Skip to main content

Molecular surveillance of the Plasmodium vivax multidrug resistance 1 gene in Peru between 2006 and 2015

Abstract

Background

The high incidence of Plasmodium vivax infections associated with clinical severity and the emergence of chloroquine (CQ) resistance has posed a challenge to control efforts aimed at eliminating this disease. Despite conflicting evidence regarding the role of mutations of P. vivax multidrug resistance 1 gene (pvmdr1) in drug resistance, this gene can be a tool for molecular surveillance due to its variability and spatial patterns.

Methods

Blood samples were collected from studies conducted between 2006 and 2015 in the Northern and Southern Amazon Basin and the North Coast of Peru. Thick and thin blood smears were prepared for malaria diagnosis by microscopy and PCR was performed for detection of P. vivax monoinfections. The pvmdr1 gene was subsequently sequenced and the genetic data was used for haplotype and diversity analysis.

Results

A total of 550 positive P. vivax samples were sequenced; 445 from the Northern Amazon Basin, 48 from the Southern Amazon Basin and 57 from the North Coast. Eight non-synonymous mutations and three synonymous mutations were analysed in 4,395 bp of pvmdr1. Amino acid changes at positions 976F and 1076L were detected in the Northern Amazon Basin (12.8%) and the Southern Amazon Basin (4.2%) with fluctuations in the prevalence of both mutations in the Northern Amazon Basin during the course of the study that seemed to correspond with a malaria control programme implemented in the region. A total of 13 pvmdr1 haplotypes with non-synonymous mutations were estimated in Peru and an overall nucleotide diversity of π = 0.00054. The Northern Amazon Basin was the most diverse region (π = 0.00055) followed by the Southern Amazon and the North Coast (π = 0.00035 and π = 0.00014, respectively).

Conclusion

This study showed a high variability in the frequencies of the 976F and 1076L polymorphisms in the Northern Amazon Basin between 2006 and 2015. The low and heterogeneous diversity of pvmdr1 found in this study underscores the need for additional research that can elucidate the role of this gene on P. vivax drug resistance as well as in vitro and clinical data that can clarify the extend of CQ resistance in Peru.

Background

In the Americas, Plasmodium falciparum and Plasmodium vivax are responsible for 25.9% and 74.1% of all malaria cases reported in the region, respectively [1]. Although P. falciparum is associated with higher mortality rates, P. vivax is attributed with the highest morbidity in the continent and particularly in the Amazon Basin with a ratio of at least 4:1 cases in relation to P. falciparum [2].

According to the Peruvian CDC, the Northern Amazon Basin accounted for most of all malaria cases reported in 2018 (Loreto region; 95%) followed by the North Coast (Piura and Tumbes Region; 0.02%) and the Southern Amazon Basin (Madre de Dios; 0.01%). Since 2006, the Northern Amazon Basin experienced a reduction in the incidence of P. vivax malaria from 56,171 cases in that year to 26,846 cases in 2010 due to the implementation of the Global fund’s Malaria Project (PAMAFRO), a community programme focused on active case detection and treatment [3]. After the end of the programme in 2010, malaria cases increased each year from 11,779 in 2011 up to 60,268 in 2015.

At the time when samples were collected, there were two treatment schemes for P. falciparum that were administered according to the geographical area. In the North coast, standard treatment consisted of sulfadoxine–pyrimethamine (SP) in combination with artesunate (ART), while in the rest of the country it was changed to ART and mefloquine (MQ) as first-line treatment after a brief period of SP [4, 5].

Cases caused by P. vivax are primarily treated with CQ and primaquine (PQ) in order to eliminate sexual, asexual and dormant P. vivax stages. However, there is accumulating evidence suggesting emergence of P. vivax anti-malarial resistance in Peru. For instance, four cases of P. vivax recurrence on days 21 and 28 after oral anti-malarial treatment with CQ (10 mg/kg on days 0 and 1 and 5 mg/kg on day 2) and PQ (0.5 mg/kg/day for seven days) were described during a study conducted in 177 patients in the Peruvian Amazon region between 1998 and 2001 [6]. Additionally, Graf et al., reported in 2012 a possible case of chloroquine-resistant P. vivax which presented recrudescence on day 28th after treatment with a combination of CQ (25 mg base/kg divided into single daily doses over 3 days) and PQ (0.25 mg/kg daily for 14 days) [7].

Cases of CQ drug resistance in P. vivax were first reported in 1989 in Papua New Guinea [8] that subsequently extended to other endemic regions including South America [9,10,11]. However, the exact mechanisms that modulate drug resistance are still poorly understood. Single nucleotide polymorphisms (SNPs) in pfmdr1, pfdhps, pfdhfr and pfcrt have been shown to be involved in P. falciparum anti-malarial resistance in vivo and in vitro [12,13,14]. Likewise, mutations on the P. vivax orthologous genes pvmdr1, pvdhps, pvdhfr and pvcrt have also been associated with anti-malarial resistance although a clear role in treatment failure remains to be fully elucidated [15,16,17,18,19,20,21,22].

In addition, other studies have shown that changes in gene copy number and expression profiles of pvmdr1 and pvcrt could be associated with CQ resistance [23, 24]. The association in the case of pvcrt is further supported by recent evidence that 5′UTR and intronic changes in pvcrt are linked with increased pvcrt expression in parasite CQ resistant cross lines tested in non-human primate models [25]. In the specific case of pvmdr1, its role in P. vivax resistance is also under discussion in the scientific community with some studies suggesting convergent evolution without an implication in resistance [26,27,28] whereas others propose a direct association with P. vivax resistance [22]. In this regard, studies showed an association between the prevalence of the pvmdr1 976F mutation with increased tolerance to CQ and decreased in vitro susceptibility to MQ and ART in areas with high levels of CQ resistance and the opposite in areas where CQ continues to be used as primary treatment [22, 29, 30]. This evidence suggests the presence of competitive evolutionary pressures on pvmdr1 [29] caused by CQ, MQ and ART and that pvmdr1 mutations might have significant fitness costs [31].

Many of the initial associations of pvmdr1 with resistance to CQ were primarily based on molecular tests and epidemiological or clinical information. However, these assessments suffer from important confounding factors [32]. Stronger evidence have been obtained from studies that combined molecular data from pvmdr1 (SNPs and changes in gene copy number) with in vitro drug susceptibility tests against anti-malarial drugs used in the study area such as CQ and MQ [22, 29] or with clinical data [18, 33].

This study aimed to extend the genetic characterization of pvmdr1 in South America by assessing changes over time in the prevalence of pvmdr1 polymorphisms in samples collected across the Amazon basin and other endemic regions from Peru between 2006 and 2015. The information collected will provide a picture of the dynamics of genetic variability of pvmdr1 in Peru, which can be used as a baseline for surveillance in response to new regional initiatives of malaria control and elimination.

Methods

Study design

Samples were collected from four IRB-approved studies conducted in Peru between 2006 and 2015 in different communities of the Northern Amazon Basin (region of Loreto; 2006–2015), Southern Amazon Basin (region of Madre de Dios; 2012–2013) and North Coast (regions of Piura and Tumbes; 2008–2010) (Fig. 1). Although the Peruvian Amazon and the North Coast of Peru are different environments, they share common characteristics such as proximity to massive water sources like rivers and lakes and having a warm humid climate.

Fig. 1
figure 1

Geographic distribution of the pvmdr1 haplotypes. The graphic was made using 550 samples collected in the Northern Amazon Basin (n = 445), the Southern Amazon Basin (n = 48) and the North Coast (n = 57). Pie charts indicate the pvmdr1 haplotype present in a region and their prevalence. The M908L/T958M haplotype was distributed in all sites while the T958M/F1076L haplotype was specific for the North Coast

Sample collection and preparation

Blood samples were collected by finger prick or venipuncture and thick and thin blood smears were prepared for malaria microscopy and read at the Naval Medical Research Unit 6 (NAMRU-6) facilities. DNA was extracted from either 200 µl venous blood or ~ 100 µl dried blood spots using the QIAamp® DNA Blood Mini Kit (GmbH, Hilden, Germany) following the manufacturer’s instructions and stored at − 20 °C. Plasmodium vivax monoinfection was confirmed by polymerase chain reaction of the 18S small subunit ribosomal RNA gene (18S SSU rRNA) as previously described [34].

Identification of single nucleotide polymorphisms (SNP) in the pvmdr1 gene

The whole pvmdr1 gene was amplified by PCR in three overlapping fragments (Additional file 1: Fig. S1, Additional file 2: Table S1). PCR reaction was carried out in a 50 µl reaction volume containing 5 µl of gDNA (~ 25 ng), 1X buffer, 2 mM MgCL2, 125 µM dNTP’s, 250 nM of each primer and 1 unit of Taq Polymerase (Invitrogen). PCR products were purified using QIAquick™ silica‐based spin columns (GmbH, Hilden, Germany) and 30 ng of purified PCR product was used to amplify in a standard reaction (20 µl) of BigDye™ Terminator v3.1. The resulting products were sequenced on an ABI3130xl (Applied Biosystems) and analysed in the program Sequencher 4.1 (Gen Codes Corporation) using the reference Sal I strain of P. vivax (Salvador I, GenBank accession number AY571984).

Data analysis

Data was entered in Microsoft Excel and analysed in STATA 13.0 for Windows. Absolute and relative frequency of mutant and wild type alleles and haplotypes of the pvmdr1 gene were calculated for each region. Chi square and 2-tailed Fisher’s exact tests were used to assess statistical significant differences in proportions according to geographic regions. Nucleotide diversity, Tajima´s D test and FST values were calculated in dnaSP v5.1 [28, 35]. In addition, Mega v7.0 was used to assess for natural selection using the modified Nei-Gojobori method [36]. Finally, a haplotype network was constructed in PopArt 1.7 [37] to assess the relatedness of all isolates based on their pvmdr1 sequences.

Results

Polymorphisms on the pvmdr1 gene

The pvmdr1 gene was sequenced on 550 Plasmodium vivax samples with no mixed genotypes from the three study areas. The majority of sequenced samples were collected from the Northern Amazon Basin (n = 445, 80.9%) followed by the Northern Coast (n = 57, 10.4%) and the Southern Amazon basin (n = 48, 8.7%).

A total of eleven mutations were found in pvmdr1 (eight non-synonymous and three synonymous mutations) (Additional file 3: Table S3). The non-synonymous mutations were L186W (2.5%), V221L (12.4%), D500N (5.5%), M908L (99.1%), T958M (94.0%), Y976F (7.5%), F1070L (4.0%) and F1076L (10.5%). From all these mutations, statistical significant differences on the prevalence of SNPs across regions were found on five polymorphisms (Fig. 2 and Table 1).

Fig. 2
figure 2

Haplotype network for P. vivax mdr1 using 8 non-synonymous mutations for the study areas of Peru (n = 550). Each circle represent an independent haplotype, the lines connect nearby haplotypes and the cross line represent one non-synonymous mutation

Table 1 Prevalence of nonsynonymous mutations by regions 2006–2015

The Y976F and F1076L mutations that are associated with CQ resistance were present in the Northern Amazon Basin in the 908L/958 M/976F/1076L and 908L/958 M/1076L haplotypes (Y976F + F1076L = 9.2%, F1076L = 3.4%) and in the 908L/958 M/1076L and 958 M/1076L haplotypes from the Southern Amazon Basin (F1076L = 2.1%) (Additional file 3: Table S3). Synonymous mutations on pvmdr1 were found at positions T529 (ACA > ACG), L1022 (CTA > TTA) and K1355 (AAA > AAG), which were present in 57.8%, 18.0% and 3.4% of the samples, respectively.

The frequencies of the Y976F and F1076L mutations fluctuated over time during the course of the study (Fig. 3). In this regard, there was a continuing decrease in the distribution of both mutations from 17.8% in 2006 to less than 2% in 2009. The highest prevalence for both mutations was recorded in 2011 (Y976F = 16.7% and F1076L = 20.8%) followed by a downward trend until 2013 (Y976F = 4% and F1076L = 6.7%).

Fig. 3
figure 3

Fluctuation of non-synonymous mutations in the Northern Amazon Basin between 2006 and 2015. The graphic shows the dynamics of the 976F and 1076L polymorphisms and the number of reported cases in the Northern Amazon Basin. The “y” axis on the left shows the number of reported cases for the barplot. The “y” axis on the right depicts the percentage of the 976F and 1076L mutations for the lineplot whereas the “x” axis indicate the years. (1) Global fund’s Malaria project “PAMAFRO” (2005–2010). Plasmodium vivax data from 2010 was not included in the graphic because of the low sample size for that year (n = 3)

Geographic distribution of pvmdr1 haplotypes

Twenty-seven haplotypes consisting of either synonymous and non-synonymous mutations were identified in pvmdr1 (Additional file 3: Table S3). Out of those, 13 haplotypes consisting of non-synonymous mutation were identified on pvmdr1 with all haplotypes but one presenting at least one non-synonymous mutation (Table 2).

Table 2 Prevalence of haplotypes by regions 2006–2015

Significant differences were found in the distribution of the 908L/958M haplotype among the three sites (p < 0.05). The Northern Amazon Basin presented the highest number of different haplotypes (92.3%) followed by the Southern Amazon basin (23%) and the North Coast (23%) (Fig. 1, Table 2). A common haplotype that was differentially (p < 0.05) distributed in all sites was M908L/T958M, which accounted for 61.1% of cases in the Northern Amazon Basin, 95.8% in the Southern Amazon and 80.7% in the North coast.

Single mutation was observed in 11 isolates (2%), double mutation in 389 (70.7%), triple mutation in 90 (16.2%) and quadruple mutation in 59 samples (10.7%). The F1076L mutation was present in three haplotypes (10.6% of the samples), while the combination Y976F/F1076L was present in one haplotype (908L/958M/976F/1076L) exclusive from the Northern Amazon Basin (7.5% of the samples).

The haplotype network of pvmdr1 that comprised 8 non-synonymous mutations showed that all the haplotypes were closely related with most of them being only one mutational step from each other (Fig. 2).

Genetic diversity of the pvmdr1 locus

The overall nucleotide diversity of pvmdr1 in Peru was π = 0.00054 with differences according to geographical location. In this regard, the Northern Amazon Basin was the most diverse (π = 0.00055) followed by the Southern Amazon basin (π = 0.00035) and the North coast (π = 0.00014).

FST values were high across all regions: 0.399 between the North coast and the Southern Amazon basin, 0.227 between North Coast and the Northern Amazon Basin, and 0.316 between the Southern Amazon basin and the Northern Amazon Basin. Moreover, Tajima´s D test was positive (1.321) although not statistically significant when samples were analyzed globally. At the regional level, Tajima’s D was also not significant across all regions, the Northern Amazon Basin (2.072), Southern Amazon basin (0.925) and North coast (− 0.612). This is also supported by a non-significant dN-dS ratio for Peru (− 0.709; P = 1.000) and at the regional level: the Northern Amazon Basin (− 0.720; P = 1.000), the North Coast (− 1.106; P = 1.000) and Southern Amazon Basin (1.587; P = 0.058).

Discussion

Different studies indicate that P. vivax has recently suffered an important evolutionary pressure driven by the use of antifolate drugs [38, 39]. Furthermore, in areas with P. vivax CQ susceptibility and P. falciparum CQ resistance, P. vivax is subjected to indirect selection pressures during the treatment of P. falciparum or mixed infections [40].

Although most mutations in pvmdr1 play no role in parasite resistance, the Y976F and F1076L mutations are still controversial in the scientific community due to conflicting evidence [17, 18, 21, 22, 24, 41]. For instance, Thailand, whose anti-malarial treatment regime for P. vivax is based in CQ and PQ as in Peru [42], presented a frequency of 25% of the Y976F mutation [22], 21% of pvmdr1 amplifications [29], CQ susceptibility and reduced susceptibility to MQ and ART in P. vivax [43]

In contrast, countries such as Indonesia or New Guinea showed up to 95% of the 976F mutation in pvmdr1, no evidence of pvmdr1 amplification, CQ resistance and susceptibility to MQ and ART [29]. Furthermore, the Y976F mutation was associated with a fourfold higher chloroquine IC50 and 5 to 8 folds lower IC50 for ART and MQ, respectively [22]. A similar situation has been reported in French Guyana that presented a decrease in pvmdr1 amplification from 71.3% when MQ was used against P. falciparum (1995–2002) to 12.8% after a change in the anti-malarial regime. This change was accompanied by an increase in 976F haplotypes after the use of MQ [40]. Unfortunately, the present study was not able to assess changes in pvmdr1 copy number and evaluate if there was a relation between 976F haplotypes and pvmdr1 amplification.

In the case of Peru, it appears that drug pressure and changes in the incidence of malaria have also affected the prevalence of the Y976F and F1076L genotypes in the Northern Amazon Basin, which decreased from 17.8% in 2006 to 1.2% in 2008. This decrease coincides with the implementation of a radical P. vivax treatment scheme in 2005 in the Northern Peruvian Amazon, which changed from 3 days of CQ and 14 days of PQ (0.25 mg/kg/day) to 3 days of CQ and 7 days of PQ (0.5 mg/kg/day) [2, 44]. Therefore, it is highly likely that the new treatment scheme coupled with increased access to diagnosis and treatment accessibility provided by PAMAFRO could have exerted strong selection pressures in the P. vivax population. This is further supported by the fact that Peru reached a malaria incidence rate lower than 1 case/1,000 inhabitants towards the end of PAMAFRO in 2010 [2]. However, after the end of PAMAFRO, the frequency of the Y976F and F1076L genotypes radically increased until reaching 20.83% in 2011 to initiate a gradual reduction down to 5% in 2015. This rapid fluctuation in the frequencies of these genotypes could be due to local variability in malaria control activities rather than reversion of the mutated genotype.

The 908L/958M haplotype that was the most frequent in our study presented a continuous increase in the Northern Amazon Basin from 48% in 2006 to 71% in 2015. No major changes in the frequencies of the other haplotypes were found. In this regard, studies carried out in P. falciparum suggest that CQ-wildtype alleles in pfmdr1 and pfcrt have a selective advantage over resistant genotypes when CQ pressure is not exerted [45,46,47]. In this regard, regions where CQ was discontinued experienced an increase of wildtype pfcrt and pfmdr1 P. falciparum strains [45, 46, 48]. However, this is unlikely to happen with the P. vivax population in the Peruvian Amazon due to the continued use of CQ as first-line treatment. Therefore, it is possible that the changes in the prevalence of pvmdr1 genotypes correspond to expansion or contraction of the wildtype parasite population in response to local and temporal variations of malaria control.

The M908L and T958M mutations that are two of the most frequent polymorphisms in our study (99.1% and 94.0%, respectively) were reported with 100% prevalence in Thailand and Madagascar and with 28% and 100% prevalence in Brazil, respectively [9, 15, 24, 49]. Also, these studies suggested that these polymorphisms do not have any association with resistance to chloroquine and are likely to be related to the evolutionary history of P. vivax [9, 15, 24, 26, 27, 49].

In terms of diversity, the Northern Amazon Basin presented 12 of the 13 haplotypes found in Peru being the region with the highest diversity for pvmdr1 (π = 0.00055) compared to the global value of pvmdr1 in Peru (π = 0.00054) [50]. This level of diversity is lower to the ones from other endemic areas such as India (π = 0.0012), Brazil (π = 0.0016) and Ecuador (π = 0.0009) [28]. In this regard, the low diversity in Peru and Ecuador could be explained by frequent inbreeding and low recombination rates which are characteristic of low transmission regions in contrast to the high recombination rates between P. vivax genotypes in India and Brazil [39, 51, 52].

The North Coast (π = 0.00014) and Southern Amazon Basin (π = 0.00035) had a lower diversity which is consistent with previous studies in the North Coast that showed a low parasite diversity and a homogeneous population structure [52]. This is likely due to the presence of the Andes mountain range that acts as a geographical barrier that blocks migration of infected Amazonian vectors. Another factor that could influence the low parasite diversity in the North Coast is the drastic fluctuation of malaria prevalence over time due to the El Niño Southern Oscillation, the presence of a different malaria vector than the one in the Peruvian Amazon basin and to the low human migration between both sites [2].

In the case of the Southern Amazon basin, distribution and transmission of vector-borne diseases including malaria has suffered rapid fluctuations during the last decade due to illegal mining, logging and agriculture, which have drastically changed the environment [2, 53]. In addition, it is also possible that differences in diversity could be explained by the genetic background of circulating populations in the distinct regions of Peru.

However, it is important to consider that a single marker and a potential target under selective pressure might have low resolution to accurately estimate genetic diversity. This is particularly important in low transmission regions such as Peru where there is already strong evidence of low genetic diversity. Therefore, further studies targeting multiple markers or using next generation sequencing are needed to confirm our results for pvmdr1.

Finally, although there is a strong association between molecular markers and resistance to anti-malarials in P. falciparum, this relationship is not clear for P. vivax and its orthologs [22, 28, 47]. This demands further research to assess their effect at the clinical level in order to confirm variants associated with resistance and therapeutic failure. Furthermore, continuous monitoring of pvmdr1 together with in vitro susceptibility tests would help to assess changes in the transmission of malaria overtime as a result of human and environmental variables to support initiatives for malaria control and elimination.

Conclusion

This research showed a regional diversification of pvmdr1 across endemic malaria regions in Peru and changes over time in the frequency of pvmdr1 genotypes in the Northern Amazon Basin between 2006 and 2015. This information is relevant for future epidemiological surveillance to measure the emergence of resistance and changes on the parasite population in this and other endemic areas. However, additional research is needed to elucidate the role of pvmdr1 in P. vivax resistance complemented with ex vivo and phenotypic data.

Availability of data and materials

All data generated and/or analyzed during this study are included in this published article and its additional files.

Abbreviations

IRB:

Institutional review board

PAMAFRO:

Global Fund Malaria Project

PCR:

Polymerase chain reaction

CQ:

Chloroquine

PQ:

Primaquine

ART:

Artesunate

SP:

Sulfadoxine-pyrimethamine

MQ:

Mefloquine

References

  1. WHO Global malaria programme. World malaria report 2017. Geneva: World Health Organization; 2018.

    Google Scholar 

  2. Rosas-Aguirre A, Gamboa D, Manrique P, Conn JE, Moreno M, Lescano AG, et al. Epidemiology of Plasmodium vivax malaria in Peru. Am J Trop Med Hyg. 2016;95:133–44.

    PubMed  PubMed Central  Article  Google Scholar 

  3. Isturiz O, Arias K, Restrepo B, Rosas-Aguirre A, Vargas D, Alvariño G. Compartiendo lecciones aprendidas. Proyecto control de malaria en zonas fronterizas de la región andina: un enfoque comunitario-PAMAFRO. 2009.

  4. Bacon DJ, McCollum AM, Griffing SM, Salas C, Soberon V, Santolalla M, et al. Dynamics of malaria drug resistance patterns in the Amazon basin region following changes in Peruvian national treatment policy for uncomplicated malaria. Antimicrob Agents Chemother. 2009;53:2042–51.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  5. Ruebush TK 2nd, Neyra D, Cabezas C. Modifying national malaria treatment policies in Peru. J Public Health Policy. 2004;25:328–45.

    PubMed  Article  Google Scholar 

  6. Ruebush TK 2nd, Zegarra J, Cairo J, Andersen EM, Green M, Pillai DR, et al. Chloroquine-resistant Plasmodium vivax malaria in Peru. Am J Trop Med Hyg. 2003;69:548–52.

    PubMed  Article  Google Scholar 

  7. Graf PC, Durand S, Alvarez Antonio C, Montalvan C, Galves Montoya M, Green MD, et al. Failure of supervised chloroquine and primaquine regimen for the treatment of Plasmodium vivax in the Peruvian Amazon. Malar Res Treat. 2012;2012:936067.

    PubMed  PubMed Central  Google Scholar 

  8. Whitby M, Wood G, Veenendaal JR, Rieckmann K. Chloroquine-resistant Plasmodium vivax. Lancet. 1989;2:1395.

    CAS  PubMed  Article  Google Scholar 

  9. Marques MM, Costa MR, Santana Filho FS, Vieira JL, Nascimento MT, Brasil LW, et al. Plasmodium vivax chloroquine resistance and anemia in the western Brazilian Amazon. Antimicrob Agents Chemother. 2014;58:342–7.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  10. Sumawinata IW, Bernadeta, Leksana B, Sutamihardja A, Purnomo, Subianto B, et al. Very high risk of therapeutic failure with chloroquine for uncomplicated Plasmodium falciparum and P. vivax malaria in Indonesian Papua. Am J Trop Med Hyg. 2003;68:416–20.

  11. Waheed AA, Ghanchi NK, Rehman KA, Raza A, Mahmood SF, Beg MA. Vivax malaria and chloroquine resistance: a neglected disease as an emerging threat. Malar J. 2015;14:146.

    PubMed  PubMed Central  Article  Google Scholar 

  12. Juma DW, Omondi AA, Ingasia L, Opot B, Cheruiyot A, Yeda R, et al. Trends in drug resistance codons in Plasmodium falciparum dihydrofolate reductase and dihydropteroate synthase genes in Kenyan parasites from 2008 to 2012. Malar J. 2014;13:250.

    PubMed  PubMed Central  Article  Google Scholar 

  13. Rouhani M, Zakeri S, Pirahmadi S, Raeisi A, Djadid ND. High prevalence of pfdhfr-pfdhps triple mutations associated with anti-malarial drugs resistance in Plasmodium falciparum isolates seven years after the adoption of sulfadoxine-pyrimethamine in combination with artesunate as first-line treatment in Iran. Infect Genet Evol. 2015;31:183–9.

    CAS  PubMed  Article  Google Scholar 

  14. Inoue J, Lopes D, do Rosario V, Machado M, Hristov AD, Lima GF, et al. Analysis of polymorphisms in Plasmodium falciparum genes related to drug resistance: a survey over four decades under different treatment policies in Brazil. Malar J. 2014;13:372.

  15. Barnadas C, Ratsimbasoa A, Tichit M, Bouchier C, Jahevitra M, Picot S, et al. Plasmodium vivax resistance to chloroquine in Madagascar: clinical efficacy and polymorphisms in pvmdr1 and pvcrt-o genes. Antimicrob Agents Chemother. 2008;52:4233–40.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  16. Brega S, Meslin B, de Monbrison F, Severini C, Gradoni L, Udomsangpetch R, et al. Identification of the Plasmodium vivax mdr-like gene (pvmdr1) and analysis of single-nucleotide polymorphisms among isolates from different areas of endemicity. J Infect Dis. 2005;191:272–7.

    CAS  PubMed  Article  Google Scholar 

  17. Chung DI, Jeong S, Dinzouna-Boutamba SD, Yang HW, Yeo SG, Hong Y, et al. Evaluation of single nucleotide polymorphisms of pvmdr1 and microsatellite genotype in Plasmodium vivax isolates from Republic of Korea military personnel. Malar J. 2015;14:336.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  18. Fernandez-Becerra C, Pinazo MJ, Gonzalez A, Alonso PL, del Portillo HA, Gascon J. Increased expression levels of the pvcrt-o and pvmdr1 genes in a patient with severe Plasmodium vivax malaria. Malar J. 2009;8:55.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  19. Gama BE, Oliveira NK, Souza JM, Daniel-Ribeiro CT, Ferreira-da-Cruz MF. Characterisation of pvmdr1 and pvdhfr genes associated with chemoresistance in Brazilian Plasmodium vivax isolates. Mem Inst Oswaldo Cruz. 2009;104:1009–11.

    CAS  PubMed  Article  Google Scholar 

  20. Golassa L, Erko B, Baliraine FN, Aseffa A, Swedberg G. Polymorphisms in chloroquine resistance-associated genes in Plasmodium vivax in Ethiopia. Malar J. 2015;14:164.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  21. Mint Lekweiry K, Ould Mohamed Salem Boukhary A, Gaillard T, Wurtz N, Bogreau H, Hafid JE, et al. Molecular surveillance of drug-resistant Plasmodium vivax using pvdhfr, pvdhps and pvmdr1 markers in Nouakchott, Mauritania. J Antimicrob Chemother. 2012;67:367–74.

  22. Suwanarusk R, Russell B, Chavchich M, Chalfein F, Kenangalem E, Kosaisavee V, et al. Chloroquine resistant Plasmodium vivax: in vitro characterisation and association with molecular polymorphisms. PLoS ONE. 2007;2:e1089.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  23. Lu F, Lim CS, Nam DH, Kim K, Lin K, Kim TS, et al. Genetic polymorphism in pvmdr1 and pvcrt-o genes in relation to in vitro drug susceptibility of Plasmodium vivax isolates from malaria-endemic countries. Acta Trop. 2011;117:69–75.

    CAS  PubMed  Article  Google Scholar 

  24. Rungsihirunrat K, Muhamad P, Chaijaroenkul W, Kuesap J, Na-Bangchang K. Plasmodium vivax drug resistance genes; Pvmdr1 and Pvcrt-o polymorphisms in relation to chloroquine sensitivity from a malaria endemic area of Thailand. Korean J Parasitol. 2015;53:43–9.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  25. Sa JM, Kaslow SR, Moraes Barros RR, Brazeau NF, Parobek CM, Tao D, et al. Plasmodium vivax chloroquine resistance links to pvcrt transcription in a genetic cross. Nat Commun. 2019;10:4300.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  26. Sa JM, Nomura T, Neves J, Baird JK, Wellems TE, del Portillo HA. Plasmodium vivax: allele variants of the mdr1 gene do not associate with chloroquine resistance among isolates from Brazil, Papua, and monkey-adapted strains. Exp Parasitol. 2005;109:256–9.

    CAS  PubMed  Article  Google Scholar 

  27. Schousboe ML, Ranjitkar S, Rajakaruna RS, Amerasinghe PH, Morales F, Pearce R, et al. Multiple origins of mutations in the mdr1 gene--A Putative marker of chloroquine resistance in P. vivax. PLoS Negl Trop Dis. 2015;9:e0004196.

  28. González-Cerón L, Montoya A, Corzo-Gómez JC, Cerritos R, Santillán F, Sandoval MA. Genetic diversity and natural selection of Plasmodium vivax multi-drug resistant gene (pvmdr1) in Mesoamerica. Malar J. 2017;16:261.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  29. Suwanarusk R, Chavchich M, Russell B, Jaidee A, Chalfein F, Barends M, et al. Amplification of pvmdr1 associated with multidrug-resistant Plasmodium vivax. J Infect Dis. 2008;198:1558–64.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  30. Vargas-Rodríguez RdCM, da Silva Bastos M, Menezes MJ, Orjuela-Sánchez P, Ferreira MU. Single-nucleotide polymorphism and copy number variation of the multidrug resistance-1 locus of Plasmodium vivax: local and global patterns. Am J Trop Med Hyg. 2012;87:813–21.

  31. Auburn S, Serre D, Pearson RD, Amato R, Sriprawat K, To S, et al. Genomic analysis reveals a common breakpoint in amplifications of the Plasmodium vivax multidrug resistance 1 locus in Thailand. J Infect Dis. 2016;214:1235–42.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  32. Russell B, Chalfein F, Prasetyorini B, Kenangalem E, Piera K, Suwanarusk R, et al. Determinants of in vitro drug susceptibility testing of Plasmodium vivax. Antimicrob agents chemother. 2008;52:1040–5.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  33. Melo GC, Monteiro WM, Siqueira AM, Silva SR, Magalhaes BM, Alencar AC, et al. Expression levels of pvcrt-o and pvmdr-1 are associated with chloroquine resistance and severe Plasmodium vivax malaria in patients of the Brazilian Amazon. PLoS ONE. 2014;9:e105922.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  34. Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, et al. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parasitol. 1993;61:315–20.

  35. Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bioinformatics. 2009;25:1451–2.

    CAS  Article  Google Scholar 

  36. Kumar S, Stecher G, Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol Biol Evol. 2016;33:1870–4.

  37. Leigh JW, Bryant D. popart: full-feature software for haplotype network construction. Methods Ecol Evol. 2015;6:1110–6.

    Article  Google Scholar 

  38. Hupalo DN, Luo Z, Melnikov A, Sutton PL, Rogov P, Escalante A, et al. Population genomics studies identify signatures of global dispersal and drug resistance in Plasmodium vivax. Nat Genet. 2016;48:953–8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  39. de Oliveira TC, Rodrigues PT, Menezes MJ, Goncalves-Lopes RM, Bastos MS, Lima NF, et al. Genome-wide diversity and differentiation in New World populations of the human malaria parasite Plasmodium vivax. PLoS Negl Trop Dis. 2017;11:e0005824.

    PubMed  PubMed Central  Article  Google Scholar 

  40. Faway E, Musset L, Pelleau S, Volney B, Casteras J, Caro V, et al. Plasmodium vivax multidrug resistance-1 gene polymorphism in French Guiana. Malar J. 2016;15:540.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  41. Kittichai V, Nguitragool W, Ngassa Mbenda HG, Sattabongkot J, Cui L. Genetic diversity of the Plasmodium vivax multidrug resistance 1 gene in Thai parasite populations. Infect Genet Evol. 2018;64:168–77.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  42. Recht J, Siqueira AM, Monteiro WM, Herrera SM, Herrera S, Lacerda MVG. Malaria in Brazil, Colombia, Peru and Venezuela: current challenges in malaria control and elimination. Malar J. 2017;16:273.

    PubMed  PubMed Central  Article  Google Scholar 

  43. Khim N, Andrianaranjaka V, Popovici J, Kim S, Ratsimbasoa A, Benedet C, et al. Effects of mefloquine use on Plasmodium vivax multidrug resistance. Emerg Infect Dis. 2014;20:1637–44.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  44. Durand S, Cabezas C, Lescano AG, Galvez M, Gutierrez S, Arrospide N, et al. Efficacy of three different regimens of primaquine for the prevention of relapses of Plasmodium vivax malaria in the Amazon Basin of Peru. Am J Trop Med Hyg. 2014;91:18–26.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  45. Frosch AE, Laufer MK, Mathanga DP, Takala-Harrison S, Skarbinski J, Claassen CW, et al. Return of widespread chloroquine-sensitive Plasmodium falciparum to Malawi. J Infect Dis. 2014;210:1110–4.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  46. Balikagala B, Sakurai-Yatsushiro M, Tachibana SI, Ikeda M, Yamauchi M, Katuro OT, et al. Recovery and stable persistence of chloroquine sensitivity in Plasmodium falciparum parasites after its discontinued use in Northern Uganda. Malar J. 2020;19:76.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  47. Mekonnen SK, Aseffa A, Berhe N, Teklehaymanot T, Clouse RM, Gebru T, et al. Return of chloroquine-sensitive Plasmodium falciparum parasites and emergence of chloroquine-resistant Plasmodium vivax in Ethiopia. Malar J. 2014;13:244.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  48. Huang B, Wang Q, Deng C, Wang J, Yang T, Huang S, et al. Prevalence of crt and mdr-1 mutations in Plasmodium falciparum isolates from Grande Comore island after withdrawal of chloroquine. Malar J. 2016;15:414.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  49. Rungsihirunrat K, Keusap J, Chaijaroenkul W, Mungthin M, Na-Bangchang K. Pvmdr1 polymorphisms of Plasmodium vivax isolates from malaria endemic areas in southern provinces of Thailand. Thai J Pharmacol. 2015;37:5–12.

    Google Scholar 

  50. Van den Eede P, Van der Auwera G, Delgado C, Huyse T, Soto-Calle VE, Gamboa D, et al. Multilocus genotyping reveals high heterogeneity and strong local population structure of the Plasmodium vivax population in the Peruvian Amazon. Malar J. 2010;9:151.

    PubMed  PubMed Central  Article  Google Scholar 

  51. Ngassa Mbenda HG, Wang M, Guo J, Siddiqui FA, Hu Y, Yang Z, et al. Evolution of the Plasmodium vivax multidrug resistance 1 gene in the Greater Mekong Subregion during malaria elimination. Parasit Vectors. 2020;13:67.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  52. Ventocilla JA, Nunez J, Tapia LL, Lucas CM, Manock SR, Lescano AG, et al. Genetic Variability of Plasmodium vivax in the North Coast of Peru and the Ecuadorian Amazon Basin. Am J Trop Med Hyg. 2018;99:27–32.

    PubMed  PubMed Central  Article  Google Scholar 

  53. Conn JE, Moreno M, Saavedra M, Bickersmith SA, Knoll E, Fernandez R, et al. Molecular taxonomy of Anopheles (Nyssorhynchus) benarrochi (Diptera: Culicidae) and malaria epidemiology in southern Amazonian Peru. Am J Trop Med Hyg. 2013;88:319–24.

    PubMed  PubMed Central  Article  Google Scholar 

Download references

Acknowledgments

We would like to express our gratitude to “Dirección Regional de Salud Piura” for the logistic support and in executing the blood samples collection for the “NMRCD.2010.0010” protocol.

AGL is sponsored by the training grant D43 TW007393 awarded by the Fogarty International Center of the US National Institutes of Health awarded to Emerge, the Emerging Diseases and Climate Change Research Unit of the School and Public Health Administration at Universidad Peruana Cayetano Heredia. Meddly Santolalla received a scholarship from CAPES – Coordination for the Improvement of Higher Education Personnel. Jorge L. Maguiña is a doctoral student studying Epidemiological Research at Universidad Peruana Cayetano Heredia under FONDECYT/CIENCIACTIVA scholarship EF033-235-2015 and supported by training grant D43 TW007393 awarded by the Fogarty International Center of the US National Institutes of Health.

We thank to Dionicia Gamboa, PhD and Joseph Vinetz, MD and the investigators, field team members and study participants from the Amazonia Center of Excellence in Malaria Research, funded by cooperative agreement U19AI089681 from the United States Public Health Service, NIH/NIAID.

Funding

This study was supported by funding from the US Department of Defense Health Agency-Armed Forces Health Surveillance Division (AFHSD), Global Emerging Infection Surveillance (GEIS) under PROMIS ID P0106_18_N6_02.

Author information

Authors and Affiliations

Authors

Contributions

FV, JM, MS, JA, AL, participated in all components of the study; design, interpretation and data analysis. EP, blood sample collection. DB, HV performed the genetic analysis. All authors contributed to the writing of the manuscript. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Fredy E. Villena.

Ethics declarations

Ethics approval and consent to participate

The study protocols were approved by the Naval Medical Research Center Institutional Review Board in compliance with all applicable Federal regulations governing the protection of human subjects (protocols NMRCD.2010.0010, NMRCD.2007.0004, NAMRU6.2008.0004, NMRCD.2005.0005 and NMRCD.2012.0006).

Consent for publication

Not relevant.

Competing interests

The authors declare that no competing interests exist.

Disclaimer

The views expressed in this article are those of the authors and do not necessarily reflect the official policy or position of the Department of the Navy, Department of Defense, nor the U.S. Government.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Additional file 1: Fig. S1.

Amplified and sequenced regions of the pvmdr1 gene.

Additional file 2: Table S1.

Primers for conventional PCR, Nested PCR and sequencing ofpvmdr1 gene.

Additional file 3: Table S3.

Prevalence of haplotypes consisting of synonymous and nonsynonymousmutation.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Villena, F.E., Maguiña, J.L., Santolalla, M.L. et al. Molecular surveillance of the Plasmodium vivax multidrug resistance 1 gene in Peru between 2006 and 2015. Malar J 19, 450 (2020). https://doi.org/10.1186/s12936-020-03519-8

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s12936-020-03519-8

Keywords

  • Malaria
  • Plasmodium vivax
  • Single nucleotide polymorphisms
  • Drug resistance