai–Myanmar border area, remains an obstacle to malaria elimination since the risk of malaria reintroduction from the migrant population still exists. Though P. vivax malaria was common along the Thailand–Myanmar border, P. vivax infections were still important causes of hospitalization. Molecular surveillance is therefore important to monitor the prevalence of drug resistance that is associated with drug efficacy in this region. This study provided up-to-date evidence on drug resistance in P. vivax at Kanchanaburi province along Thai–Myanmar border through the analysis of the polymorphisms of antimalarial drug resistance in 2023.
There are increasing worldwide reports of chloroquine treatment failures of P. vivax malaria [29, 52–54]. Though new drugs are now available to replace CQ plus PQ, such as mefloquine (MQ), tafenoquine (TQ), and artemisinin-based combination therapy (ATC) [22], those drugs are more costly than CQ plus primaquine [55]. Alternative treatment regimens such as artemether-lumefantrine for treating CQR in vivax malaria have also been established and evaluated for their antimalarial activities [56]. It was shown that artemether-lumefantrine was as effective as CQ against vivax malaria [56]. These drugs may be used as the first-line drug for the treatment of vivax malaria in a mixed infection of falciparum and vivax malaria areas. However, alternative regimens are now not generally available. These alternative treatment policies have not been proposed based on clinical studies or therapeutic efficacy. Efficient methods are required to assess the potential for resistance to current standard treatments. Molecular markers have not been confirmed yet. Therefore, monitoring P. vivax’s resistance to anti-malarial drugs is still important for an earlier warning system. Unlike P. falciparum, in vitro, studies to assess drug resistance in P. vivax are still difficult to conduct due to the asynchronous blood stage of the parasite, including relapse, recrudescence, and reinfection of this parasite [29]. The molecular mechanisms of drug resistance in P. vivax remain ambiguous. A genetic marker of CQR in P. vivax is also still not clear. The search for CQ resistance markers in P. vivax relied on P. falciparum ortholog, namely Pvmdr1 in P. vivax [57]. However, based on the currently available data on drug resistance markers in P. vivax, Pvmdr1 mutations have been suggested to be candidate markers of resistance to the frontline treatment drug CQ in P. vivax [23, 58]. This study found the T985M mutation in all isolates (100%), similar to a previous report in Thailand that showed the T985M allele is dominant on the Thai–Myanmar and Thai–Cambodia borders [20, 29] (Fig 3A). This allelic variant is fixed in Asia and Africa isolates but is not responsible for CQR in P. vivax [59]. The F1076L mutation showed a prevalence of 23%. There was a decrease in the prevalence of the 1076L mutant allele on the Thai–Myanmar border with statistical significance. In contrast, this allelic variant was still found in a high prevalence on the Thai–Cambodia borders between 2008 and 2014 (Fig 3A, S2 Table) [20]. The mutation Y976F, which has been found an association with decreased susceptibility to CQ in in vitro studies [23, 27] was detected in 21%. This mutation is less common along the Thai-Myanmar border, similar to a previous report [20, 29, 60] compared to the Thai–Cambodia borders. Haplotype analysis showed that the single mutant (M958) is more frequent in this study (75%), a result congruent with previous research in the Thai–Myanmar border [20, 29] (Fig 3B). The prevalence of this haplotype has increased (35.4% in 2008 and 75% in 2023) with statistical significance (Fig 3B, S3 Table). In contrast, at the Thai–Cambodia border, the triple mutant (M958F976L1076) was dominant [20]. The double-mutant (a) (M958F976) and triple mutant (M958F976L1076) alleles that carried the Y976F mutation were detected in both borders. Notably, these two haplotypes were more prevalent along the Thai–Cambodia border than the Thai–Myanmar border (Fig 3B). Even though there are few case reports of CQR P. vivax carried the double (F976L1076) mutations in Thailand [13], CQ remains the drug of choice and an effective treatment for most P. vivax malaria cases in Thailand [12]. CQ-sensitive strains of P. vivax are mostly found in this study. In contrast, a high prevalence of CQR in P. vivax has occurred in Indonesia, Malaysia, and Papua New Guinea, which has been linked with life-threatening complications of malaria due to treatment failure. Reports of drug resistant were found in this study, indicating that resistant phenotypes would continue to spread in these regions. However, the F976L1076 mutations in clinical CQR have not been evaluated yet in patients infected with mutant Plasmodium species. Clinical studies at different geographic sites have reported that Pvmdr1 mutations are not linked with CQR [61, 62]. The functional studies for Pvmdr1 mutations also need to be fully understood. In addition, susceptibility to MQ was significantly lower in P. vivax after the introduction of MQ regimens in Cambodia during 2017–2018 [63]. The associations between susceptibility and mutation in Pvmdr1 were not only in CQR but also in mefloquine resistance (MQR). MQ has been used against CQR and SP-resistant falciparum malaria at the Thai-Cambodian border [64]. MQR was associated with the Pfmdr1 gene in P. falciparum [58]. Ongoing selection of decreased susceptibility to MQ in P. vivax in a neighboring country should be aware. This was evidence of a cross-resistance to other antimalarials due to Pvmdr1 mutations. Thus, close supervision of the efficacy of CQ together with in vitro susceptibility tests to determine at the clinical level in order to confirm gene polymorphism-related drug resistance and therapeutic failure should encourage constant surveillance for monitoring the occurrence of CQR, and possible MQR in P. vivax in Thailand.
Mutant allele and haplotype frequencies of Pvmdr1 (a, b), Pvdhfr (c, d), and Pvdhps (e, f) in P. vivax isolates from a Kanchanaburi province, western Thailand. Data were combined from previously published surveys on the Thai–Myanmar and Thai–Cambodia borders between 2008 [20], 2008/2 [19], 2010 [29], 2011 [21], 2014 [20] and 2023 in this study. Mutant allele frequencies (a, c, e) and haplotype frequencies (b, d, f) are represented as percentages on the y-axis, while the x-axis indicate time (in years). The different bar charts indicate different mutant alleles. Mutant amino acids are shown in bold with underlined letters. The difference in the mutant allele and haplotype frequencies between year was calculated by Chi square and 2-tailed Fisher’s exact test with statistically significant difference at P-value < 0.001.
Due to the wide use of SP to treat mixed P. falciparum and P. vivax infections, the emergence of SP resistance in P. vivax has been reported worldwide. Mutation in the Pvdhfr and Pvdhps has been widely reported and associated with spreading SP-resistance in the P. vivax parasite in countries within the Greater Mekong Sub-region (GMS) [40]. The result in this study aggregated with previously published survey showed high level resistance of P. vivax parasite to SP at Thai–Myanmar border, which was concordant with previous reports [19–21]. For Pvdhfr gene, confer resistance to pyrimethamine, the mutations at codons 57, 58, 61 and 117 were the most prevalent on the Thai–Myanmar border, similar to a previous report from Thailand (Fig 3C) [19–21]. In contrast at the Thai–Cambodia border, codons 58, 61 and 117 were predominant. In this study, the prevalence of mutation frequency at codons 57, 58, 61 and 117 were very similar (95–96%), compared with a previous report on this border region [20, 21] (Fig 3C). Haplotype analysis of Pvdhfr revealed a high prevalence of quadruple mutant (I/L57R58M61T117) (95%), where is commonly prevalent along the Thai–Myanmar border (Fig 3D) [19–21, 65]. Quadruple mutant (a) (I57R58M61T117) was the most prevalent (84%) in this study, followed by quadruple mutant (b) (L57R58M61T117) (11%), similar to those reported at the same border region [19, 20] (Fig 3D). Compared to previous reports at the Thai–Myanmar border, this study found a double-mutant allele (R58N117) in very low frequency (1%) with statistical significance (Fig 3D, S3 Table). On the contrary, a very high prevalence of this allele was observed at the Thai–Cambodia border. Unlike previous reports [20, 21], wild-type alleles (F57S58T61S117) were detected (4%) in this study. This allele carried non-synonymous mutations at codon Y69 (TTT→TAC). Double-mutant allele (R58N117) and quadruple-mutant allele (I/L57R58M61T117) are likely associated with reducing their susceptibility to pyrimethamine [33, 66, 67]. The high prevalence of these mutant alleles in this study agreed with the report of other previous studies [19–21, 68]. Our finding indicated that pyrimethamine resistance is still high over time on the Thai–Myanmar border.
The tandem repeat region between the amino acid positions 82 and 109 was identified in the Pvdhfr gene (Fig 4A). The type 1, with three copies of GGDN repeats, is the majority of the isolates (99%), commonly occurring with quadruple-mutant Pvdhfr alleles [17, 19, 20]. Meanwhile, type 2, with a deletion of six amino acids, was detected in one isolate, and it usually co-existed with the double mutations (R58N117) [17, 19, 20]. Combined with previous surveys of the prevalence of tandem repeat region in Pvdhfr mutations from the Thai–Myanmar border, our results are consistent with previously reported studies [19, 20], where type 1 is predominant (Fig 4B, S4 Table). In contrast, type 2 was found in all isolates (100%) at the Thai–Cambodia border. At the Thai–Myanmar border, Type 2 has decreased (22.6% in 2008, 29.9% in 2014 and 1% in 2023) with statistical significance (Fig 4B, S4 Table). Tandem repeat variations in Pvdhfr are possibly linked with increased resistance to pyrimethamine resistance [19, 65]; however, the mechanism of these repeat regions that confer the risk of drug resistance remains unclear [20].
Data were combined from previously published surveys on the Thai–Myanmar border and Thai–Cambodia borders between 2008 [20], 2008/2 [19], 2014 [20] and 2023 in this study. (a) Sequences alignment of tandem repeat variants in the Pvdhfr gene between amino acid positions 82 and 109. Type 1 contains three copies of GGDN repeats. Type 2 represents deletion of six amino acids from positions 98 to 103. GGDN repeats are shown in text highlight color. The amino acid deletions are shown as Dashes (-). (b) Prevalence of tandem repeat variants are shown as percentages on the y axis, while the x axis indicate time (in years). The difference in the tandem repeat variants frequencies between year was calculated by Chi square and 2-tailed Fisher’s exact tests with statistically significant difference at P-value < 0.001.
For Pvdhps, mutations at codon S382A, A383G, K512M/E, A553G, and V585A have been associated with resistance to sulfadoxine [16, 24, 34, 40]. Among these codons, the single (G383) and double (G383G553) mutant Pvdhps alleles are directly related to sulfadoxine resistance by which a disruption in the affinity between sulfadoxine-binding site in P. vivax [34]. G383 and G553 codons prevailed in 95% of the isolates, similar to that reported by other studies that these codon mutations were the majority from the Thai–Myanmar border [20, 21]. In contrast, codon G383 was more prevalent from the Thai–Cambodia border (Fig 3E). The prevalence of G383 at the Thai–Myanmar border between 2011, 2014 and 2023 increased from 46.3%, 46.3% to 95%, respectively with statistical significance (Fig 3E, S2 Table). Haplotype analysis of Pvdhps revealed a high prevalence of double mutant (G383G553) in this study (48%), a result congruent with previous studies on the Thai–Myanmar border [19, 20]. In contrast, only WT and single mutant (G383) were detected (Fig 3F). In this study, high prevalence of the triple mutant (G383M512G553) Pvdhps alleles were observed in 47%, which is inconsistent with those previous reports at the same border region [19, 20]. Double mutant (G383G553) has been linked with high-level resistance in sulfadoxine compared to the wildtype [69]. Though M512 allele might not involve with sulfadoxine resistance, the double (G383G553) and triple (G383M512G553) Pvdhps mutant alleles that carried mutation at codon 383 and 553 were highly prevalent in this region. It was indicated that the resistance of the P. vivax parasite to sulfadoxine at the Thai–Myanmar border is still high.
Distributions of Pvdhfr and Pvdhps allelic combinations among P. vivax isolates were identified in this study. The combination of Pvdhfr quadruple mutant (I57R58M61T117) and double-mutant (G383G553) Pvdhps alleles–forming sextuple mutated haplotypes was detected as the most prevalence of 48% (Fig 1). The result was concordant with previous reports, in which sextuple mutated haplotypes in the isolates were found in most samples obtained from areas along the Thai–Myanmar border [19]. The clinical response to SP depends on the Pvdhfr and Pvdhps genotype [65]. Sextuple mutations have been associated with high-grade drug resistance, which contributed to the clinical failure of SP treatment in P. vivax infection [70]. Vivax parasites harboring the sextuple mutated haplotypes were eliminated more slowly after receiving SP treatment than less-mutated parasites. Moreover, early treatment failures in patients were linked to infection with multiple mutants in the combination of the Pvdhfr and Pvdhps genes [70]. The mutant Pvdhps M512 allele was found in a high frequency (47%) in this study, while it was not found in the previous study [19]. Therefore, a triple-mutant allele (G383M512G553) of Pvdhps combined a quadruple-mutant (a) (I57R58M61T117), a quadruple-mutant (b) (L57R58M61T117), and a double mutant allele (R58N117) of Pvdhfr was detected in 35%, 11% and 1%, respectively. In contrast, a double mutant (G383G553) Pvdhps combined with a double mutant (R58N117) or a quadruple mutant (b) (L57R58M61T117) Pvdhfr alleles were found at 10.7% and 7.1%, respectively from the previous survey [19]. Additionally, the LD analysis showed statistically significant intragenic and intergenic associations among the SNP pairs between the Pvdhfr and Pvdhps genes (Fig 2). These results indicate that the alleles located in both genes were dependent on each other. Even though SP has never been prescribed against P. vivax in Thailand, the high prevalence of sextuple and septuple mutated haplotypes the Pvdhfr and Pvdhps genes was observed in P. vivax isolates during this study. Sextuple mutations found in this study indicated that the selection pressure came from widespread use of SP in this region. A combination of SP with other antimalarial drugs has been recommended to treat uncomplicated falciparum malaria, where co-infection with P. falciparum and P. vivax is common. The use of SP may have placed the P. vivax parasite under SP drug selection pressure [70, 71]. SP mutations reflect that P. vivax has had exposure to SP in the past. Moreover, it may be a consequence of misdiagnosis of malaria in the absence of clinical suspicion. Misdiagnosed patients might receive unnecessary medication. Self-treatment of P. vivax infection with SP might be associated with SP resistance in endemic areas [20]. Labor migration of the population from neighboring countries along the international borders might also cause the introduction of resistant parasites. This study’s results, combined with a previously published survey on the Thai–Myanmar border, demonstrated that diverse patterns of Pvdhfr and Pvdhps mutant alleles in P. vivax were found and changed over time in this malaria-endemic region. The prevalence and patterns of combined mutations in Pvdhfr and Pvdhps in this study indicated that SP resistance in P. vivax is still prevalent in these areas. The surveillance of the distribution of combined Pvdhfr and Pvdhps polymorphisms is thus essential to evaluate the development and spread of SP resistance in P. vivax in this region.
Though the prevalence of drug resistance in P. vivax parasites on the Thai-Myanmar border in this study was congruent with previously published results from 2008 to 2014 in Thailand, the limitations of this study should be considered. The study samples were included from one geographical area. Only symptomatic patients who had presented to an accessible hospital were obtained for the analysis. The samples from the different provinces within border regions should be enrolled to represent the true prevalence of resistance alleles. The data reported in this study may not be construed as indicating the overall prevalence of malaria in Thailand. Thus, ongoing and comprehensive molecular surveillance, together with using whole-genome sequencing for further studies targeting multiple markers, is required to inform policymakers and enable the stated Thai malaria elimination strategy.
Conclusions
This study reported an up-to-date overview of the prevalence of three potential drug-resistant markers, Pvmdr1, Pvdhfr, and Pvdhps in P. vivax isolates from a Kanchanaburi province, along the Thai–Myanmar border. High mutations in these drug-resistance genes are prevalent in this region. Further assessment of drug resistance from different geographical locations and the expansion of the sample size would be necessary to provide a more representative conclusion of the frequencies of the mutations. Thus, these data will serve as a baseline for further monitoring of drug-resistant malaria.
Supporting information
S1 Table. Analysis of SNPs in P. vivax genetic markers.
https://doi.org/10.1371/journal.pone.0304337.s001
(XLSX)
S2 Table. Prevalence of target mutation conferring antimalarial drug resistance in P. vivax isolates from a Kanchanaburi province, Thailand collected between January to May 2023, combined from previously published surveys on the Thai–Myanmar and Thai–Cambodia borders between 2008, 2010, 2011 and 2014.
https://doi.org/10.1371/journal.pone.0304337.s002
(PDF)
S3 Table. Prevalence of P. vivax Pvmdr1, Pvdhfr and Pvdhps haplotypes in isolates collected from a Kanchanaburi province, Thailand during January to May 2023, combined from previously published surveys on the Thai–Myanmar and Thai–Cambodia borders between 2008, 2008/2 and 2014.
https://doi.org/10.1371/journal.pone.0304337.s003
(PDF)
S4 Table. Prevalence of tandem repeat variants Pvdhfr in isolates collected from a Kanchanaburi province, Thailand during January to May 2023, combined from previously published surveys on the Thai–Myanmar and Thai–Cambodia borders between 2008, 2008/2 and 2014.
https://doi.org/10.1371/journal.pone.0304337.s004
(PDF)
S1 File. The gene sequences reported in this study were shown in FASTA format.
https://doi.org/10.1371/journal.pone.0304337.s005
(RAR)
Acknowledgments
We thank all the participants who provided blood samples for this study. We also thank Dr. James M. Brimson (Research Unit for Innovation & international affairs, Faculty of Allied Health Sciences Chulalongkorn University) for a critical reading and grammar check of this manuscript.
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