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Antimalarial Drug Resistance: Literature Review and Activities and Findings of the ICEMR Network

Liwang Cui, Sungano Mharakurwa, Daouda Ndiaye and 2 more

The American Journal of Tropical Medicine and Hygiene · 2015 · Review

doi:10.4269/ajtmh.15-0007

Abstract

Antimalarial drugs are key tools for the control and elimination of malaria. Recent decreases in the global malaria burden are likely due, in part, to the deployment of artemisinin-based combination therapies. Therefore, the emergence and potential spread of artemisinin-resistant parasites in southeast Asia and changes in sensitivities to artemisinin partner drugs have raised concerns. In recognition of this urgent threat, the International Centers of Excellence for Malaria Research (ICEMRs) are closely monitoring antimalarial drug efficacy and studying the mechanisms underlying drug resistance. At multiple sentinel sites of the global ICEMR network, research activities include clinical studies to track the efficacies of antimalarial drugs, ex vivo/in vitro assays to measure drug susceptibilities of parasite isolates, and characterization of resistance-mediating parasite polymorphisms. Taken together, these efforts offer an increasingly comprehensive assessment of the efficacies of antimalarial therapies, and enable us to predict the emergence of drug resistance and to guide local antimalarial drug policies. Here we briefly review worldwide antimalarial drug resistance concerns, summarize research activities of the ICEMRs related to drug resistance, and assess the global impacts of the ICEMR programs.

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Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 20, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Despite important gains in some areas, malaria remains a major problem in most of the tropical world, and it continues to cause hundreds of millions of illnesses and hundreds of thousands of deaths each year. Most serious illnesses and deaths from malaria and also most drug-resistant infections are due to infection with Plasmodium falciparum, the most virulent human malaria parasite. In addition, there is increasing appreciation of the importance of Plasmodium vivax, the other common human malaria parasite, as a cause of serious illnesses, and its resistance to antimalarial drugs is now well described.

The control and eventual eradication of malaria depend on a small set of tools. For control of anopheline mosquito vectors the values of insecticide-impregnated bednets and indoor residual spraying of insecticides have been clearly demonstrated, but their efficacy will be limited without coincident efforts directed against malaria parasites. An effective vaccine against malaria would be extremely valuable. Unfortunately, although the RTS,S vaccine, which has offered modest protection against malaria in African children, may be available in a few years, no highly effective vaccine is on the horizon. Thus, appropriate use of antimalarial drugs remains a cornerstone of malaria control. Drugs have two key roles for malaria control. First, prompt and effective treatment of malaria prevents progression to severe disease and limits the development of gametocytes, thus blocking transmission to mosquitoes. Second, drugs can be used to prevent malaria in endemic populations, including various strategies of chemoprophylaxis, intermittent preventive therapy, and mass drug administration.

The ICEMR network includes 10 groups focused on malaria surveillance and related activities in 10 different malaria-endemic regions. This article discusses current knowledge of antimalarial drug resistance, including activities of ICEMR groups to assess and characterize resistance in different regions.

Antimalarial Drugs

Antimalarial drugs act principally to eliminate the erythrocytic stages of malaria parasites that are responsible for human illness. Drug regimens for treatment of the two most prevalent malaria parasites, P. falciparum and P. vivax, are different. With frequent resistance to older drugs, artemisinin-based combination therapy (ACT) is now recommended for the treatment of uncomplicated falciparum malaria in nearly all areas. Chloroquine plus primaquine remains the first-line regimen for radical cure of vivax malaria in most regions. ACT consists of a potent artemisinin component, which rapidly clears most parasites, plus a longer acting partner drug, which eliminates remaining parasites and limits selection of artemisinin resistance. The ACTs recommended by the World Health Organization (WHO) are artemether/lumefantrine, artesunate/amodiaquine, artesunate/mefloquine, dihydroartemisinin/piperaquine, artesunate/pyronaridine, and artesunate/sulfadoxine–pyrimethamine. ACTs are also effective against erythrocytic stages of non-falciparum malaria parasites. Multiple drugs are used to prevent malaria. Recommendations for travelers from nonendemic to endemic areas generally advocate use of atovaquone/proguanil (Malarone), mefloquine, or doxycycline in chemoprophylactic regimens. In Africa, intermittent preventive therapy is advocated in some high-risk populations, including sulfadoxine/pyrimethamine during pregnancy and amodiaquine/sulfadoxine–pyrimethamine as seasonal malaria chemoprophylaxis in the Sahel subregion, where there is little resistance to these drugs.

Available antimalarial drugs can be divided into multiple classes (Table 1). The 4-aminoquinoline, chloroquine, was the gold standard for the treatment of uncomplicated malaria for many years, but it is no longer appropriate for the treatment of falciparum malaria in nearly all areas because of drug resistance. In Indonesia, increased resistance to chloroquine in P. vivax prompted a policy change to ACTs for vivax malaria. Amodiaquine appears to be subject to the same resistance mechanisms as chloroquine, but due to improved potency it provides adequate efficacy against many chloroquine-resistant parasites, and it is a component of the widely used ACT artesunate/amodiaquine. A third 4-aminoquinoline, piperaquine, was widely used to treat and prevent malaria in China a few decades ago, but it then fell into disfavor because of increasing drug resistance. More recently, piperaquine has become a component of another ACT, dihydroartemisinin/piperaquine. The 8-aminoquinoline, primaquine, has some activity against erythrocytic parasites, but it is used principally to eliminate parasite liver stages, including the exoerythrocytic forms that precede erythrocytic infection in all species and the hypnozoites that cause latent infections with P. vivax and Plasmodium ovale. Primaquine also acts against gametocytes, thereby lowering transmission of parasites to mosquito vectors. Quinine is an arylamino alcohol that is the oldest antimalarial drug, used as cinchona bark since the 1600s and in its pure form since 1820. Quinine is quite hard to tolerate, and its use is best limited to the treatment of severe malaria. Important malaria-related drugs are mefloquine and lumefantrine, components of the ACTs artesunate/mefloquine and artemether/lumefantrine.

Antifolates target parasite dihydrofolate reductase (DHFR) and dihydropteroate synthase (DHPS). Sulfadoxine/pyrimethamine has the distinct advantage of single-dose therapy, but its treatment efficacy is seriously limited by drug resistance. The naphthoquinone atovaquone acts against the mitochondrial cytochrome bc1 complex. Combined with the DHFR inhibitor proguanil as Malarone, it offers effective albeit expensive therapy and chemoprophylaxis for falciparum malaria, although it is noteworthy that the synergy of this combination appears to be independent of inhibition of folate synthesis. Several antibiotics that are prokaryotic protein synthesis inhibitors have antimalarial activity because of action against the protein synthesis machinery of the apicoplast organelle. Doxycycline is used for chemoprophylaxis against malaria, and doxycycline or clindamycin are combined with quinine to treat falciparum malaria.

The most important new class of antimalarials is the artemisinins, which were developed from a natural product remedy in China. Artemisinin is a potent antimalarial, but the derivatives artesunate, artemether, and dihydroartemisinin are most widely used as components of ACT regimens. Indeed, the use of artemisinins outside combination regimens is strongly discouraged by the WHO because of fear of selecting for resistance to this important class of drugs. Artemisinins are highly effective against acute malaria, but short acting, so combination with longer-acting drugs in ACTs allows short (3-day) courses of treatment that protect against the selection of resistance to the artemisinin component. Because of its rapid action, intravenous artesunate is also the new gold standard for the treatment of severe falciparum malaria, with documented survival advantages compared with intravenous quinine.,

Antimalarial Drug Resistance

The efficacies of many antimalarial drugs are limited by drug resistance, and recent evidence suggests that parasites are becoming resistant to the newest agents. However, the extent of resistance varies, such that in many cases drugs with resistance concerns are nonetheless offering good effectiveness for the treatment and control of malaria. Resistance has been described for nearly all available drugs, as is discussed below. For many drugs the extent of resistance is uncertain and mechanisms of resistance are unknown, and thus the opportunity to glean data from the 10 ICEMR surveillance sites is highly valuable. Resistance can be assessed by clinical trials comparing antimalarial efficacies of different agents, ex vivo/in vitro assessment of sensitivities of cultured P. falciparum, evaluation of genetic polymorphisms associated with resistance, or by assessing the selective pressure of antimalarial treatment on subsequent infections. Studies considering all of these factors have shed light on the extent of resistance and on mechanisms of resistance.

Resistance mediated by transporter mutations.

The P. falciparum genome encodes multiple predicted transporters. Polymorphisms in transport proteins can mediate resistance to many agents active against cancer and infectious diseases via enhancing efflux of the drugs from cells. It appears that a number of plasmodial proteins transport different drugs and that polymorphisms in these proteins may impact on drug sensitivity.

pfmdr1.

Polymorphisms in the P. falciparum multidrug resistance-1 (pfmdr1) gene, which encodes the P-glycoprotein homolog, impact on sensitivity to multiple antimalarial drugs.– In humans, P-glycoprotein polymorphisms are associated with resistance to cancer drugs. In P. falciparum, the function of the pfmdr1 product is unknown, but the protein localizes to the membrane of the food vacuole, the site of action of a number of drugs, suggesting that it is a drug transporter. Data on associations between pfmdr1 polymorphisms and drug sensitivity are complex, but overall suggest that changes in pfmdr1 sequence or copy number alter transport of multiple drugs in or out of the parasite food vacuole, with individual polymorphisms leading to opposite effects on different drugs. Increased copy number of pfmdr1, which is prevalent in southeast Asia, has been associated with extensive use of mefloquine. Experimental evidence indicates that pfmdr1 amplification also leads to decreased sensitivity to quinine, lumefantrine, and artemisinin. Mutations at pfmdr1 N86Y and D1246Y (for this and other P. falciparum genes, wild type sequence is based on the 3D7 reference strain), which are common in Africa, have been linked to decreased sensitivity to chloroquine and amodiaquine, but increased sensitivity to lumefantrine, mefloquine, and artemisinins.– Other polymorphisms primarily seen outside Africa (including 1034C and 1042D) are associated with altered sensitivity to lumefantrine, mefloquine, and artemisinins.,,– Considering infections that emerge soon after prior therapy, amodiaquine-containing regimens selected for the 86Y and 1246Y mutant alleles– and for parasites with decreased in vitro sensitivity to the active metabolite monodesethylamodiaquine in subsequent infections. The selective pressure of the related aminoquinoline piperaquine seems less marked than that of amodiaquine, but prior use of the drug also selected for the pfmdr1 86Y and 1246Y mutations., In contrast, therapy with artemether/lumefantrine selected for the N86 and D1246 wild type alleles in subsequent infections within 60 days of prior therapy.,,,– Importantly, impacts of pfmdr1 polymorphisms on drug sensitivity are modest, correlations between particular polymorphisms and treatment efficacy have not been seen, and the ACTs artesunate/amodiaquine and artemether/lumefantrine remain highly efficacious for the treatment of uncomplicated falciparum malaria in Africa., However, as seen for chloroquine and amodiaquine, pfmdr1 polymorphisms may contribute, with additional polymorphisms, to resistance to increasingly used components of ACTs.

pfcrt.

Soon after the identification of pfmdr1, it became clear that polymorphisms in this gene are not the primary mediators of chloroquine resistance. Subsequently, analysis of progeny of a genetic cross between chloroquine sensitive and resistant strains led to the identification of pfcrt, which encodes a food vacuole membrane protein that is predicted to be a member of the drug/metabolite transporter superfamily., The function of pfcrt is unknown, but apparently essential, as disruption of the gene has not been possible. pfcrt is highly polymorphic, but one single nucleotide polymorphism (SNP), K76T, is the primary mediator of chloroquine resistance., The 76T mutation appears to act principally by increasing the export of chloroquine from the food vacuole, but the mechanism of pfcrt 76T-mediated chloroquine resistance is incompletely understood. Other pfcrt SNPs always accompany 76T in field isolates, and these likely encode compensatory mutations that allow parasites containing 76T to maintain adequate fitness; some other SNPs may also contribute directly to the drug resistant phenotype. The 76T mutation also mediates decreased sensitivity to monodesethylamodiaquine, and studies with genetically modified parasites have shown it to mediate increased susceptibility to mefloquine and artemisinins, suggesting the same reciprocal relationship between sensitivities to aminoquinolines and other drugs as described for certain pfmdr1 polymorphisms.

Pfmrp1.

Plasmodium falciparum multidrug resistance protein-1 (Pfmrp1) is a member of the ABC transporter superfamily. In studies of culture-adapted P. falciparum, SNPs in pfmrp1 were linked to decreased sensitivity to chloroquine and quinine. Two SNPs that appear to be common in African parasites, I876V and K1466R, were selected by prior treatment with artemether/lumefantrine and sulfadoxine/pyrimethamine, respectively, although these SNPs were not associated with altered drug sensitivity in African isolates. pfmrp1 mutations appear to differ between continents; some SNPs in northeast Myanmar isolates were associated with reduced susceptibilities to chloroquine, mefloquine, pyronaridine, and lumefantrine. Disruption of the pfmrp1 gene yielded parasites with diminished growth and increased sensitivity to chloroquine and other drugs, suggesting a role for this protein in the efflux of antimalarial drugs from the parasite and in parasite fitness.

Sodium transporters.

Quantitative trait locus analysis identified three genes predicted to play roles in the responsiveness of P. falciparum to quinine, pfcrt, pfmdr1, and pfnhe1, which encode a putative sodium–hydrogen exchanger and are highly polymorphic. pfatp4 encodes a P. falciparum plasma membrane protein that appears to be a sodium efflux pump. Recent studies have shown that three different classes of potent antimalarial compounds, spiroindolones, pyrazoleamides, and dihydroisoquinolones, all target pfatp4.– Mutations in pfatp4 have been linked to altered sensitivity to these candidate antimalarials.– In addition, screening of the “Malaria Box” chemical library identified 28 compounds of diverse chemotypes that affected parasite Na+ and pH regulation in a manner consistent with PfATP4 inhibition. A recent clinical trial of the spiroindolone KAE609 demonstrated excellent efficacy against falciparum and vivax malaria.

Resistance to quinine.

Resistance to quinine, the oldest antimalarial drug, was reported first in Brazil and later in southeast Asia., Quinine resistance is associated with polymorphisms in several transporters. As stated earlier, SNPs in pfmdr1, pfcrt, and pfmrp1 are linked to decreased sensitivity to quinine. In addition, pfmdr1 gene amplification can also lead to quinine resistance. Recent studies evaluating associations between polymorphisms in a pfnhe1 microsatellite, in vitro parasite sensitivity, and clinical responses to various drugs have been inconsistent, but these polymorphisms appear to have a modest impact on sensitivity of parasites to quinine, and possibly other drugs.– In vitro allelic exchange to reduce the expression of pfnhe1 by ~50% led to a 30% increase in quinine sensitivity in some, but not other parasite strains.

Resistance to antifolates.

The parasite-specific antimetabolite, pyrimethamine, is usually discussed in combination with its partner drug sulfadoxine (known as SP or Fansidar). Pyrimethamine was first used as an individual drug, but resistance was seen within a year in both P. vivax and P. falciparum., The combination of sulfa drugs and pyrimethamine proved to be potent in the laboratory, as well as in the field against chloroquine-resistant uncomplicated malaria but, again, resistance appeared rapidly in the Asia Pacific regions in the late 1970s, as well as in South America.–

Molecular genetic studies attribute pyrimethamine and sulfa resistance to mutations in the genes coding for the target enzymes DHFR and DHPS., These markers have been useful for tracking sulfadoxine/pyrimethamine resistance across the globe, and show particular promise with new multiplex strategies. In the 1990s, sulfadoxine/pyrimethamine found increasing use in Africa to treat widespread chloroquine-resistant malaria, before sulfadoxine/pyrimethamine resistance followed. Molecular epidemiology studies utilizing DNA microsatellite sequences flanking the dhfr gene point to transfer of pyrimethamine resistance from Asia to Africa, possibly from a single ancestor and possibly before sulfadoxine/pyrimethamine use even began in Africa. In contrast, resistance to sulfa partners through dhps mutations seemed to occur through de novo events both in sub-Saharan Africa and in Asia., Sulfadoxine/pyrimethamine is no longer recommended as a first-line drug for the treatment of P. falciparum. However, it continues to be used in ACT combinations in most parts of India, for intermittent preventive therapy in pregnant women in Africa, and for seasonal malaria chemoprevention in children in the sub-Sahel of Africa, although widespread resistance limits these interventions., Perhaps not surprisingly, sulfadoxine/pyrimethamine resistance in P. vivax appeared in Asia and the Pacific Islands, where P. falciparum and P. vivax coexist.–

Resistance to artemisinin family drugs.

Since artemisinins play an indispensable role in current malaria therapies, artemisinin resistance has received wide recent attention. In the Cambodia–Thailand border region of southeast Asia, an epicenter of antimalarial drug resistance, declining efficacy of the artesunate/mefloquine combination was noted, and clinical resistance to artesunate, manifested as delayed clearance of parasitemia after therapy, but not generally as full-blown treatment failure, was documented in 2008.– The delayed parasite clearance phenotype does not correspond to increased artemisinin half maximal inhibitory concentration (IC50) values, as determined by standard in vitro assays, but does correspond to decreased susceptibility assessed 72 hours after a pulse of dihydroartemisinin using the new ring-stage survival assay., Attempts to select resistance to artemisinins in vitro using constant or pulsed drug selection pressure initially identified pfmdr1 amplification and increased antioxidant levels in selected parasites., In field-based studies, genome-wide association studies identified regions on chromosome 13 linked to delayed parasite clearance., Using a combined resistance selection and genomic approach, Ariey and others identified mutations in the propeller domain of the P. falciparum kelch (K13) gene (PF3D7_1343700) associated with delayed parasite clearance after artemisinin therapy in southeast Asia. Very recently, using clinical and molecular data, the extent of artemisinin resistance has been delineated, with delayed clearance and K13 mutations common in parts of Cambodia, Thailand, Myanmar, and Vietnam, but not in other areas of Asia or Africa.– Other reports from Cambodia have shown recrudescent infections after treatment with dihydroartemisinin/piperaquine, raising the concern that resistance to artemisinin partner drugs has been facilitated by the spread of artemisinin resistance. In African parasites, although K13 gene polymorphisms are common, including some mutations in the propeller domain, the specific mutations associated with artemisinin resistance in southeast Asia have not been seen.– Parasites with introduced K13 mutations showed enhanced survival after a dihydroartemisinin pulse, confirming the role of these mutations in resistance. The transcriptomes of resistant parasites showed increased expression of unfolded protein response pathways and prolonged ring-stage development, offering insights into the mechanism of artemisinin resistance.

Resistance to Malarone.

Atovaquone is a potent inhibitor of electron transport, and studies identified the target of this drug as the critical quinone-binding sites of cytochrome b., When the drug is used alone, resistance develops rapidly and recrudescence after therapy is common. Resistance is conferred by single-point mutations in the cytochrome b (Pfcytb) gene. Pfcytb mutations 268S and 268N were associated with Malarone treatment failure., However, treatment failure has also been reported in the absence of these mutations.–

ICEMR Data Cocncerning Antimalarial Drug Resistance

Data from ICEMR sites, collected both before and during enactment of the ICEMR programs, offer insight into global drug resistance trends. Research activities at different ICEMR sites entail clinical studies, ex vivo/in vitro assays, and molecular studies (Table 2 ).

Clinical observations.

Clinical trials in west Africa, Uganda, south Asia, and Papua New Guinea (PNG) have generally shown excellent antimalarial efficacy for the ACTs artemether/lumefantrine, artesunate/amodiaquine, and dihydroartemisinin/piperaquine (Supplemental Table 1). In high transmission settings, new infections after ACT therapy may be common, but true recrudescences after treatment have been very uncommon. In India, particularly in the northeast along the Myanmar border, where artesunate/sulfadoxine–pyrimethamine combinations are being discontinued, there have been excellent responses to artesunate/mefloquine, artesunate/amodiaquine, and dihydroartemisinin/piperaquine (Supplemental Table 1). Even in low-endemicity areas at the China–Myanmar border, where artemisinins have the longest history of use, excellent efficacy of the ACTs for treatment of falciparum malaria has been seen. On the other hand, an effectiveness study in Uganda showed a failure rate of 31% after treatment with quinine. In regions of P. falciparum/P. vivax co-endemicity, P. falciparum typically shows rapid responses to control efforts, whereas P. vivax prevalence subsides more slowly. In northeast Myanmar, follow-ups of P. vivax cases after chloroquine/primaquine treatment indicated an increase in cases with recurrent parasitemia within 28 days compared with a prior report, suggesting the emergence of chloroquine resistance. Ongoing clinical efficacy studies at the ICEMR sentinel sites will be important to offer a longitudinal appreciation of drug efficacy and provide a scientific basis to guide local drug policy (Table 3 ).

Ex vivo and in vitro studies.

Studies on ex vivo (parasites studied immediately after collection from infected patients) or in vitro (parasites studied after culture adaptation) antimalarial drug sensitivity of P. falciparum have been conducted by ICEMR groups from west Africa, Uganda, south Asia, southeast Asia, and PNG (Supplemental Table 2). Ex vivo studies have the advantage of testing samples directly from patients without potential selection biases due to constraints of culturing and cryopreservation. However, the results may be confounded by the presence of multiple clones of parasites that differ in sensitivities to the test drugs. In vitro assays performed with culture-adapted parasite clones allow assays to include multiple biological replications and provide better opportunities for subsequent genetic analysis. In general, these studies have shown that African parasites have varied sensitivities to chloroquine and amodiaquine, and good sensitivities to dihydroartemisinin, the active metabolite of all artemisinin derivatives, and to the ACT partner drugs lumefantrine, mefloquine, and piperaquine.,, In Uganda, increased deployment of artemether/lumefantrine was linked to some decrease in in vitro susceptibility to lumefantrine. In Thailand, reduced lumefantrine susceptibility might have resulted from extensive use of mefloquine, another aminoalcohol. Clinical and in vitro resistance to quinine has been seen in southeast Asia, but not consistently in Africa. In Senegal and Uganda, for example, P. falciparum parasites appeared to be susceptible to quinine in vitro., In comparison, data from southeast Asia showed a mean IC50 greater than 500 nM. Similar to African parasites, southeast Asian isolates were generally sensitive to artemisinins and the ACT partner drugs lumefantrine and mefloquine. Yet, longitudinal studies revealed gradual decrease of susceptibility to piperaquine, and a high correlation between chloroquine and piperaquine IC50 values. However, only limited results are available, and considerations of ex vivo/in vitro results is complicated by varied methodologies used by different groups, difficulties of interpreting results for polyclonal infections, and uncertain correlations between in vitro findings and clinical efficacy. Commonly used ex vivo/in vitro assays measure the parasite histidine-rich protein-2 by enzyme-linked immunosorbent assay, replication of parasite DNA by isotope incorporation, or use of a fluorescent dye such as SBYR Green I., To enhance comparisons among sites, ex vivo/in vitro assays should consider the inclusion of a standard laboratory strain (such as 3D7) as an internal control. Further, the new ring-stage survival assay should be adopted in multiple ICEMR sites to monitor the emergence and spread of artemisinin resistance. Some ICEMRs also have prevalent transmission of vivax malaria. Ex vivo drug assays for P. vivax are also being conducted (Table 3), but the assays are constrained by difficulties of P. vivax culture and the appreciation that assays for certain drugs (e.g., chloroquine) require a high proportion of parasites at the ring stage and a high parasitemia.

Genotyping drug resistance–mediating polymorphisms.

Studies of genetic polymorphisms associated with drug resistance are technically much simpler than in vitro studies of parasite sensitivity, and so results are more widely available. Studies of P. falciparum genetic polymorphisms have been conducted by ICEMR groups from west Africa, southern Africa, Uganda, south Asia, southeast Asia, and PNG (Supplemental Tables 3 and 4). As has already been well documented in past studies, the prevalence of a number of polymorphisms that impact on drug sensitivity varies greatly around the world. Also of interest are changes in polymorphism prevalence over time. In Uganda, parasites have demonstrated marked changes in the prevalence of some key polymorphisms over the last decade, coincident with changes in treatment practices for malaria from chloroquine to chloroquine/sulfadoxine–pyrimethamine to artemether/lumefantrine. Most notably, the prevalence of three wild type alleles, pfcrt K76, pfmdr1 N86, and pfmdr1 D1246, has all increased markedly in recent years and this increase was greater in children treated with artemether/lumefantrine for all episodes of malaria than in those treated with dihydroartemisinin/piperaquine. This is in sharp contrast to the P. falciparum parasites at the China–Myanmar border area, where pfcrt 76T and 220S remained almost fixed in a recent study. Recently, the Uganda ICEMR group showed that therapy with artemether/lumefantrine selects for the wild-type polymorphisms associated with decreased lumefantrine efficacy and, in ex vivo studies, for parasites with diminished lumefantrine sensitivity. Importantly, despite these changes, sensitivity to lumefantrine remains quite good, and artemether/lumefantrine treatment efficacy is excellent. However, recent unpublished trials (Yeka and others, unpublished data) showed that in 2013–2014 artemether/lumefantrine was less efficacious than artesunate/amodiaquine at three different sites in Uganda, a change in relative treatment efficacy compared with older findings., These results suggest that recent changes in treatment practices have led to changes in P. falciparum in Uganda that have mediated decreased antimalarial efficacy of artemether/lumefantrine, the first-line antimalarial drug in the country. An urgent issue that the ICEMRs are addressing is monitoring of the emergence and/or spread of artemisinin resistance. The identification of the K13 gene as a molecular marker for artemisinin resistance will facilitate large-scale surveillance.

Resistance and Fitness

One of the challenges to studying the interplay between parasite drug resistance and fitness is the lack of a direct measure for fitness. Comparison of relative growth rates in vitro or ex vivo is the commonly used approach, although growth rates may not represent relative fitness in the natural host. Assessment of parasite survival in the field provides an improved measure, although analyses are challenging. It has long been observed that P. falciparum genetic mutations that confer drug resistance are associated with altered biological fitness of the parasite.– However, various investigators have reported both increased– and decreased– fitness in resistant parasites. The latter would seem intuitive from initial low prevalence of innate resistance observed in the field for some antimalarials such as mefloquine and atovaquone, presumably owing to the mutant parasites being outcompeted by the wild type before start of drug use.,, More compelling evidence for a fitness cost of resistance has been documented by reemergence of sensitive parasites, virtually replacing highly prevalent resistant strains, following withdrawal of drug (chloroquine or sulfadoxine–pyrimethamine) pressure in the population.– The rub is that in other areas under similar conditions, reemergence has occurred much more slowly or not at all.– Similarly cogent evidence of a fitness cost to resistance has been demonstrated by decreased prevalence of resistant parasites after the dry season in west Africa, when there is relatively little drug selection pressure.– Again, other studies have had contrasting results.,

Global data from the ICEMRs afford an opportunity for a concerted evidence base on associations between resistance and fitness, and potentially on the de facto risk factors for the emergence or suppression of drug resistance in the field. So far drug selection pressure, herd immunity,, and ecological differences,, have been shown to impinge on relative fitness of drug-resistant and drug-sensitive parasites in the wild. Although combination of antimalarial compounds with opposing resistance mechanisms have been used to suppress the emergence of drug resistance in laboratory isolates, opposite resistance selection has also been observed in the field between 4-aminoquinolines (chloroquine, amodiaquine) and artemisinins.,– Data from Uganda showed significantly lower prevalence of symptoms among children infected with parasites containing chloroquine resistance mutations compared with those infected with wild-type parasites, consistent with greater virulence for wild-type parasites. Field data from the southern Africa ICEMR suggest a role for the vector in selecting drug resistance polymorphisms, with significant differences in prevalence of SNPs that mediate resistance to aminoquinolines and antifolates between parasites infecting mosquitoes and people., So far, Anopheles arabiensis, and more recently Anopheles funestus (Matsena and others, unpublished data) have both been shown to exert selection on drug resistance polymorphisms. This would seem to explain the role of ecology in governing resistance,, which can differ between different regions of the same country. The unexpected link between vector control and prevalence of drug-resistant malaria parasites in some areas– but not others also seems consistent with vector selection. More detailed data from multiple ICEMRs will be instrumental in improving our understanding of the interplay between drug resistance and fitness, and hopefully the development of more effective strategies for the containment of drug-resistant malaria.

Conclusion and Future Studies

In an evolutionary arms race between malaria parasites and a series of therapeutic interventions, the parasites have consistently been able to develop resistance to each new class of drugs. The emergence of parasites resistant to artemisinins in southeast Asia and altered sensitivities to artemisinin partner drugs pose great threats to efforts to control and, eventually, eradicate malaria. Specifically, previous failures of the ACTs artesunate/mefloquine and artesunate/amodiaquine have recently been followed by frequent failures of dihydroartemisinin/piperaquine in parts of Cambodia, and decreasing sensitivity to lumefantrine may further threaten artemether/lumefantrine. It is thus of high priority to continue surveillance of ACT efficacy, the ex vivo and in vitro activities of ACT components, and molecular markers that may mediate resistance to these drugs. For artemisinins, mutations in the K13 gene offer markers for the delayed parasite clearance phenotype that is now common in parts of southeast Asia. Mutations in the putative drug transporters pfmdr1 and pfcrt mediate altered sensitivity to multiple artemisinin partner drugs, including amodiaquine, mefloquine, lumefantrine, and piperaquine, although different drugs are impacted in opposite directions. Additional parasite polymorphisms are likely important in drug responsiveness, and an improved understanding of the roles of these polymorphisms is an important goal. Additional studies on the influence of drug resistance on parasite fitness may enable the identification of optimal dosing strategies, including, possibly, rotating of regimens. Further, understanding how mosquitoes mediate the spread of drug resistance and use of evolution-proof mosquito control measures may help to deter resistance spread, enabling the regional elimination and eventual eradication of malaria.

References 178

  1. 1.

    Murray CJ, Rosenfeld LC, Lim SS, Andrews KG, Foreman KJ, Haring D, Fullman N, Naghavi M, Lozano R, Lopez AD. Global malaria mortality between 1980 and 2010: a systematic analysis. Lancet. 2012;379:413-431.

    Not in our library10.1016/s0140-6736(12)60034-8

  2. 2.
    Multidrug-Resistant Plasmodium vivax Associated with Severe and Fatal Malaria: A Prospective Study in Papua, Indonesia · 2008
    No notice
  3. 3.
    Combining indoor residual spraying and insecticide-treated nets for malaria control in Africa: a review of possible outcomes and an outline of suggestions for the future · 2011
    No notice
  4. 4.

    Olotu A, Fegan G, Wambua J, Nyangweso G, Awuondo KO, Leach A, Lievens M, Leboulleux D, Njuguna P, Peshu N, Marsh K, Bejon P. Four-year efficacy of RTS,S/AS01E and its interaction with malaria exposure. N Engl J Med. 2013;368:1111-1120.

    Not in our library10.1056/nejmoa1207564

  5. 5.

    Gosling RD, Okell L, Mosha J, Chandramohan D. The role of antimalarial treatment in the elimination of malaria. Clin Microbiol Infect. 2011;17:1617-1623.

    Not in our library10.1111/j.1469-0691.2011.03660.x

  6. 6.

    Greenwood B. Anti-malarial drugs and the prevention of malaria in the population of malaria endemic areas. Malar J. 2010;9(Suppl 3):S2.

    Not in our library10.1186/1475-2875-9-s3-s2

  7. 7.

    Nosten F, White NJ. Artemisinin-based combination treatment of falciparum malaria. Am J Trop Med Hyg. 2007;77:181-192.

    Not in our library

  8. 8.

    Schlagenhauf P, Petersen E. Malaria chemoprophylaxis: strategies for risk groups. Clin Microbiol Rev. 2008;21:466-472.

    Not in our library10.1128/cmr.00059-07

  9. 9.

    Baird JK. Resistance to chloroquine unhinges vivax malaria therapeutics. Antimicrob Agents Chemother. 2011;55:1827-1830.

    Not in our library10.1128/aac.01296-10

  10. 10.

    Davis TM, Hung TY, Sim IK, Karunajeewa HA, Ilett KF. Piperaquine: a resurgent antimalarial drug. Drugs. 2005;65:75-87.

    Not in our library10.2165/00003495-200565010-00004

  11. 11.

    Meshnick SR, Dobson MJ, Rosenthal PJ. The history of antimalarial drugs. Antimalarial Chemotherapy: Mechanisms of Action, Resistance, and New Directions in Drug Discovery. 2001;15-25.

    Not in our library

  12. 12.

    Dahl EL, Rosenthal PJ. Apicoplast translation, transcription and genome replication: targets for antimalarial antibiotics. Trends Parasitol. 2008;24:279-284.

    Not in our library10.1016/j.pt.2008.03.007

  13. 13.

    Cui L, Su XZ. Discovery, mechanisms of action and combination therapy of artemisinin. Expert Rev Anti Infect Ther. 2009;7:999-1013.

    Not in our library10.1586/eri.09.68

  14. 14.

    Dondorp A, Nosten F, Stepniewska K, Day N, White N. Artesunate versus quinine for treatment of severe falciparum malaria: a randomised trial. Lancet. 2005;366:717-725.

    Not in our library10.1016/s0140-6736(05)67176-0

  15. 15.

    Dondorp AM, Fanello CI, Hendriksen IC, Gomes E, Seni A, Chhaganlal KD, Bojang K, Olaosebikan R, Anunobi N, Maitland K, Kivaya E, Agbenyega T, Nguah SB, Evans J, Gesase S, Kahabuka C, Mtove G, Nadjm B, Deen J, Mwanga-Amumpaire J, Nansumba M, Karema C, Umulisa N, Uwimana A, Mokuolu OA, Adedoyin OT, Johnson WB, Tshefu AK, Onyamboko MA, Sakulthaew T, Ngum WP, Silamut K, Stepniewska K, Woodrow CJ, Bethell D, Wills B, Oneko M, Peto TE, von Seidlein L, Day NP, White NJ. Artesunate versus quinine in the treatment of severe falciparum malaria in African children (AQUAMAT): an open-label, randomised trial. Lancet. 2010;376:1647-1657.

    Not in our library10.1016/s0140-6736(10)61924-1

  16. 16.

    Mu J, Ferdig MT, Feng X, Joy DA, Duan J, Furuya T, Subramanian G, Aravind L, Cooper RA, Wootton JC, Xiong M, Su XZ. Multiple transporters associated with malaria parasite responses to chloroquine and quinine. Mol Microbiol. 2003;49:977-989.

    Not in our library10.1046/j.1365-2958.2003.03627.x

  17. 17.

    Borges-Walmsley MI, McKeegan KS, Walmsley AR. Structure and function of efflux pumps that confer resistance to drugs. Biochem J. 2003;376:313-338.

    Not in our library10.1042/bj20020957

  18. 18.
    A systematic review and meta-analysis of evidence for correlation between molecular markers of parasite resistance and treatment outcome in falciparum malaria · 2009
    No notice
  19. 19.

    Foote SJ, Kyle DE, Martin RK, Oduola AM, Forsyth K, Kemp DJ, Cowman AF. Several alleles of the multidrug-resistance gene are closely linked to chloroquine resistance in Plasmodium falciparum. Nature. 1990;345:255-258.

    Not in our library10.1038/345255a0

  20. 20.

    Sanchez CP, Dave A, Stein WD, Lanzer M. Transporters as mediators of drug resistance in Plasmodium falciparum. Int J Parasitol. 2010;40:1109-1118.

    Not in our library10.1016/j.ijpara.2010.04.001

  21. 21.

    Valderramos SG, Fidock DA. Transporters involved in resistance to antimalarial drugs. Trends Pharmacol Sci. 2006;27:594-601.

    Not in our library10.1016/j.tips.2006.09.005

  22. 22.

    Sharom FJ. The P-glycoprotein multidrug transporter. Essays Biochem. 2011;50:161-178.

    Not in our library10.1042/bse0500161

  23. 23.

    Cowman AF, Karcz S, Galatis D, Culvenor JG. A P-glycoprotein homologue of Plasmodium falciparum is localized on the digestive vacuole. J Cell Biol. 1991;113:1033-1042.

    Not in our library10.1083/jcb.113.5.1033

  24. 24.

    Koenderink JB, Kavishe RA, Rijpma SR, Russel FG. The ABCs of multidrug resistance in malaria. Trends Parasitol. 2010;26:440-446.

    Not in our library10.1016/j.pt.2010.05.002

  25. 25.

    Price RN, Uhlemann AC, Brockman A, McGready R, Ashley E, Phaipun L, Patel R, Laing K, Looareesuwan S, White NJ, Nosten F, Krishna S. Mefloquine resistance in Plasmodium falciparum and increased pfmdr1 gene copy number. Lancet. 2004;364:438-447.

    Not in our library10.1016/s0140-6736(04)16767-6

  26. 26.

    Sidhu AB, Uhlemann AC, Valderramos SG, Valderramos JC, Krishna S, Fidock DA. Decreasing pfmdr1 copy number in Plasmodium falciparum malaria heightens susceptibility to mefloquine, lumefantrine, halofantrine, quinine, and artemisinin. J Infect Dis. 2006;194:528-535.

    Not in our library10.1086/507115

  27. 27.

    Duraisingh MT, Jones P, Sambou I, von Seidlein L, Pinder M, Warhurst DC. The tyrosine-86 allele of the pfmdr1 gene of Plasmodium falciparum is associated with increased sensitivity to the anti-malarials mefloquine and artemisinin. Mol Biochem Parasitol. 2000;108:13-23.

    Not in our library10.1016/s0166-6851(00)00201-2

  28. 28.

    Duraisingh MT, Roper C, Walliker D, Warhurst DC. Increased sensitivity to the antimalarials mefloquine and artemisinin is conferred by mutations in the pfmdr1 gene of Plasmodium falciparum. Mol Microbiol. 2000;36:955-961.

    Not in our library10.1046/j.1365-2958.2000.01914.x

  29. 29.

    Reed MB, Saliba KJ, Caruana SR, Kirk K, Cowman AF. Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature. 2000;403:906-909.

    Not in our library10.1038/35002615

  30. 30.

    Mwai L, Kiara SM, Abdirahman A, Pole L, Rippert A, Diriye A, Bull P, Marsh K, Borrmann S, Nzila A. In vitro activity of piperaquine, lumefantrine and dihydroartemisinin in Kenyan Plasmodium falciparum isolates and polymorphisms in Pfcrt and Pfmdr1. Antimicrob Agents Chemother. 2009;55:5069-5073.

    Not in our library10.1128/aac.00638-09

  31. 31.

    Tumwebaze P, Conrad MD, Walakira A, LeClair N, Byaruhanga O, Nakazibwe C, Kozak B, Bloome J, Okiring J, Kakuru A, Bigira V, Kapisi J, Legac J, Gut J, Cooper RA, Kamya MR, Havlir DV, Dorsey G, Greenhouse B, Nsobya SL, Rosenthal PJ. Impact of antimalarial treatment and chemoprevention on the drug sensitivity of malaria parasites isolated from Ugandan children. Antimicrob Agents Chemother. 2015;59:3018-3030.

    Not in our library10.1128/aac.05141-14

  32. 32.

    Pickard AL, Wongsrichanalai C, Purfield A, Kamwendo D, Emery K, Zalewski C, Kawamoto F, Miller RS, Meshnick SR. Resistance to antimalarials in southeast Asia and genetic polymorphisms in pfmdr1. Antimicrob Agents Chemother. 2003;47:2418-2423.

    Not in our library10.1128/aac.47.8.2418-2423.2003

  33. 33.

    Sidhu AB, Valderramos SG, Fidock DA. pfmdr1 mutations contribute to quinine resistance and enhance mefloquine and artemisinin sensitivity in Plasmodium falciparum. Mol Microbiol. 2005;57:913-926.

    Not in our library10.1111/j.1365-2958.2005.04729.x

  34. 34.

    Veiga MI, Ferreira PE, Jornhagen L, Malmberg M, Kone A, Schmidt BA, Petzold M, Bjorkman A, Nosten F, Gil JP. Novel polymorphisms in Plasmodium falciparum ABC transporter genes are associated with major ACT antimalarial drug resistance. PLoS One. 2011;6:e20212.

    Not in our library10.1371/journal.pone.0020212

  35. 35.

    Humphreys GS, Merinopoulos I, Ahmed J, Whitty CJ, Mutabingwa TK, Sutherland CJ, Hallett RL. Amodiaquine and artemether-lumefantrine select distinct alleles of the Plasmodium falciparum mdr1 gene in Tanzanian children treated for uncomplicated malaria. Antimicrob Agents Chemother. 2007;51:991-997.

    Not in our library10.1128/aac.00875-06

  36. 36.

    Zongo I, Dorsey G, Rouamba N, Tinto H, Dokomajilar C, Guiguemde RT, Rosenthal PJ, Ouedraogo JB. Artemether-lumefantrine versus amodiaquine plus sulfadoxine-pyrimethamine for uncomplicated falciparum malaria in Burkina Faso: a randomised non-inferiority trial. Lancet. 2007;369:491-498.

    Not in our library10.1016/s0140-6736(07)60236-0

  37. 37.

    Nsobya SL, Dokomajilar C, Joloba M, Dorsey G, Rosenthal PJ. Resistance-mediating Plasmodium falciparum pfcrt and pfmdr1 alleles after treatment with artesunate-amodiaquine in Uganda. Antimicrob Agents Chemother. 2007;51:3023-3025.

    Not in our library10.1128/aac.00012-07

  38. 38.

    Nawaz F, Nsobya SL, Kiggundu M, Joloba M, Rosenthal PJ. Selection of parasites with diminished drug susceptibility by amodiaquine-containing antimalarial regimens in Uganda. J Infect Dis. 2009;200:1650-1657.

    Not in our library10.1086/647988

  39. 39.

    Conrad MD, LeClair N, Arinaitwe E, Wanzira H, Kakuru A, Bigira V, Muhindo M, Kamya MR, Tappero JW, Greenhouse B, Dorsey G, Rosenthal PJ. Comparative impacts over 5 years of artemisinin-based combination therapies on Plasmodium falciparum polymorphisms that modulate drug sensitivity in Ugandan children. J Infect Dis. 2014;210:344-353.

    Not in our library10.1093/infdis/jiu141

  40. 40.

    Sisowath C, Stromberg J, Martensson A, Msellem M, Obondo C, Bjorkman A, Gil JP. In vivo selection of Plasmodium falciparum pfmdr1 86N coding alleles by artemether-lumefantrine (Coartem). J Infect Dis. 2005;191:1014-1017.

    Not in our library10.1086/427997

  41. 41.

    Happi CT, Gbotosho GO, Folarin OA, Sowunmi A, Hudson T, O'Neil M, Milhous W, Wirth DF, Oduola AM. Selection of Plasmodium falciparum multidrug resistance gene 1 alleles in asexual stages and gametocytes by artemether-lumefantrine in Nigerian children with uncomplicated falciparum malaria. Antimicrob Agents Chemother. 2009;53:888-895.

    Not in our library10.1128/aac.00968-08

  42. 42.

    Some AF, Sere YY, Dokomajilar C, Zongo I, Rouamba N, Greenhouse B, Ouedraogo JB, Rosenthal PJ. Selection of known Plasmodium falciparum resistance-mediating polymorphisms by artemether-lumefantrine and amodiaquine-sulfadoxine-pyrimethamine but not dihydroartemisinin-piperaquine in Burkina Faso. Antimicrob Agents Chemother. 2010;54:1949-1954.

    Not in our library10.1128/aac.01413-09

  43. 43.

    Baliraine FN, Rosenthal PJ. Prolonged selection of pfmdr1 polymorphisms after treatment of falciparum malaria with artemether-lumefantrine in Uganda. J Infect Dis. 2011;204:1120-1124.

    Not in our library10.1093/infdis/jir486

  44. 44.

    Dorsey G, Staedke S, Clark TD, Njama-Meya D, Nzarubara B, Maiteki-Sebuguzi C, Dokomajilar C, Kamya MR, Rosenthal PJ. Combination therapy for uncomplicated falciparum malaria in Ugandan children: a randomized trial. JAMA. 2007;297:2210-2219.

    Not in our library10.1001/jama.297.20.2210

  45. 45.
    A Head-to-Head Comparison of Four Artemisinin-Based Combinations for Treating Uncomplicated Malaria in African Children: A Randomized Trial · 2011
    No notice
  46. 46.

    Fidock DA, Nomura T, Talley AK, Cooper RA, Dzekunov SM, Ferdig MT, Ursos LM, Sidhu AB, Naude B, Deitsch KW, Su XZ, Wootton JC, Roepe PD, Wellems TE. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol Cell. 2000;6:861-871.

    Not in our library10.1016/s1097-2765(05)00077-8

  47. 47.

    Martin RE, Kirk K. The malaria parasite's chloroquine resistance transporter is a member of the drug/metabolite transporter superfamily. Mol Biol Evol. 2004;21:1938-1949.

    Not in our library10.1093/molbev/msh205

  48. 48.

    Tran CV, Saier MH. The principal chloroquine resistance protein of Plasmodium falciparum is a member of the drug/metabolite transporter superfamily. Microbiology. 2004;150:1-3.

    Not in our library10.1099/mic.0.26818-0

  49. 49.

    Ecker A, Lehane AM, Clain J, Fidock DA. PfCRT and its role in antimalarial drug resistance. Trends Parasitol. 2012;28:504-514.

    Not in our library10.1016/j.pt.2012.08.002

  50. 50.

    Lakshmanan V, Bray PG, Verdier-Pinard D, Johnson DJ, Horrocks P, Muhle RA, Alakpa GE, Hughes RH, Ward SA, Krogstad DJ, Sidhu AB, Fidock DA. A critical role for PfCRT K76T in Plasmodium falciparum verapamil-reversible chloroquine resistance. EMBO J. 2005;24:2294-2305.

    Not in our library10.1038/sj.emboj.7600681

  51. 51.

    Sidhu AB, Verdier-Pinard D, Fidock DA. Chloroquine resistance in Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science. 2002;298:210-213.

    Not in our library10.1126/science.1074045

  52. 52.

    Dahlstrom S, Ferreira PE, Veiga MI, Sedighi N, Wiklund L, Martensson A, Farnert A, Sisowath C, Osorio L, Darban H, Andersson B, Kaneko A, Conseil G, Bjorkman A, Gil JP. Plasmodium falciparum multidrug resistance protein 1 and artemisinin-based combination therapy in Africa. J Infect Dis. 2009;200:1456-1464.

    Not in our library10.1086/606009

  53. 53.

    Dahlstrom S, Veiga MI, Martensson A, Bjorkman A, Gil JP. Polymorphism in PfMRP1 (Plasmodium falciparum multidrug resistance protein 1) amino acid 1466 associated with resistance to sulfadoxine-pyrimethamine treatment. Antimicrob Agents Chemother. 2009;53:2553-2556.

    Not in our library10.1128/aac.00091-09

  54. 54.

    Gupta B, Xu S, Wang Z, Sun L, Miao J, Cui L, Yang Z. Plasmodium falciparum multidrug resistance protein 1 (pfmrp1) gene and its association with in vitro drug susceptibility of parasite isolates from north-east Myanmar. J Antimicrob Chemother. 2014;69:2110-2117.

    Not in our library10.1093/jac/dku125

  55. 55.

    Raj DK, Mu J, Jiang H, Kabat J, Singh S, Sullivan M, Fay MP, McCutchan TF, Su XZ. Disruption of a Plasmodium falciparum multidrug resistance-associated protein (PfMRP) alters its fitness and transport of antimalarial drugs and glutathione. J Biol Chem. 2009;284:7687-7696.

    Not in our library10.1074/jbc.m806944200

  56. 56.

    Ferdig MT, Cooper RA, Mu J, Deng B, Joy DA, Su XZ, Wellems TE. Dissecting the loci of low-level quinine resistance in malaria parasites. Mol Microbiol. 2004;52:985-997.

    Not in our library10.1111/j.1365-2958.2004.04035.x

  57. 57.
    Na+ Regulation in the Malaria Parasite Plasmodium falciparum Involves the Cation ATPase PfATP4 and Is a Target of the Spiroindolone Antimalarials · 2013
    No notice
  58. 58.

    Vaidya AB, Morrisey JM, Zhang Z, Das S, Daly TM, Otto TD, Spillman NJ, Wyvratt M, Siegl P, Marfurt J, Wirjanata G, Sebayang BF, Price RN, Chatterjee A, Nagle A, Stasiak M, Charman SA, Angulo-Barturen I, Ferrer S, Belen Jimenez-Diaz M, Martinez MS, Gamo FJ, Avery VM, Ruecker A, Delves M, Kirk K, Berriman M, Kortagere S, Burrows J, Fan E, Bergman LW. Pyrazoleamide compounds are potent antimalarials that target Na+ homeostasis in intraerythrocytic Plasmodium falciparum. Nat Commun. 2014;5:5521.

    Not in our library10.1038/ncomms6521

  59. 59.

    Jimenez-Diaz MB, Ebert D, Salinas Y, Pradhan A, Lehane AM, Myrand-Lapierre ME, O'Loughlin KG, Shackleford DM, Justino de Almeida M, Carrillo AK, Clark JA, Dennis AS, Diep J, Deng X, Duffy S, Endsley AN, Fedewa G, Guiguemde WA, Gomez MG, Holbrook G, Horst J, Kim CC, Liu J, Lee MC, Matheny A, Martinez MS, Miller G, Rodriguez-Alejandre A, Sanz L, Sigal M, Spillman NJ, Stein PD, Wang Z, Zhu F, Waterson D, Knapp S, Shelat A, Avery VM, Fidock DA, Gamo FJ, Charman SA, Mirsalis JC, Ma H, Ferrer S, Kirk K, Angulo-Barturen I, Kyle DE, DeRisi JL, Floyd DM, Guy RK. (+)-SJ733, a clinical candidate for malaria that acts through ATP4 to induce rapid host-mediated clearance of Plasmodium. Proc Natl Acad Sci USA. 2014;111:E5455-E5462.

    Not in our library10.1073/pnas.1414221111

  60. 60.

    Rottmann M, McNamara C, Yeung BK, Lee MC, Zou B, Russell B, Seitz P, Plouffe DM, Dharia NV, Tan J, Cohen SB, Spencer KR, Gonzalez-Paez GE, Lakshminarayana SB, Goh A, Suwanarusk R, Jegla T, Schmitt EK, Beck HP, Brun R, Nosten F, Renia L, Dartois V, Keller TH, Fidock DA, Winzeler EA, Diagana TT. Spiroindolones, a potent compound class for the treatment of malaria. Science. 2010;329:1175-1180.

    Not in our library10.1126/science.1193225

  61. 61.

    Lehane AM, Ridgway MC, Baker E, Kirk K. Diverse chemotypes disrupt ion homeostasis in the malaria parasite. Mol Microbiol. 2014;94:327-339.

    Not in our library10.1111/mmi.12765

  62. 62.

    White NJ, Pukrittayakamee S, Phyo AP, Rueangweerayut R, Nosten F, Jittamala P, Jeeyapant A, Jain JP, Lefevre G, Li R, Magnusson B, Diagana TT, Leong FJ. Spiroindolone KAE609 for falciparum and vivax malaria. N Engl J Med. 2014;371:403-410.

    Not in our library10.1056/nejmoa1315860

  63. 63.

    da Silva AF, Benchimol JL. Malaria and quinine resistance: a medical and scientific issue between Brazil and Germany (1907–19). Med Hist. 2014;58:1-26.

    Not in our library10.1017/mdh.2013.69

  64. 64.

    Giboda M, Denis MB. Response of Kampuchean strains of Plasmodium falciparum to antimalarials: in-vivo assessment of quinine and quinine plus tetracycline; multiple drug resistance in vitro. J Trop Med Hyg. 1988;91:205-211.

    Not in our library

  65. 65.

    Pukrittayakamee S, Supanaranond W, Looareesuwan S, Vanijanonta S, White NJ. Quinine in severe falciparum malaria: evidence of declining efficacy in Thailand. Trans R Soc Trop Med Hyg. 1994;88:324-327.

    Not in our library10.1016/0035-9203(94)90102-3

  66. 66.

    Cowman AF, Galatis D, Thompson JK. Selection for mefloquine resistance in Plasmodium falciparum is linked to amplification of the pfmdr1 gene and cross-resistance to halofantrine and quinine. Proc Natl Acad Sci USA. 1994;91:1143-1147.

    Not in our library10.1073/pnas.91.3.1143

  67. 67.

    Henry M, Briolant S, Zettor A, Pelleau S, Baragatti M, Baret E, Mosnier J, Amalvict R, Fusai T, Rogier C, Pradines B. Plasmodium falciparum Na+/H+ exchanger 1 transporter is involved in reduced susceptibility to quinine. Antimicrob Agents Chemother. 2009;53:1926-1930.

    Not in our library10.1128/aac.01243-08

  68. 68.

    Meng H, Zhang R, Yang H, Fan Q, Su X, Miao J, Cui L, Yang Z. In vitro sensitivity of Plasmodium falciparum clinical isolates from the China-Myanmar border area to quinine and association with polymorphism in the Na+/H+ exchanger. Antimicrob Agents Chemother. 2010;54:4306-4313.

    Not in our library10.1128/aac.00321-10

  69. 69.

    Okombo J, Kiara SM, Rono J, Mwai L, Pole L, Ohuma E, Borrmann S, Ochola LI, Nzila A. In vitro activities of quinine and other antimalarials and pfnhe polymorphisms in Plasmodium isolates from Kenya. Antimicrob Agents Chemother. 2010;54:3302-3307.

    Not in our library10.1128/aac.00325-10

  70. 70.

    Andriantsoanirina V, Menard D, Rabearimanana S, Hubert V, Bouchier C, Tichit M, Bras JL, Durand R. Association of microsatellite variations of Plasmodium falciparum Na+/H+ exchanger (Pfnhe-1) gene with reduced in vitro susceptibility to quinine: lack of confirmation in clinical isolates from Africa. Am J Trop Med Hyg. 2010;82:782-787.

    Not in our library10.4269/ajtmh.2010.09-0327

  71. 71.

    Baliraine FN, Nsobya SL, Achan J, Tibenderana JK, Talisuna AO, Greenhouse B, Rosenthal PJ. Limited ability of Plasmodium falciparum pfcrt, pfmdr1, and pfnhe1 polymorphisms to predict quinine in vitro sensitivity or clinical effectiveness in Uganda. Antimicrob Agents Chemother. 2011;55:615-622.

    Not in our library10.1128/aac.00954-10

  72. 72.

    Sinou V, Quang le H, Pelleau S, Huong VN, Huong NT, Tai le M, Bertaux L, Desbordes M, Latour C, Long LQ, Thanh NX, Parzy D. Polymorphism of Plasmodium falciparum Na+/H+ exchanger is indicative of a low in vitro quinine susceptibility in isolates from Viet Nam. Malar J. 2011;10:164.

    Not in our library10.1186/1475-2875-10-164

  73. 73.

    Nkrumah LJ, Riegelhaupt PM, Moura P, Johnson DJ, Patel J, Hayton K, Ferdig MT, Wellems TE, Akabas MH, Fidock DA. Probing the multifactorial basis of Plasmodium falciparum quinine resistance: evidence for a strain-specific contribution of the sodium-proton exchanger PfNHE. Mol Biochem Parasitol. 2009;165:122-131.

    Not in our library10.1016/j.molbiopara.2009.01.011

  74. 74.

    Hernandez T, Myatt AV, Coatney GR, Jeffery GM. Studies in human malaria. XXXIV. Acquired resistance to pyrimethamine (Daraprim) by the Chesson strain of Plasmodium vivax. Am J Trop Med Hyg. 1953;2:797-804.

    Not in our library

  75. 75.

    Jones SA. Experiment to determine if a proguanil-resistant strain of P. falciparum would respond to large doses of pyrimethamine. BMJ. 1953;1:977.

    Not in our library10.1136/bmj.1.4817.977

  76. 76.

    Ferraroni JJ, Hayes J. Drug-resistant falciparum malaria among the Mayongong Indians in the Brazilian Amazon. Am J Trop Med Hyg. 1979;28:909-911.

    Not in our library

  77. 77.

    Nurse GT. Fansidar-resistant falciparum malaria in Papua New Guinea. Lancet. 1981;1:36-37.

    Not in our library10.1016/s0140-6736(81)90133-1

  78. 78.

    Rumans LW, Dennis DT, Atmosoedjono S. Fansidar resistant falciparum malaria in Indonesia. Lancet. 1979;2:580-581.

    Not in our library10.1016/s0140-6736(79)91633-7

  79. 79.

    Hurwitz ES, Johnson D, Campbell CC. Resistance of Plasmodium falciparum malaria to sulfadoxine-pyrimethamine (‘Fansidar’) in a refugee camp in Thailand. Lancet. 1981;1:1068-1070.

    Not in our library10.1016/s0140-6736(81)92239-x

  80. 80.

    Cowman AF, Morry MJ, Biggs BA, Cross GA, Foote SJ. Amino acid changes linked to pyrimethamine resistance in the dihydrofolate reductase-thymidylate synthase gene of Plasmodium falciparum. Proc Natl Acad Sci USA. 1988;85:9109-9113.

    Not in our library10.1073/pnas.85.23.9109

  81. 81.

    Peterson DS, Walliker D, Wellems TE. Evidence that a point mutation in dihydrofolate reductase-thymidylate synthase confers resistance to pyrimethamine in falciparum malaria. Proc Natl Acad Sci USA. 1988;85:9114-9118.

    Not in our library10.1073/pnas.85.23.9114

  82. 82.

    Nankoberanyi S, Mbogo GW, LeClair NP, Conrad MD, Tumwebaze P, Tukwasibwe S, Kamya MR, Tappero J, Nsobya SL, Rosenthal PJ. Validation of the ligase detection reaction fluorescent microsphere assay for the detection of Plasmodium falciparum resistance mediating polymorphisms in Uganda. Malar J. 2014;13:95.

    Not in our library10.1186/1475-2875-13-95

  83. 83.

    Roper C, Pearce R, Nair S, Sharp B, Nosten F, Anderson T. Intercontinental spread of pyrimethamine-resistant malaria. Science. 2004;305:1124.

    Not in our library10.1126/science.1098876

  84. 84.

    Lumb V, Das MK, Singh N, Dev V, Khan W, Sharma YD. Multiple origins of Plasmodium falciparum dihydropteroate synthetase mutant alleles associated with sulfadoxine resistance in India. Antimicrob Agents Chemother. 2011;55:2813-2817.

    Not in our library10.1128/aac.01151-10

  85. 85.

    Alifrangis M, Nag S, Schousboe ML, Ishengoma D, Lusingu J, Pota H, Kavishe RA, Pearce R, Ord R, Lynch C, Dejene S, Cox J, Rwakimari J, Minja DT, Lemnge MM, Roper C. Independent origin of Plasmodium falciparum antifolate super-resistance, Uganda, Tanzania, and Ethiopia. Emerg Infect Dis. 2014;20:1280-1286.

    Not in our library10.3201/eid2008.131897

  86. 86.

    Jain V, Basak S, Bhandari S, Bharti PK, Thomas T, Singh MP, Singh N. Burden of complicated malaria in a densely forested Bastar region of Chhattisgarh State (central India). PLoS One. 2014;9:e115266.

    Not in our library10.1371/journal.pone.0115266

  87. 87.

    ter Kuile FO, van Eijk AM, Filler SJ. Effect of sulfadoxine-pyrimethamine resistance on the efficacy of intermittent preventive therapy for malaria control during pregnancy: a systematic review. JAMA. 2007;297:2603-2616.

    Not in our library10.1001/jama.297.23.2603

  88. 88.

    Konate AT, Yaro JB, Ouedraogo AZ, Diarra A, Gansane A, Soulama I, Kangoye DT, Kabore Y, Ouedraogo E, Ouedraogo A, Tiono AB, Ouedraogo IN, Chandramohan D, Cousens S, Milligan PJ, Sirima SB, Greenwood B, Diallo DA. Intermittent preventive treatment of malaria provides substantial protection against malaria in children already protected by an insecticide-treated bednet in Burkina Faso: a randomised, double-blind, placebo-controlled trial. PLoS Med. 2011;8:e1000408.

    Not in our library10.1371/journal.pmed.1000408

  89. 89.

    Naidoo I, Roper C. Mapping ‘partially resistant’, ‘fully resistant’, and ‘super resistant’ malaria. Trends Parasitol. 2013;29:505-515.

    Not in our library10.1016/j.pt.2013.08.002

  90. 90.

    Parikh S, Rosenthal PJ. Intermittent preventive therapy for malaria in pregnancy: is sulfadoxine-pyrimethamine the right drug? Clin Pharmacol Ther. 2010;87:160-162.

    Not in our library10.1038/clpt.2009.284

  91. 91.

    Miao M, Yang Z, Cui L, Ahlum J, Huang Y, Cui L. Different allele prevalence in the dihydrofolate reductase and dihydropteroate synthase genes in Plasmodium vivax populations from China. Am J Trop Med Hyg. 2010;83:1206-1211.

    Not in our library10.4269/ajtmh.2010.10-0259

  92. 92.

    Auliff A, Wilson DW, Russell B, Gao Q, Chen N, Anh le N, Maguire J, Bell D, O'Neil MT, Cheng Q. Amino acid mutations in Plasmodium vivax dhfr and dhps from several geographical regions and susceptibility to antifolate drugs. Am J Trop Med Hyg. 2006;75:617-621.

    Not in our library

  93. 93.

    Prajapati SK, Joshi H, Dev V, Dua VK. Molecular epidemiology of Plasmodium vivax anti-folate resistance in India. Malar J. 2011;10:102.

    Not in our library10.1186/1475-2875-10-102

  94. 94.

    Ganguly S, Saha P, Chatterjee M, Maji AK. Prevalence of polymorphisms in antifolate drug resistance molecular marker genes pvdhfr and pvdhps in clinical isolates of Plasmodium vivax from Kolkata, India. Antimicrob Agents Chemother. 2014;58:196-200.

    Not in our library10.1128/aac.01171-13

  95. 95.
    Declining Artesunate-Mefloquine Efficacy against Falciparum Malaria on the Cambodia–Thailand Border · 2008
    No notice
  96. 96.

    Noedl H, Se Y, Schaecher K, Smith BL, Socheat D, Fukuda MM. Evidence of artemisinin-resistant malaria in western Cambodia. N Engl J Med. 2008;359:2619-2620.

    Not in our library10.1056/nejmc0805011

  97. 97.

    Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, Lwin KM, Ariey F, Hanpithakpong W, Lee SJ, Ringwald P, Silamut K, Imwong M, Chotivanich K, Lim P, Herdman T, An SS, Yeung S, Singhasivanon P, Day NP, Lindegardh N, Socheat D, White NJ. Artemisinin resistance in Plasmodium falciparum malaria. N Engl J Med. 2009;361:455-467.

    Not in our library10.1056/nejmoa0808859

  98. 98.

    Phyo AP, Nkhoma S, Stepniewska K, Ashley EA, Nair S, McGready R, ler Moo C, Al-Saai S, Dondorp AM, Lwin KM, Singhasivanon P, Day NP, White NJ, Anderson TJ, Nosten F. Emergence of artemisinin-resistant malaria on the western border of Thailand: a longitudinal study. Lancet. 2012;379:1960-1966.

    Not in our library10.1016/s0140-6736(12)60484-x

  99. 99.

    Witkowski B, Amaratunga C, Khim N, Sreng S, Chim P, Kim S, Lim P, Mao S, Sopha C, Sam B, Anderson JM, Duong S, Chuor CM, Taylor WR, Suon S, Mercereau-Puijalon O, Fairhurst RM, Menard D. Novel phenotypic assays for the detection of artemisinin-resistant Plasmodium falciparum malaria in Cambodia: in-vitro and ex-vivo drug-response studies. Lancet Infect Dis. 2013;13:1043-1049.

    Not in our library10.1016/s1473-3099(13)70252-4

  100. 100.

    Witkowski B, Khim N, Chim P, Kim S, Ke S, Kloeung N, Chy S, Duong S, Leang R, Ringwald P, Dondorp AM, Tripura R, Benoit-Vical F, Berry A, Gorgette O, Ariey F, Barale JC, Mercereau-Puijalon O, Menard D. Reduced artemisinin susceptibility of Plasmodium falciparum ring stages in western Cambodia. Antimicrob Agents Chemother. 2013;57:914-923.

    Not in our library10.1128/aac.01868-12

  101. 101.

    Chen N, Chavchich M, Peters JM, Kyle DE, Gatton ML, Cheng Q. Deamplification of pfmdr1-containing amplicon on chromosome 5 in Plasmodium falciparum is associated with reduced resistance to artelinic acid in vitro. Antimicrob Agents Chemother. 2010;54:3395-3401.

    Not in our library10.1128/aac.01421-09

  102. 102.

    Cui L, Wang Z, Miao J, Miao M, Chandra R, Jiang H, Su XZ, Cui L. Mechanisms of in vitro resistance to dihydroartemisinin in Plasmodium falciparum. Mol Microbiol. 2012;86:111-128.

    Not in our library10.1111/j.1365-2958.2012.08180.x

  103. 103.

    Cheeseman IH, Miller BA, Nair S, Nkhoma S, Tan A, Tan JC, Al Saai S, Phyo AP, Moo CL, Lwin KM, McGready R, Ashley E, Imwong M, Stepniewska K, Yi P, Dondorp AM, Mayxay M, Newton PN, White NJ, Nosten F, Ferdig MT, Anderson TJ. A major genome region underlying artemisinin resistance in malaria. Science. 2012;336:79-82.

    Not in our library10.1126/science.1215966

  104. 104.

    Takala-Harrison S, Clark TG, Jacob CG, Cummings MP, Miotto O, Dondorp AM, Fukuda MM, Nosten F, Noedl H, Imwong M, Bethell D, Se Y, Lon C, Tyner SD, Saunders DL, Socheat D, Ariey F, Phyo AP, Starzengruber P, Fuehrer HP, Swoboda P, Stepniewska K, Flegg J, Arze C, Cerqueira GC, Silva JC, Ricklefs SM, Porcella SF, Stephens RM, Adams M, Kenefic LJ, Campino S, Auburn S, MacInnis B, Kwiatkowski DP, Su XZ, White NJ, Ringwald P, Plowe CV. Genetic loci associated with delayed clearance of Plasmodium falciparum following artemisinin treatment in southeast Asia. Proc Natl Acad Sci USA. 2013;110:240-245.

    Not in our library10.1073/pnas.1211205110

  105. 105.

    Ariey F, Witkowski B, Amaratunga C, Beghain J, Langlois AC, Khim N, Kim S, Duru V, Bouchier C, Ma L, Lim P, Leang R, Duong S, Sreng S, Suon S, Chuor CM, Bout DM, Menard S, Rogers WO, Genton B, Fandeur T, Miotto O, Ringwald P, Le Bras J, Berry A, Barale JC, Fairhurst RM, Benoit-Vical F, Mercereau-Puijalon O, Menard D. A molecular marker of artemisinin-resistant Plasmodium falciparum malaria. Nature. 2014;505:50-55.

    Not in our library10.1038/nature12876

  106. 106.

    Ashley EA, Dhorda M, Fairhurst RM, Amaratunga C, Lim P, Suon S, Sreng S, Anderson JM, Mao S, Sam B, Sopha C, Chuor CM, Nguon C, Sovannaroth S, Pukrittayakamee S, Jittamala P, Chotivanich K, Chutasmit K, Suchatsoonthorn C, Runcharoen R, Hien TT, Thuy-Nhien NT, Thanh NV, Phu NH, Htut Y, Han KT, Aye KH, Mokuolu OA, Olaosebikan RR, Folaranmi OO, Mayxay M, Khanthavong M, Hongvanthong B, Newton PN, Onyamboko MA, Fanello CI, Tshefu AK, Mishra N, Valecha N, Phyo AP, Nosten F, Yi P, Tripura R, Borrmann S, Bashraheil M, Peshu J, Faiz MA, Ghose A, Hossain MA, Samad R, Rahman MR, Hasan MM, Islam A, Miotto O, Amato R, MacInnis B, Stalker J, Kwiatkowski DP, Bozdech Z, Jeeyapant A, Cheah PY, Sakulthaew T, Chalk J, Intharabut B, Silamut K, Lee SJ, Vihokhern B, Kunasol C, Imwong M, Tarning J, Taylor WJ, Yeung S, Woodrow CJ, Flegg JA, Das D, Smith J, Venkatesan M, Plowe CV, Stepniewska K, Guerin PJ, Dondorp AM, Day NP, White NJ, Tracking Resistance to Artemisinin Collaboration. Spread of artemisinin resistance in Plasmodium falciparum malaria. N Engl J Med. 2014;371:411-423.

    Not in our library10.1056/nejmoa1314981

  107. 107.

    Bosman P, Stassijns J, Nackers F, Canier L, Kim N, Khim S, Alipon SC, Chuor Char M, Chea N, Dysoley L, Van den Bergh R, Etienne W, De Smet M, Menard D, Kindermans JM. Plasmodium prevalence and artemisinin-resistant falciparum malaria in Preah Vihear Province, Cambodia: a cross-sectional population-based study. Malar J. 2014;13:394.

    Not in our library10.1186/1475-2875-13-394

  108. 108.
    Spread of artemisinin-resistant Plasmodium falciparum in Myanmar: a cross-sectional survey of the K13 molecular marker · 2015
    No notice
  109. 109.

    Saunders DL, Vanachayangkul P, Lon C. Dihydroartemisinin-piperaquine failure in Cambodia. N Engl J Med. 2014;371:484-485.

    Not in our library10.1056/nejmc1403007

  110. 110.
    Polymorphisms in K13 and falcipain-2 associated with artemisinin resistance are not prevalent in Plasmodium falciparum isolated from Ugandan children. · 2014
    Not checked
  111. 111.

    Kamau E, Campino S, Amenga-Etego L, Drury E, Ishengoma D, Johnson K, Mumba D, Kekre M, Yavo W, Mead D, Bouyou-Akotet M, Apinjoh T, Golassa L, Randrianarivelojosia M, Andagalu B, Maiga-Ascofare O, Amambua-Ngwa A, Tindana P, Ghansah A, MacInnis B, Kwiatkowski D, Djimde AA. K13-propeller polymorphisms in Plasmodium falciparum parasites from sub-Saharan Africa. J Infect Dis. 2014;211:1352-1355.

    Not in our library10.1093/infdis/jiu608

  112. 112.

    Taylor SM, Parobek CM, DeConti DK, Kayentao K, Coulibaly SO, Greenwood BM, Tagbor H, Williams J, Bojang K, Njie F, Desai M, Kariuki S, Gutman J, Mathanga DP, Martensson A, Ngasala B, Conrad MD, Rosenthal PJ, Tshefu AK, Moormann AM, Vulule JM, Doumbo OK, Ter Kuile FO, Meshnick SR, Bailey JA, Juliano JJ. Absence of putative artemisinin resistance mutations among Plasmodium falciparum in sub-Saharan Africa: a molecular epidemiologic study. J Infect Dis. 2014;211:680-688.

    Not in our library10.1093/infdis/jiu467

  113. 113.

    Straimer J, Gnadig NF, Witkowski B, Amaratunga C, Duru V, Ramadani AP, Dacheux M, Khim N, Zhang L, Lam S, Gregory PD, Urnov FD, Mercereau-Puijalon O, Benoit-Vical F, Fairhurst RM, Menard D, Fidock DA. K13-propeller mutations confer artemisinin resistance in Plasmodium falciparum clinical isolates. Science. 2014;347:428-431.

    Not in our library10.1126/science.1260867

  114. 114.

    Mok S, Ashley EA, Ferreira PE, Zhu L, Lin Z, Yeo T, Chotivanich K, Imwong M, Pukrittayakamee S, Dhorda M, Nguon C, Lim P, Amaratunga C, Suon S, Hien TT, Htut Y, Faiz MA, Onyamboko MA, Mayxay M, Newton PN, Tripura R, Woodrow CJ, Miotto O, Kwiatkowski DP, Nosten F, Day NP, Preiser PR, White NJ, Dondorp AM, Fairhurst RM, Bozdech Z. Population transcriptomics of human malaria parasites reveals the mechanism of artemisinin resistance. Science. 2014;347:431-435.

    Not in our library10.1126/science.1260403

  115. 115.

    Vaidya AB, Lashgari MS, Pologe LG, Morrisey J. Structural features of Plasmodium cytochrome b that may underlie susceptibility to 8-aminoquinolines and hydroxynaphthoquinones. Mol Biochem Parasitol. 1993;58:33-42.

    Not in our library10.1016/0166-6851(93)90088-f

  116. 116.

    Srivastava IK, Rottenberg H, Vaidya AB. Atovaquone, a broad spectrum antiparasitic drug, collapses mitochondrial membrane potential in a malarial parasite. J Biol Chem. 1997;272:3961-3966.

    Not in our library10.1074/jbc.272.7.3961

  117. 117.

    Fivelman QL, Butcher GA, Adagu IS, Warhurst DC, Pasvol G. Malarone treatment failure and in vitro confirmation of resistance of Plasmodium falciparum isolate from Lagos, Nigeria. Malar J. 2002;1:1.

    Not in our library10.1186/1475-2875-1-1

  118. 118.

    Korsinczky M, Chen N, Kotecka B, Saul A, Rieckmann K, Cheng Q. Mutations in Plasmodium falciparum cytochrome b that are associated with atovaquone resistance are located at a putative drug-binding site. Antimicrob Agents Chemother. 2000;44:2100-2108.

    Not in our library10.1128/aac.44.8.2100-2108.2000

  119. 119.

    Wichmann O, Muehlen M, Gruss H, Mockenhaupt FP, Suttorp N, Jelinek T. Malarone treatment failure not associated with previously described mutations in the cytochrome b gene. Malar J. 2004;3:14.

    Not in our library10.1186/1475-2875-3-14

  120. 120.

    Musset L, Bouchaud O, Matheron S, Massias L, Le Bras J. Clinical atovaquone-proguanil resistance of Plasmodium falciparum associated with cytochrome b codon 268 mutations. Microbes Infect. 2006;8:2599-2604.

    Not in our library10.1016/j.micinf.2006.07.011

  121. 121.

    Wichmann O, Muehlberger N, Jelinek T, Alifrangis M, Peyerl-Hoffmann G, Muhlen M, Grobusch MP, Gascon J, Matteelli A, Laferl H, Bisoffi Z, Ehrhardt S, Cuadros J, Hatz C, Gjorup I, McWhinney P, Beran J, da Cunha S, Schulze M, Kollaritsch H, Kern P, Fry G, Richter J, European Network on Surveillance of Imported Infectious Diseases. Screening for mutations related to atovaquone/proguanil resistance in treatment failures and other imported isolates of Plasmodium falciparum in Europe. J Infect Dis. 2004;190:1541-1546.

    Not in our library10.1086/424469

  122. 122.

    Wang Y, Yang Z, Yuan L, Zhou G, Lee M-C, Fan Q, Xiao Y, Cao Y, Yan G, Cui L. Clinical efficacy of dihydroartemisinin-piperaquine for the treatment of uncomplicated Plasmodium falciparum malaria at the China-Myanmar border. Am J Trop Med Hyg. 2015;93:577-583.

    Not in our library10.4269/ajtmh.15-0029

  123. 123.

    Achan J, Tibenderana JK, Kyabayinze D, Wabwire Mangen F, Kamya MR, Dorsey G, D'Alessandro U, Rosenthal PJ, Talisuna AO. Effectiveness of quinine versus artemether-lumefantrine for treating uncomplicated falciparum malaria in Ugandan children: randomised trial. BMJ. 2009;339:b2763.

    Not in our library10.1136/bmj.b2763

  124. 124.

    Liang GL, Sun XD, Wang J, Zhang ZX. Sensitivity of Plasmodium vivax to chloroquine in Laza City, Myanmar. Chin J Parasitol Parasitic Dis. 2009;27:175-176.

    Not in our library

  125. 125.

    Yuan L, Wang Y, Parker DM, Gupta B, Yang Z, Liu H, Fan Q, Cao Y, Xiao Y, Lee MC, Zhou G, Yan G, Baird JK, Cui L. Therapeutic responses of Plasmodium vivax malaria to chloroquine and primaquine treatment in northeastern Myanmar. Antimicrob Agents Chemother. 2015;59:1230-1235.

    Not in our library10.1128/aac.04270-14

  126. 126.

    Nsobya SL, Kiggundu M, Nanyunja S, Joloba M, Greenhouse B, Rosenthal PJ. In vitro sensitivities of Plasmodium falciparum to different antimalarial drugs in Uganda. Antimicrob Agents Chemother. 2010;54:1200-1206.

    Not in our library10.1128/aac.01412-09

  127. 127.

    Van Tyne D, Dieye B, Valim C, Daniels RF, Sene PD, Lukens AK, Ndiaye M, Bei AK, Ndiaye YD, Hamilton EJ, Ndir O, Mboup S, Volkman SK, Wirth DF, Ndiaye D. Changes in drug sensitivity and anti-malarial drug resistance mutations over time among Plasmodium falciparum parasites in Senegal. Malar J. 2013;12:441.

    Not in our library10.1186/1475-2875-12-441

  128. 128.

    Mungthin M, Khositnithikul R, Sitthichot N, Suwandittakul N, Wattanaveeradej V, Ward SA, Na-Bangchang K. Association between the pfmdr1 gene and in vitro artemether and lumefantrine sensitivity in Thai isolates of Plasmodium falciparum. Am J Trop Med Hyg. 2010;83:1005-1009.

    Not in our library10.4269/ajtmh.2010.10-0339

  129. 129.

    Ndiaye D, Patel V, Demas A, LeRoux M, Ndir O, Mboup S, Clardy J, Lakshmanan V, Daily JP, Wirth DF. A non-radioactive DAPI-based high-throughput in vitro assay to assess Plasmodium falciparum responsiveness to antimalarials—increased sensitivity of P. falciparum to chloroquine in Senegal. Am J Trop Med Hyg. 2010;82:228-230.

    Not in our library10.4269/ajtmh.2010.09-0470

  130. 130.

    Wang Z, Parker D, Meng H, Wu L, Li J, Zhao Z, Zhang R, Fan Q, Wang H, Cui L, Yang Z. In vitro sensitivity of Plasmodium falciparum from China-Myanmar border area to major ACT drugs and polymorphisms in potential target genes. PLoS One. 2012;7:e30927.

    Not in our library10.1371/journal.pone.0030927

  131. 131.

    Hao M, Jia D, Li Q, He Y, Yuan L, Xu S, Chen K, Wu J, Shen L, Sun L, Zhao H, Yang Z, Cui L. In vitro sensitivities of Plasmodium falciparum isolates from the China-Myanmar border to piperaquine and association with polymorphisms in candidate genes. Antimicrob Agents Chemother. 2013;57:1723-1729.

    Not in our library10.1128/aac.02306-12

  132. 132.

    Noedl H, Wongsrichanalai C, Wernsdorfer WH. Malaria drug-sensitivity testing: new assays, new perspectives. Trends Parasitol. 2003;19:175-181.

    Not in our library10.1016/s1471-4922(03)00028-x

  133. 133.

    Abdul-Ghani R, Al-Maktari MT, Al-Shibani LA, Allam AF. A better resolution for integrating methods for monitoring Plasmodium falciparum resistance to antimalarial drugs. Acta Trop. 2014;137:44-57.

    Not in our library10.1016/j.actatropica.2014.04.031

  134. 134.

    Mbogo GW, Nankoberanyi S, Tukwasibwe S, Baliraine FN, Nsobya SL, Conrad MD, Arinaitwe E, Kamya M, Tappero J, Staedke SG, Dorsey G, Greenhouse B, Rosenthal PJ. Temporal changes in prevalence of molecular markers mediating antimalarial drug resistance in a high malaria transmission setting in Uganda. Am J Trop Med Hyg. 2014;91:54-61.

    Not in our library10.4269/ajtmh.13-0647

  135. 135.

    Yang Z, Li C, Miao M, Zhang Z, Sun X, Meng H, Li J, Fan Q, Cui L. Multidrug-resistant genotypes of Plasmodium falciparum, Myanmar. Emerg Infect Dis. 2011;17:498-501.

    Not in our library10.3201/eid1703.100870

  136. 136.

    Bukirwa H, Yeka A, Kamya MR, Talisuna A, Banek K, Bakyaita N, Rwakimari JB, Rosenthal PJ, Wabwire-Mangen F, Dorsey G, Staedke SG. Artemisinin combination therapies for treatment of uncomplicated malaria in Uganda. PLoS Clin Trials. 2006;1:e7.

    Not in our library10.1371/journal.pctr.0010007

  137. 137.

    Ashley EA, Dhorda M, Fairhurst RM, Amaratunga C, Lim P, Suon S, Sreng S, Anderson JM, Mao S, Sam B, Sopha C, Chuor CM, Nguon C, Sovannaroth S, Pukrittayakamee S, Jittamala P, Chotivanich K, Chutasmit K, Suchatsoonthorn C, Runcharoen R, Hien TT, Thuy-Nhien NT, Thanh NV, Phu NH, Htut Y, Han KT, Aye KH, Mokuolu OA, Olaosebikan RR, Folaranmi OO, Mayxay M, Khanthavong M, Hongvanthong B, Newton PN, Onyamboko MA, Fanello CI, Tshefu AK, Mishra N, Valecha N, Phyo AP, Nosten F, Yi P, Tripura R, Borrmann S, Bashraheil M, Peshu J, Faiz MA, Ghose A, Hossain MA, Samad R, Rahman MR, Hasan MM, Islam A, Miotto O, Amato R, MacInnis B, Stalker J, Kwiatkowski DP, Bozdech Z, Jeeyapant A, Cheah PY, Sakulthaew T, Chalk J, Intharabut B, Silamut K, Lee SJ, Vihokhern B, Kunasol C, Imwong M, Tarning J, Taylor WJ, Yeung S, Woodrow CJ, Flegg JA, Das D, Smith J, Venkatesan M, Plowe CV, Stepniewska K, Guerin PJ, Dondorp AM, Day NP, White NJ. Spread of artemisinin resistance in Plasmodium falciparum malaria. N Engl J Med. 2014;371:411-423.

    Not in our library10.1056/nejmoa1314981

  138. 138.

    Felger I, Beck HP. Fitness costs of resistance to antimalarial drugs. Trends Parasitol. 2008;24:331-333.

    Not in our library10.1016/j.pt.2008.05.004

  139. 139.

    Hastings IM, Donnelly MJ. The impact of antimalarial drug resistance mutations on parasite fitness, and its implications for the evolution of resistance. Drug Resist Updat. 2005;8:43-50.

    Not in our library10.1016/j.drup.2005.03.003

  140. 140.

    Rosenthal PJ. The interplay between drug resistance and fitness in malaria parasites. Mol Microbiol. 2013;89:1025-1038.

    Not in our library10.1111/mmi.12349

  141. 141.

    Marks F, Evans J, Meyer CG, Browne EN, Flessner C, von Kalckreuth V, Eggelte TA, Horstmann RD, May J. High prevalence of markers for sulfadoxine and pyrimethamine resistance in Plasmodium falciparum in the absence of drug pressure in the Ashanti region of Ghana. Antimicrob Agents Chemother. 2005;49:1101-1105.

    Not in our library10.1128/aac.49.3.1101-1105.2005

  142. 142.

    Brown KM, Costanzo MS, Xu W, Roy S, Lozovsky ER, Hartl DL. Compensatory mutations restore fitness during the evolution of dihydrofolate reductase. Mol Biol Evol. 2010;27:2682-2690.

    Not in our library10.1093/molbev/msq160

  143. 143.

    Sandefur CI, Wooden JM, Quaye IK, Sirawaraporn W, Sibley CH. Pyrimethamine-resistant dihydrofolate reductase enzymes of Plasmodium falciparum are not enzymatically compromised in vitro. Mol Biochem Parasitol. 2007;154:1-5.

    Not in our library10.1016/j.molbiopara.2007.03.009

  144. 144.

    Abdel-Muhsin AM, Mackinnon MJ, Ali E, Nassir el-KA, Suleiman S, Ahmed S, Walliker D, Babiker HA. Evolution of drug-resistance genes in Plasmodium falciparum in an area of seasonal malaria transmission in eastern Sudan. J Infect Dis. 2004;189:1239-1244.

    Not in our library10.1086/382509

  145. 145.

    Preechapornkul P, Imwong M, Chotivanich K, Pongtavornpinyo W, Dondorp AM, Day NP, White NJ, Pukrittayakamee S. Plasmodium falciparum pfmdr1 amplification, mefloquine resistance, and parasite fitness. Antimicrob Agents Chemother. 2009;53:1509-1515.

    Not in our library10.1128/aac.00241-08

  146. 146.

    Temu EA, Kimani I, Tuno N, Kawada H, Minjas JN, Takagi M. Monitoring chloroquine resistance using Plasmodium falciparum parasites isolated from wild mosquitoes in Tanzania. Am J Trop Med Hyg. 2006;75:1182-1187.

    Not in our library

  147. 147.

    Oduola AM, Sowunmi A, Milhous WK, Kyle DE, Martin RK, Walker O, Salako LA. Innate resistance to new antimalarial drugs in Plasmodium falciparum from Nigeria. Trans R Soc Trop Med Hyg. 1992;86:123-126.

    Not in our library10.1016/0035-9203(92)90533-i

  148. 148.

    WHO. Advances in Malaria Chemotherapy. 1984.

    Not in our library

  149. 149.

    Hailemeskel E, Kassa M, Taddesse G, Mohammed H, Woyessa A, Tasew G, Sleshi M, Kebede A, Petros B. Prevalence of sulfadoxine-pyrimethamine resistance-associated mutations in dhfr and dhps genes of Plasmodium falciparum three years after SP withdrawal in Bahir Dar, northwest Ethiopia. Acta Trop. 2013;128:636-641.

    Not in our library10.1016/j.actatropica.2013.09.010

  150. 150.

    Ndiaye M, Faye B, Tine R, Ndiaye JL, Lo A, Abiola A, Dieng Y, Ndiaye D, Hallett R, Alifrangis M, Gaye O. Assessment of the molecular marker of Plasmodium falciparum chloroquine resistance (Pfcrt) in Senegal after several years of chloroquine withdrawal. Am J Trop Med Hyg. 2012;87:640-645.

    Not in our library10.4269/ajtmh.2012.11-0709

  151. 151.

    Mang'era CM, Mbai FN, Omedo IA, Mireji PO, Omar SA. Changes in genotypes of Plasmodium falciparum human malaria parasite following withdrawal of chloroquine in Tiwi, Kenya. Acta Trop. 2012;123:202-207.

    Not in our library10.1016/j.actatropica.2012.05.007

  152. 152.

    Mita T, Kaneko A, Lum JK, Bwijo B, Takechi M, Zungu IL, Tsukahara T, Tanabe K, Kobayakawa T, Bjorkman A. Recovery of chloroquine sensitivity and low prevalence of the Plasmodium falciparum chloroquine resistance transporter gene mutation K76T following the discontinuance of chloroquine use in Malawi. Am J Trop Med Hyg. 2003;68:413-415.

    Not in our library

  153. 153.

    Kublin JG, Cortese JF, Njunju EM, Mukadam RA, Wirima JJ, Kazembe PN, Djimde AA, Kouriba B, Taylor TE, Plowe CV. Reemergence of chloroquine-sensitive Plasmodium falciparum malaria after cessation of chloroquine use in Malawi. J Infect Dis. 2003;187:1870-1875.

    Not in our library10.1086/375419

  154. 154.

    Thaithong S, Suebsaeng L, Rooney W, Beale GH. Evidence of increased chloroquine sensitivity in Thai isolates of Plasmodium falciparum. Trans R Soc Trop Med Hyg. 1988;82:37-38.

    Not in our library10.1016/0035-9203(88)90255-6

  155. 155.

    de Almeida A, Arez AP, Cravo PV, do Rosario VE. Analysis of genetic mutations associated with anti-malarial drug resistance in Plasmodium falciparum from the Democratic Republic of East Timor. Malar J. 2009;8:59.

    Not in our library10.1186/1475-2875-8-59

  156. 156.

    Frank M, Lehners N, Mayengue PI, Gabor J, Dal-Bianco M, Kombila DU, Ngoma GM, Supan C, Lell B, Ntoumi F, Grobusch MP, Dietz K, Kremsner PG. A thirteen-year analysis of Plasmodium falciparum populations reveals high conservation of the mutant pfcrt haplotype despite the withdrawal of chloroquine from national treatment guidelines in Gabon. Malar J. 2011;10:304.

    Not in our library10.1186/1475-2875-10-304

  157. 157.

    Kamugisha E, Bujila I, Lahdo M, Pello-Esso S, Minde M, Kongola G, Naiwumbwe H, Kiwuwa S, Kaddumukasa M, Kironde F, Swedberg G. Acta Trop. 2012;121:148-151.

    Not in our library10.1016/j.actatropica.2011.11.004

  158. 158.
    Chloroquine resistance before and after its withdrawal in Kenya · 2009
    No notice
  159. 159.

    McCollum AM, Mueller K, Villegas L, Udhayakumar V, Escalante AA. Common origin and fixation of Plasmodium falciparum dhfr and dhps mutations associated with sulfadoxine-pyrimethamine resistance in a low-transmission area in South America. Antimicrob Agents Chemother. 2007;51:2085-2091.

    Not in our library10.1128/aac.01228-06

  160. 160.

    Babiker HA. Seasonal fluctuation of drug-resistant malaria parasites: a sign of fitness cost. Trends Parasitol. 2009;25:351-352.

    Not in our library10.1016/j.pt.2009.05.006

  161. 161.

    Babiker HA, Hastings IM, Swedberg G. Impaired fitness of drug-resistant malaria parasites: evidence and implication on drug-deployment policies. Expert Rev Anti Infect Ther. 2009;7:581-593.

    Not in our library10.1586/eri.09.29

  162. 162.

    Ord R, Alexander N, Dunyo S, Hallett R, Jawara M, Targett G, Drakeley CJ, Sutherland CJ. Seasonal carriage of pfcrt and pfmdr1 alleles in Gambian Plasmodium falciparum imply reduced fitness of chloroquine-resistant parasites. J Infect Dis. 2007;196:1613-1619.

    Not in our library10.1086/522154

  163. 163.
    No seasonal accumulation of resistant P. falciparum when high-dose chloroquine is used. · 2009
    Not checked
  164. 164.

    Asih PB, Rogers WO, Susanti AI, Rahmat A, Rozi IE, Kusumaningtyas MA, Krisin, Sekartuti, Dewi RM, Coutrier FN, Sutamihardja A, van der Ven AJ, Sauerwein RW, Syafruddin D. Seasonal distribution of anti-malarial drug resistance alleles on the island of Sumba, Indonesia. Malar J. 2009;8:222.

    Not in our library10.1186/1475-2875-8-222

  165. 165.

    Mbacham WF, Evehe MS, Netongo PM, Ateh IA, Mimche PN, Ajua A, Nji AM, Irenee D, Echouffo-Tcheugui JB, Tawe B, Hallett R, Roper C, Targett G, Greenwood B. Efficacy of amodiaquine, sulphadoxine-pyrimethamine and their combination for the treatment of uncomplicated Plasmodium falciparum malaria in children in Cameroon at the time of policy change to artemisinin-based combination therapy. Malar J. 2010;9:34.

    Not in our library10.1186/1475-2875-9-34

  166. 166.

    Bhumiratana A, Intarapuk A, Sorosjinda-Nunthawarasilp P, Maneekan P, Koyadun S. Border malaria associated with multidrug resistance on Thailand-Myanmar and Thailand-Cambodia borders: transmission dynamic, vulnerability, and surveillance. BioMed Res Int. 2013;2013:363417.

    Not in our library10.1155/2013/363417

  167. 167.

    Lukens AK, Ross LS, Heidebrecht R, Javier Gamo F, Lafuente-Monasterio MJ, Booker ML, Hartl DL, Wiegand RC, Wirth DF. Harnessing evolutionary fitness in Plasmodium falciparum for drug discovery and suppressing resistance. Proc Natl Acad Sci USA. 2014;111:799-804.

    Not in our library10.1073/pnas.1320886110

  168. 168.

    Lelievre J, Berry A, Benoit-Vical F. Artemisinin and chloroquine: do mode of action and mechanism of resistance involve the same protagonists? Curr Opin Investig Drugs. 2007;8:117-124.

    Not in our library

  169. 169.

    Sisowath C, Petersen I, Veiga MI, Martensson A, Premji Z, Bjorkman A, Fidock DA, Gil JP. In vivo selection of Plasmodium falciparum parasites carrying the chloroquine-susceptible pfcrt K76 allele after treatment with artemether-lumefantrine in Africa. J Infect Dis. 2009;199:750-757.

    Not in our library10.1086/596738

  170. 170.

    Froberg G, Jornhagen L, Morris U, Shakely D, Msellem MI, Gil JP, Bjorkman A, Martensson A. Decreased prevalence of Plasmodium falciparum resistance markers to amodiaquine despite its wide scale use as ACT partner drug in Zanzibar. Malar J. 2012;11:321.

    Not in our library10.1186/1475-2875-11-321

  171. 171.

    Tukwasibwe S, Mugenyi L, Mbogo GW, Nankoberanyi S, Maiteki-Sebuguzi C, Joloba ML, Nsobya SL, Staedke SG, Rosenthal PJ. Differential prevalence of transporter polymorphisms in symptomatic and asymptomatic falciparum malaria infections in Uganda. J Infect Dis. 2014;210:154-157.

    Not in our library10.1093/infdis/jiu044

  172. 172.

    Mharakurwa S, Sialumano M, Liu K, Scott A, Thuma P. Selection for chloroquine-sensitive Plasmodium falciparum by wild Anopheles arabiensis in southern Zambia. Malar J. 2013;12:453.

    Not in our library10.1186/1475-2875-12-453

  173. 173.

    Mharakurwa S, Kumwenda T, Mkulama MA, Musapa M, Chishimba S, Shiff CJ, Sullivan DJ, Thuma PE, Liu K, Agre P. Malaria antifolate resistance with contrasting Plasmodium falciparum dihydrofolate reductase (DHFR) polymorphisms in humans and Anopheles mosquitoes. Proc Natl Acad Sci USA. 2011;108:18796-18801.

    Not in our library10.1073/pnas.1116162108

  174. 174.

    Mharakurwa S. Plasmodium falciparum transmission rate and selection for drug resistance: a vexed association or a key to successful control? Int J Parasitol. 2004;34:1483-1487.

    Not in our library10.1016/j.ijpara.2004.10.011

  175. 175.

    Mharakurwa S, Mutambu SL, Mudyiradima R, Chimbadzwa T, Chandiwana SK, Day KP. Association of house spraying with suppressed levels of drug resistance in Zimbabwe. Malar J. 2004;3:35.

    Not in our library10.1186/1475-2875-3-35

  176. 176.
    High frequency of <it>Plasmodium falciparum </it>chloroquine resistance marker (<it>pfcrt </it>T76 mutation) in Yemen: An urgent need to re-examine malaria drug policy · 2011
    Not checked
  177. 177.

    Alifrangis M, Lemnge MM, Ronn AM, Segeja MD, Magesa SM, Khalil IF, Bygbjerg IC. Increasing prevalence of wildtypes in the dihydrofolate reductase gene of Plasmodium falciparum in an area with high levels of sulfadoxine/pyrimethamine resistance after introduction of treated bed nets. Am J Trop Med Hyg. 2003;69:238-243.

    Not in our library

  178. 178.

    Shah M, Kariuki S, Vanden Eng J, Blackstock AJ, Garner K, Gatei W, Gimnig JE, Lindblade K, Terlouw D, ter Kuile F, Hawley WA, Phillips-Howard P, Nahlen B, Walker E, Hamel MJ, Slutsker L, Shi YP. Effect of transmission reduction by insecticide-treated bednets (ITNs) on antimalarial drug resistance in western Kenya. PLoS One. 2011;6:e26746.

    Not in our library10.1371/journal.pone.0026746

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