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Trends Pharmacol Sci. Author manuscript; available in PMC Sep 23, 2010.
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PMCID: PMC2944664

Transporters involved in resistance to antimalarial drugs


The ability to treat and control Plasmodium falciparum infection through chemotherapy has been compromised by the advent and spread of resistance to antimalarial drugs. Research in this area has identified the P. falciparum chloroquine resistance transporter (PfCRT) and the multidrug resistance-1 (PfMDR1) transporter as key determinants of decreased in vitro susceptibility to several principal antimalarial drugs. Transfection-based in vitro studies are consistent with clinical findings of an association between mutations in the pfcrt gene and failure of chloroquine treatment, and between amplification of the pfmdr1 gene and failure of mefloquine treatment. Many countries are now switching to arte-misinin-based combination therapies. These incorporate partner drugs of which some have an in vitro efficacy that can be modulated by changes in pfcrt or pfmdr1. Here, we summarize investigations of these and other recently identified P. falciparum transporters in the context of antimalarial mode of action and mechanisms of resistance.

Malaria and drug resistance

The spread of drug resistant strains of the malaria parasite Plasmodium falciparum has led to a significant resurgence of malarial morbidity and mortality, and a growing crisis in global public health [1]. P. falciparum causes an estimated 500 million clinical infections and at least one million deaths annually, primarily in sub-Saharan Africa [1,2]. Infection begins when Anopheles mosquitoes deliver sporozoite forms that invade hepatocytes and replicate as liver-stage parasites, before emerging into the blood stream and infecting red blood cells. The asexual blood-stage infection causes clinical disease characterized by cyclical fevers and shaking chills, which can lead to complications such as severe anemia or cerebral malaria [3,4]. Red blood cells can also harbor sexual-stage gametocytes that can be transmitted to mosquitoes, where they undergo genetic recombination and complete the lifecycle of the parasite.

For decades, the treatment of malaria depended on chloroquine – an extensively used 4-aminoquinoline characterized by its rapid efficacy, low toxicity, availability and affordability [5]. The eventual appearance of chloroquine resistance in Southeast Asia and South America sparked the global dissemination of resistance [6]. The first-line treatment for chloroquine-resistant (CQR) malaria – sulfadoxine–pyrimethamine – rapidly met with resistance and is also becoming increasingly ineffective [7]. Failure of P. falciparum clinical treatment resulting from confirmed in vivo parasite resistance has now been documented for all current antimalarial drugs apart from the artemisinins, placing artemisinin-based combination therapy at the forefront of current malaria control programs [8,9].

Here, we review recent studies into the P. falciparum chloroquine resistance (PfCRT) and multidrug resistance-1 (PfMDR1) transporters that are now known to be key contributors to P. falciparum antimalarial drug resistance. We also briefly discuss the Ca2+ transporter PfATP6, which has been implicated in artemisinin action [1013] (for reviews on parasite determinants of resistance to antimalarial antifolates, mitochondrial inhibitors and antibiotics, see Refs [7,14,15]).


The putative transporter PfCRT was identified through the analysis of a genetic cross between a chloroquine-sensitive (CQS) and a CQR clone, which mapped resistance to the gene pfcrt [16,17]. The 45-kDa PfCRT protein contains ten predicted transmembrane domains and is located on the membrane of the digestive vacuole – an acidic, lysosome-like compartment in which hemoglobin is degraded and detoxified, and in which the weak base chloroquine concentrates in its diprotonated form and binds hematin (a dimeric form of oxidized heme) [6,18,19]. pfcrt shows extraordinary sequence diversity among geographically distinct isolates: point mutations have been detected at 15 residues and 4–8 individual mutations are present in individual CQR lines (Figure 1). This level of diversity corresponds to at least five independent origins of mutant pfcrt [20]. The Asian and African CQR alleles seem to confer a fitness cost, as evidenced by data from Malawi and China that show a decrease in the prevalence of these alleles upon discontinued use of chloroquine [21].

Figure 1
Predicted structure and representative haplotypes of P. falciparum chloroquine resistance transporter. (a) PfCRT is predicted to have ten transmembrane domains, with its N and C termini located on the cytoplasmic side of the digestive vacuole membrane ...

By allelic exchange, CQR alleles have been shown to confer in vitro chloroquine resistance to CQS parasites [22]. This resistance phenotype is defined as a half-maximal inhibitory concentration (IC50) of ≥80 nM, decreased [3H]chloroquine accumulation and verapamil reversibility. Removal of the K76T mutation, which is ubiquitous in CQR lines, results in CQR parasites becoming fully CQS and losing their verapamil reversibility phenotype [23]. It has not proved possible to introduce this single mutation into CQS parasites, indicating that this mutation might have a detrimental effect on function that needs to be compensated by other pfcrt mutations. These in vitro results are largely consistent with in vivo findings that document a strong association between the PfCRT K76T mutation and failure of chloroquine treatment, leading to its widespread use as a molecular marker of chloroquine resistance [5,7,21]. The in vivo studies have also found that some individuals carrying parasites with mutant pfcrt have an adequate clinical response to chloroquine treatment. This has been attributed, at least in part, to synergy between partially effective chloroquine treatment and acquired immunity [5].

Interestingly, studies have shown that the PfCRT transporter can also significantly influence parasite in vitro susceptibility to many antimalarial drugs including quinine, monodesethylamodiaquine (the primary metabolite of amodiaquine), halofantrine and artemisinin [18,2224]. A significant contribution of pfcrt to parasite susceptibility to antimalarial drugs in addition to chloroquine might explain the unusual diversity of pfcrt alleles, particularly in parts of Asia such as Cambodia where chloroquine is used rarely if at all against P. falciparum infection [25,26].

To disseminate, resistance determinants must not only ensure parasite survival against drug treatment but also be successfully transmitted through the gametocyte stage into the mosquito vector. Recent findings in the Gambia show a strong selection for mutant pfcrt in gametocyte populations in individuals exposed to chloroquine [27,28]. These mutants might prevent chloroquine from killing very-early-stage gametocytes [29] or, alternatively, game-tocytes harboring mutant pfcrt might have enhanced transmissibility. Such effects would provide a compelling explanation for how mutant pfcrt became so prevalent across malaria-endemic regions [30].

PfCRT and biochemical models of chloroquine resistance

Historically, investigations into the chloroquine resistance mechanism have generated vastly differing models, including reduced chloroquine influx, increased efflux, pH effects on drug accumulation and/or receptor availability, and glutathione degradation of hematin or chloroquine–hematin complexes [6,31]. From these studies, several tenets regarding the mode of action of chloroquine and the mechanism (or mechanisms) of resistance have become widely accepted: first, chloroquine enters the acidic digestive vacuole by passive diffusion as an uncharged species and becomes trapped in the digestive vacuole in its diprotonated, membrane-impermeable form; second, chloroquine is retained in the digestive vacuole as chloroquine2+–hematin complexes that are central to its antimalarial activity [32,33]; and third, the chloroquine resistance mechanism restricts chloroquine access to hematin and leads to reduced drug levels in the digestive vacuole [34]. The availability of isogenic lines expressing variant pfcrt alleles has also demonstrated that the chloroquine resistance mechanism is dependent on replacing the positively charged PfCRT K76 residue in the first transmembrane domain with a neutral residue (such as threonine, asparagine or isoleucine) [17,18,23].

Three models of chloroquine resistance that attempt to reconcile the existing data have now come to the forefront: (i) efflux of chloroquine out of the digestive vacuole via an energy-coupled transporter [3538]; (ii) leak of chloroquine out of the digestive vacuole down its concentration gradient in a manner that is not directly coupled to energy [20,24,34]; and (iii) pH-dependent reductions in chloroquine accumulation in the digestive vacuole, possibly associated with a role for PfCRT in mediating direct transport of the drug [33,3941] (Figure 2). Whereas the first two models are mutually exclusive in their interpretation of whether drug movement is directly coupled to energy, the digestive vacuole pH model is non-exclusive and could potentially combine with either of the two other models to yield chloroquine resistance. The idea that PfCRT is directly involved in chloroquine transport in the cell is consistent with both bioinformatic analyses that place this protein in the drug-metabolite effluxer family of transporters [42,43], and data from heterologous systems suggesting that mutant PfCRT can bind to and physically transport chloroquine [44,45].

Figure 2
Mechanistic models of PfCRT-mediated chloroquine resistance. (a) Chloroquine-sensitive parasites. In sensitive parasites expressing wild-type PfCRT, the weak base chloroquine (CQ) concentrates (broken arrow) in the digestive vacuole. The acidic environment ...

Energy-coupled chloroquine efflux

In support of the first model, CQR parasites have been reported to release pre-accumulated chloroquine almost 50 times faster than have CQS parasites [35]. Investigation of the kinetics of accumulation shows a transient, rapid increase in chloroquine accumulation in resistant parasites, which resolves to little or no accumulation within minutes. By contrast, there is a continuing rise in chloroquine accumulation in CQS parasites that reaches a plateau far higher than that attained in CQR parasites [46]. In all parasites, the initial chloroquine uptake is maximal at 37–40 °C, arguing that uptake is a temperature-dependent active process. Addition of glucose markedly stimulates chloroquine accumulation in CQS parasites, but reduces steady-state accumulation of chloroquine in CQR parasites, suggesting that there are energy-coupled mechanisms of chloroquine uptake and chloroquine efflux in sensitive and resistant parasites, respectively [36,38].

Preloading CQR parasites with different concentrations of unlabeled chloroquine before adding [3H]chloroquine has recently provided intriguing evidence of a ‘trans-stimulation’ effect, whereby accumulation of [3H]chloroquine first increases at low external preloaded chloroquine concentrations and then decreases [38]. This trans-stimulation phenotype has been previously described for the human red blood cell GLUT1 transporter and is thought to distinguish active efflux carriers from channels, pores or leaks, through which solutes move by passive diffusion [47]. This effect in CQR lines is not seen in CQS lines, where external preloaded chloroquine seems to compete with [3H]chloroquine for carrier sites at all concentrations. This observation has led to the proposal that CQR parasites have an active chloroquine efflux carrier, such that pre-equilibrated, preloaded chloroquine at low concentrations competes for carrier sites, leading to an increase in [3H]chloroquine accumulation, whereas preloaded chloroquine at high concentrations saturates the carrier sites and outcompetes [3H]chloroquine in binding to its intracellular receptor (heme), leading to a reduction in [3H]chloroquine accumulation [38]. This trans-stimulated accumulation of chloroquine by CQR parasites is glucose dependent (implying that ATP is involved), is blocked by verapamil, and also occurs when preloaded chloroquine is substituted by related quinoline drugs such as amodiaquine, quinine and quinidine [37]. Recent studies with pfcrt-modified recombinant parasites have indicated that mutant PfCRT might fulfill this role of a substrate-specific, verapamil-reversible, ATP-dependent chloroquine efflux carrier [48].

Leak of charged chloroquine out of the digestive vacuole

The second model postulates that chloroquine leaves the digestive vacuole by a mechanism of facilitated diffusion driven by a large concentration gradient of the protonated forms of the drug. Here, energy is proposed to drive the digestive vacuole proton pump and to maintain the concentration gradient of protonated chloroquine, rather than being directly coupled to drug movement.

Features of this model were first proposed by Warhurst et al. [49], who indicated some similarities between PfCRT and bacterial ClC chloride channels and noted that the crucial K76T mutation removes a positively charged residue from transmembrane domain I in a putative pore, increasing the hydrophobicity and potentially providing a route for the more polar (protonated) forms of chloroquine to escape the digestive vacuole. More hydrophobic chemo-sensitizers such as verapamil have been proposed to block chloroquine efflux sterically, effectively countering the resistance mechanism [49]. Support for the importance of the charge-loss mutation has come from the demonstration that a compensatory charge substitution (S163R in transmembrane domain IV) fully restores chloroquine sensitivity in drug-pressured mutant pfcrt lines [24].

Both the charged drug leak and the energy-coupled efflux models posit that PfCRT is directly involved in chloroquine movement but with the following key difference: in the former protonated chloroquine passively leaks out of the digestive vacuole through mutant PfCRT, whereas in the latter this protein actively effluxes drug. Both models can adequately explain most of the existing chloroquine transport data, and direct comparisons of the different experimental conditions with the same isogenic pfcrt-modified lines are warranted to resolve the differences between these models.

pH-dependent physiological changes at the digestive vacuole membrane

Early models of chloroquine accumulation postulated that uptake of chloroquine into the acidic digestive vacuole was primarily due to weak-base ion-trapping in accordance with the Henderson–Hasselbach equation [5052]. These models predicted that a more acidic pH in the digestive vacuole, leading to a larger pH gradient across the digestive vacuole membrane, would result in greater accumulation of chloroquine. Innovative developments in P. falciparum single cell photometry, however, have produced the surprising result that CQR and not CQS lines have the most acidic digestive vacuoles [39,40]. One explanation is that the lower digestive vacuole pH in CQR isolates might accelerate the rate of heme aggregation and hemozoin formation, reducing the amount of hematin available to bind chloroquine and resulting in less uptake of chloroquine [41]. Some concerns have been expressed about the appropriateness of using fluorescent pH sensor dyes and the possibility of photobleaching and laser-induced disaggregation of the digestive vacuole [5355]. Nevertheless, independent studies in pfcrt-transfected Dictyostelium discoideum also provide evidence that pfcrt expression has an effect on intracellular pH and that mutant PfCRT confers a more acidic pH than does the wild-type protein [45].

Notwithstanding, it is generally thought that differences in the digestive vacuole pH are themselves not primarily responsible for CQR, and recent data from the proponents of the pH model have provided evidence that physical interactions with chloroquine are also a factor in how mutant PfCRT mediates resistance [44].


More than a decade before the discovery of pfcrt, research into the genetic basis of chloroquine resistance had focused on pfmdr1, a P. falciparum ortholog of mammalian P-glycoproteins that mediate verapamil-reversible multi-drug resistance in mammalian cancer cells [35]. pfmdr1 encodes a 162-kDa protein (PfMDR1; also known as Pgh1) that localizes to the digestive vacuole membrane and consists of two homologous halves, each with six predicted transmembrane domains and a conserved nucleotide-binding domain [56,57] (Figure 3). The transport function of PfMDR1 was evidenced by its complementation of a yeast strain defective for the STE6 transporter [58]. A recent study, using fluorescein derivatives that are widely used in surrogate assays of P-glycoprotein function, has provided intriguing evidence that PfMDR1 might function to import solutes, including some antimalarial drugs, into the digestive vacuole [59].

Figure 3
Predicted structure and genetic polymorphisms in P. falciparum multidrug resistance-1. (a) PfMDR1 has two homologous halves, each with six predicted transmembrane domains and a nucleotide-binding pocket [56]. The nucleotide-binding domains (NBD1 and NBD2; ...

Two pfmdr1 alleles have been identified in CQR field isolates: the K1 allele (containing the point mutation N86Y) and the 7G8 allele (containing Y184F, S1034C, N1042D and D1246Y). In vitro studies using field isolates or laboratory lines have identified a partial association between the N86Y mutation and chloroquine resistance [6063]. By contrast, a role for the 7G8-type-3 mutations has been harder to ascertain [6,57]. For these 3′ mutations, direct evidence was obtained in allelic exchange experiments, which showed that the mutations enhance the degree of in vitro chloroquine resistance, although they do not confer resistance to sensitive parasites [64]. Another study using similar genetic techniques, however, observed no effect of these 3′ mutations on chloroquine [65]. The interpretation that pfmdr1 mutations might enhance chloroquine resistance in some genetic backgrounds but are themselves insufficient to confer resistance has support from some clinical studies [61,62,66]. Alternatively, the increased frequency of pfmdr1 polymorphisms in CQR parasites might reflect physiological compensation for the altered function of mutant pfcrt [5].

Studies of field isolates and a genetic cross have also identified an association between pfmdr1 point mutations and the degree of parasite in vitro susceptibility to other antimalarial drugs including mefloquine, halofantrine, quinine and artemisinin [57]. These data have been largely confirmed by genetic studies, which reinforce the notion that these effects are strain dependent [64,65]. Their impact on therapeutic outcomes seems more limited [67]. Nevertheless, a recent study that monitored recrudescence of infection after treatment with a lumefantrine–artemether combination (Coartem) has reported significant selection for the pfmdr1 86N polymorphism, suggesting that this mutation might be useful as a molecular marker of lumefantrine resistance [68].

Several investigations have noted a correlation between pfmdr1 expression and drug resistance in P. falciparum, paralleling the multidrug resistance mechanism observed in mammalian tumor cells. In a study based on drug-sensitive strains, pfmdr1 transcript levels were observed to increase after treatment with chloroquine, mefloquine and quinine, but not after treatment with pyrimethamine, suggesting that induction of pfmdr1 might be a drug-specific mechanism of resistance [69]. Other studies with field isolates or drug-pressured laboratory lines have found an association between in vitro mefloquine resistance and a higher pfmdr1 copy number, which in some instances is also associated with increased susceptibility to chloroquine [57,67,70,71].

The relationship between pfmdr1 copy number and mefloquine treatment outcome has been comprehensively investigated in a large, prospective study in Thailand, where quantitative real-time PCR analysis of over 600 patient samples has shown that pfmdr1 amplification is highly associated with failure of mefloquine or mefloquine–artesunate treatment [67]. pfmdr1 amplification has also been associated with an increased risk of failure of short-term artemether–lumefantrine treatment [72]. These clinical findings have been confirmed by a genetically controlled experiment showing that an increase in pfmdr1 copy number causes a decrease in in vitro susceptibility to mefloquine, quinine, halofantrine and artemisinin [73]. Amplified pfmdr1 copy number, which is prevalent mostly in Asia, has also been observed in some earlier field isolates from Gabon, possibly because of local drug pressure [74]. How PfMDR1 mediates these effects is unclear, although a recent study indicates that pfmdr1 point mutations or changes in pfmdr1 copy number might affect the degree of drug accumulation in the digestive vacuole, which could affect the in vitro potency of the drugs [59].

Other transporters implicated in antimalarial drug resistance

Recent studies in antimalarial chemotherapy have also implicated several other transporters, most notably PfATP6 – the P. falciparum ortholog of the mammalian sarcoendoplasmic reticulum Ca2+ ATPase (SERCA). Expression of PfATP6 in Xenopus laevis oocytes revealed that its ATPase activity is inhibited by artemisinin in addition to thapsigargin – a known SERCA inhibitor [10]. Modeling of this protein against mammalian SERCA led to the finding that amino acid variants at position 263 in the predicted thapsigargin-binding pocket can ablate inhibition by artemisinin derivatives in the Xenopus oocyte system [12]. Recent studies have also reported an association between the S769N mutation in PfATP6 and increased IC50 values for artemether in field isolates from French Guyana [11]. Genetic experiments in the parasite should further define the role of PfATP6 in the mode of action of artemisinin derivatives.

Recent genomic analyses have implicated other putative transporters in modulating parasite response to antimalarial drugs [63,75,76]. For example, quantitative trait loci mapping of a P. falciparum genetic cross in which inheritance of chloroquine and quinine resistance are correlated found an association between quinine resistance and mutations in pfcrt and pfmdr1, and also implicated a locus on chromosome 13 that contains a predicted Na+–H+ exchanger (pfnhe) [75]. Analysis of microsatellite variations noted a significant association between DNNND repeats in the C-terminal cytoplasmic domain of PfNHE and in vitro quinine response. In addition, reduction of pfnhe expression by genetic manipulation has recently identified an association between PfNHE expression levels and the degree of quinine resistance in CQR parasites (L. Nkrumah et al., unpublished).

Analyses of single nucleotide polymorphisms in different parasite isolates have also identified additional candidates for antimalarial resistance genes. Linkage disequilibrium analysis of 97 culture-adapted parasite isolates from around the world found that single nucleotide polymorphisms from pfcrt, pfmdr1 and at least nine new putative transporter genes located on different chromosomes were associated with chloroquine and quinine resistance [63]. However, a follow-up study examining polymorphisms in these nine new genes in isolates from two independent population samples in Southeast Asia found only one consistent association, which was between increased artesunate IC50 values and the putative ABC transporter G7 [76]. These discrepancies might be due to variations in study design or to the different geographic origins of the parasite samples.

Concluding remarks

The above studies provide a promising platform from which to direct future research on parasite transporter proteins and drug resistance. In particular, elucidation of the mechanisms by which PfCRT and PfMDR1 mediate resistance to multiple drug classes can help to guide efforts to overcome the spread of drug resistance. Furthermore, screens for other candidate transporter loci involved in antimalarial drug resistance should be extended to assess copy number and/or expression levels, in addition to identifying single nucleotide polymorphisms. A thorough understanding of the complex interactions among malarial transport proteins, and how these interactions influence parasite response to antimalarial drugs, will be an essential tool in worldwide efforts to combat and control this important disease.


We thank Rebecca Muhle, Marcus Lee, Amar bir Singh Sidhu, Patrick Bray and Scott Bohle for comments on the manuscript. Financial support was provided by the National Institutes of Health (R01 AI50234) and a Burroughs Wellcome Fund Investigator in Pathogenesis of Infectious Disease Award (to D.A.F.).


1. Greenwood B, Mutabingwa T. Malaria in 2002. Nature. 2002;415:670–672. [PubMed]
2. Snow RW, et al. The global distribution of clinical episodes of Plasmodium falciparum malaria. Nature. 2005;434:214–217. [PMC free article] [PubMed]
3. Miller LH, et al. The pathogenic basis of malaria. Nature. 2002;415:673–679. [PubMed]
4. Planche T, Krishna S. Severe malaria: metabolic complications. Curr Mol Med. 2006;6:141–153. [PubMed]
5. Wellems TE, Plowe CV. Chloroquine-resistant malaria. J Infect Dis. 2001;184:770–776. [PubMed]
6. Uhlemann A-C, et al. Mechanisms of antimalarial drug action and resistance. In: Sherman IW, editor. Molecular Approaches to Malaria. ASM Press; 2005. pp. 429–461.
7. Wongsrichanalai C, et al. Epidemiology of drug-resistant malaria. Lancet Infect Dis. 2002;2:209–218. [PubMed]
8. White NJ. Antimalarial drug resistance. J Clin Invest. 2004;113:1084–1092. [PMC free article] [PubMed]
9. Baird JK. Effectiveness of antimalarial drugs. N Engl J Med. 2005;352:1565–1577. [PubMed]
10. Eckstein-Ludwig U, et al. Artemisinins target the SERCA of Plasmodium falciparum. Nature. 2003;424:957–961. [PubMed]
11. Jambou R, et al. Resistance of Plasmodium falciparum field isolates to in-vitro artemether and point mutations of the SERCA-type PfATPase6. Lancet. 2005;366:1960–1963. [PubMed]
12. Uhlemann AC, et al. A single amino acid residue can determine sensitivity of SERCAs to artemisinins. Nat Struct Mol Biol. 2005;12:628–629. [PubMed]
13. Krishna S, et al. Re-evaluation of how artemisinins work in light of emerging evidence of in vitro resistance. Trends Mol Med. 2006;12:200–205. [PMC free article] [PubMed]
14. Gregson A, Plowe CV. Mechanisms of resistance of malaria parasites to antifolates. Pharmacol Rev. 2005;57:117–145. [PubMed]
15. Wiesner J, et al. New antimalarial drugs. Angew Chem Int Ed Engl. 2003;42:5274–5293. [PubMed]
16. Su X, et al. Complex polymorphisms in a ~330 kDa protein are linked to chloroquine-resistant P. falciparum in Southeast Asia and Africa. Cell. 1997;91:593–603. [PubMed]
17. Fidock DA, et al. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol Cell. 2000;6:861–871. [PMC free article] [PubMed]
18. Cooper RA, et al. Alternative mutations at position 76 of the vacuolar transmembrane protein PfCRT are associated with chloroquine resistance and unique stereospecific quinine and quinidine responses in Plasmodium falciparum. Mol Pharmacol. 2002;61:35–42. [PubMed]
19. Goldberg DE. Hemoglobin degradation. Curr Top Microbiol Immunol. 2005;295:275–291. [PubMed]
20. Bray PG, et al. Defining the role of PfCRT in Plasmodium falciparum chloroquine resistance. Mol Microbiol. 2005;56:323–333. [PubMed]
21. Hyde JE. Drug-resistant malaria. Trends Parasitol. 2005;21:494–498. [PMC free article] [PubMed]
22. Sidhu AB, et al. Chloroquine resistance in Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science. 2002;298:210–213. [PMC free article] [PubMed]
23. Lakshmanan V, et al. A critical role for PfCRT K76T in Plasmodium falciparum verapamil-reversible chloroquine resistance. EMBO J. 2005;24:2294–2305. [PMC free article] [PubMed]
24. Johnson DJ, et al. Evidence for a central role for PfCRT in conferring Plasmodium falciparum resistance to diverse antimalarial agents. Mol Cell. 2004;15:867–877. [PMC free article] [PubMed]
25. Chen N, et al. pfcrt allelic types with two novel amino acid mutations in chloroquine-resistant Plasmodium falciparum isolates from the Philippines. Antimicrob Agents Chemother. 2003;47:3500–3505. [PMC free article] [PubMed]
26. Lim P, et al. pfcrt polymorphism and chloroquine resistance in Plasmodium falciparum strains isolated in Cambodia. Antimicrob Agents Chemother. 2003;47:87–94. [PMC free article] [PubMed]
27. Sutherland CJ, et al. Gambian children successfully treated with chloroquine can harbour and transmit Plasmodium falciparum gametocytes carrying resistance genes. Am J Trop Med Hyg. 2002;67:578–585. [PubMed]
28. Hallett RL, et al. Combination therapy counteracts the enhanced transmission of drug-resistant malaria parasites to mosquitoes. Antimicrob Agents Chemother. 2004;48:3940–3943. [PMC free article] [PubMed]
29. Chutmongkonkul M, et al. Plasmodium falciparum: effect of chloroquine, halofantrine and pyrimethamine on the infectivity of gametocytes for Anopheles stephensi mosquitoes. Ann Trop Med Parasitol. 1992;86:103–110. [PubMed]
30. Wootton JC, et al. Genetic diversity and chloroquine selective sweeps in Plasmodium falciparum. Nature. 2002;418:320–323. [PubMed]
31. Bray PG, Ward SA. A comparison of the phenomenology and genetics of multidrug resistance in cancer cells and quinoline resistance in Plasmodium falciparum. Pharmacol Ther. 1998;77:1–28. [PubMed]
32. Bray PG, et al. Defining the role of PfCRT in P. falciparum chloroquine resistance. Mol Microbiol. 2005;56:323–333. [PubMed]
33. Yayon A. The antimalarial mode of action of chloroquine. Rev Clin Basic Pharm. 1985;5:99–139. [PubMed]
34. Bray PG, et al. Access to hematin: the basis of chloroquine resistance. Mol Pharmacol. 1998;54:170–179. [PubMed]
35. Krogstad DJ, et al. Efflux of chloroquine from Plasmodium falciparum: mechanism of chloroquine resistance. Science. 1987;238:1283–1285. [PubMed]
36. Krogstad DJ, et al. Energy dependence of chloroquine accumulation and chloroquine efflux in Plasmodium falciparum. Biochem Pharmacol. 1992;43:57–62. [PubMed]
37. Sanchez CP, et al. Evidence for a substrate specific and inhibitable drug efflux system in chloroquine resistant Plasmodium falciparum strains. Biochemistry. 2004;43:16365–16373. [PubMed]
38. Sanchez CP, et al. Trans stimulation provides evidence for a drug efflux carrier as the mechanism of chloroquine resistance in Plasmodium falciparum. Biochemistry. 2003;42:9383–9394. [PubMed]
39. Bennett TN, et al. Drug resistance-associated pfCRT mutations confer decreased Plasmodium falciparum digestive vacuolar pH. Mol Biochem Parasitol. 2004;133:99–114. [PubMed]
40. Dzekunov SM, et al. Digestive vacuolar pH of intact intraerythrocytic P. falciparum either sensitive or resistant to chloroquine. Mol Biochem Parasitol. 2000;110:107–124. [PubMed]
41. Ursos LM, Roepe PD. Chloroquine resistance in the malarial parasite, Plasmodium falciparum. Med Res Rev. 2002;22:465–491. [PubMed]
42. 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. [PubMed]
43. Tran CV, Saier MH., Jr The principal chloroquine resistance protein of Plasmodium falciparum is a member of the drug/metabolite transporter superfamily. Microbiology. 2004;150:1–3. [PubMed]
44. Zhang H, et al. The antimalarial drug resistance protein Plasmodium falciparum chloroquine resistance transporter binds chloroquine. Biochemistry. 2004;43:8290–8296. [PubMed]
45. Naude B, et al. Dictyostelium discoideum expresses a malaria chloroquine resistance mechanism upon transfection with mutant, but not wild-type, Plasmodium falciparum transporter PfCRT. J Biol Chem. 2005;280:25596–25603. [PMC free article] [PubMed]
46. Sanchez CP, et al. Identification of a chloroquine importer in Plasmodium falciparum. Differences in import kinetics are genetically linked with the chloroquine-resistant phenotype. J Biol Chem. 1997;272:2652–2658. [PubMed]
47. Stein WD. Transport and Diffusion Across Cell Membranes. Academic Press; 1986.
48. Sanchez CP, et al. Evidence for a pfcrt-associated chloroquine efflux system in the human malarial parasite Plasmodium falciparum. Biochemistry. 2005;44:9862–9870. [PubMed]
49. Warhurst DC, et al. Lysosomes and drug resistance in malaria. Lancet. 2002;360:1527–1529. [PubMed]
50. Homewood CA, et al. Lysosomes, pH and the anti-malarial action of chloroquine. Nature. 1972;235:50–52. [PubMed]
51. Krogstad DJ, et al. Antimalarials increase vesicle pH in Plasmodium falciparum. J Cell Biol. 1985;101:2302–2309. [PMC free article] [PubMed]
52. Yayon A, et al. Identification of the acidic compartment of Plasmodium falciparum-infected human erythrocytes as the target of the antimalarial drug chloroquine. EMBO J. 1984;3:2695–2700. [PMC free article] [PubMed]
53. Bray PG, et al. Distribution of acridine orange fluorescence in Plasmodium falciparum-infected erythrocytes and its implications for the evaluation of digestive vacuole pH. Mol Biochem Parasitol. 2002;119:301–304. [PubMed]
54. Wissing F, et al. Illumination of the malaria parasite Plasmodium falciparum alters intracellular pH. Implications for live cell imaging. J Biol Chem. 2002;277:37747–37755. [PubMed]
55. Hayward R, et al. The pH of the digestive vacuole of Plasmodium falciparum is not associated with chloroquine resistance. J Cell Sci. 2006;119:1016–1025. [PubMed]
56. Peel SA. The ABC transporter genes of Plasmodium falciparum and drug resistance. Drug Resist Updat. 2001;4:66–74. [PubMed]
57. Duraisingh MT, Cowman AF. Contribution of the pfmdr1 gene to antimalarial drug-resistance. Acta Trop. 2005;94:181–190. [PubMed]
58. Volkman SK, et al. Functional complementation of the ste6 gene of Saccharomyces cerevisiae with the pfmdr1 gene of Plasmodium falciparum. Proc Natl Acad Sci U S A. 1995;92:8921–8925. [PMC free article] [PubMed]
59. Rohrbach P, et al. Genetic linkage of pfmdr1 with food vacuolar solute import in Plasmodium falciparum. EMBO J. 2006;25:3000–3011. [PMC free article] [PubMed]
60. Adagut IS, Warhust DC. Plasmodium falciparum: linkage disequilibrium between loci in chromosomes 7 and 5 and chloroquine selective pressure in Northern Nigeria. Parasitology. 2001;123:219–224. [PubMed]
61. Djimdé A, et al. A molecular marker for chloroquine resistant falciparum malaria. N Engl J Med. 2001;344:257–263. [PubMed]
62. Babiker HA, et al. High-level chloroquine resistance in Sudanese isolates of Plasmodium falciparum is associated with mutations in the chloroquine resistance transporter gene pfcrt and the multidrug resistance gene pfmdr1. J Infect Dis. 2001;183:1535–1538. [PubMed]
63. Mu J, et al. Multiple transporters associated with malaria parasite responses to chloroquine and quinine. Mol Microbiol. 2003;49:977–989. [PubMed]
64. Reed MB, et al. Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature. 2000;403:906–909. [PubMed]
65. Sidhu AB, et al. pfmdr1 mutations contribute to quinine resistance and enhance mefloquine and artemisinin sensitivity in Plasmodium falciparum. Mol Microbiol. 2005;57:913–926. [PubMed]
66. Ngo T, et al. Analysis of pfcrt, pfmdr1, dhfr, and dhps mutations and drug sensitivities in Plasmodium falciparum isolates from patients in Vietnam before and after treatment with artemisinin. Am J Trop Med Hyg. 2003;68:350–356. [PubMed]
67. Price RN, et al. Mefloquine resistance in Plasmodiumfalciparum and increased pfmdr1 gene copy number. Lancet. 2004;364:438–447. [PubMed]
68. Sisowath C, et al. In vivo selection of Plasmodium falciparum pfmdr1 86N coding alleles by artemether-lumefantrine (Coartem) J Infect Dis. 2005;191:1014–1017. [PubMed]
69. Myrick A, et al. Mapping of the Plasmodium falciparum multidrug resistance gene 5′-upstream region, and evidence of induction of transcript levels by antimalarial drugs in chloroquine sensitive parasites. Mol Microbiol. 2003;49:671–683. [PubMed]
70. Nelson AL, et al. pfmdr1 genotyping and in vivo mefloquine resistance on the Thai-Myanmar border. Am J Trop Med Hyg. 2005;72:586–592. [PubMed]
71. Price RN, et al. The pfmdr1 gene is associated with a multidrug-resistant phenotype in Plasmodium falciparum from the western border of Thailand. Antimicrob Agents Chemother. 1999;43:2943–2949. [PMC free article] [PubMed]
72. Price RN, et al. Molecular and pharmacological determinants of the therapeutic response to artemether–lumefantrine in multidrug-resistant Plasmodium falciparum malaria. Clin Infect Dis. 2006;42:1570–1577. [PubMed]
73. Sidhu ABS, et al. Decreasing pfmdr1 copy number in Plasmodium falciparum malaria heightens susceptibility to mefloquine, lumefantrine, halofantrine, quinine, and artemisinin. J Infect Dis. 2006;194:528–535. [PMC free article] [PubMed]
74. Uhlemann AC, et al. Amplification of Plasmodium falciparum multidrug resistance gene 1 in isolates from Gabon. J Infect Dis. 2005;192:1830–1835. [PubMed]
75. Ferdig MT, et al. Dissecting the loci of low-level quinine resistance in malaria parasites. Mol Microbiol. 2004;52:985–997. [PubMed]
76. Anderson TJ, et al. Are transporter genes other than the chloroquine resistance locus (pfcrt) and multidrug resistance gene (pfmdr) associated with antimalarial drug resistance? Antimicrob Agents Chemother. 2005;49:2180–2188. [PMC free article] [PubMed]
77. Howard EM, et al. A novel transporter, pfcrt, confers antimalarial drug resistance. J Membr Biol. 2002;190:1–8. [PubMed]


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