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A critical role for PfCRT K76T in Plasmodium falciparum verapamil‐reversible chloroquine resistance

Viswanathan Lakshmanan, Patrick G Bray, Dominik Verdier‐Pinard, David J Johnson, Paul Horrocks, Rebecca A Muhle, George E Alakpa, Ruth H Hughes, Steve A Ward, Donald J Krogstad, Amar Bir Singh Sidhu, David A Fidock

Author Affiliations

  1. Viswanathan Lakshmanan1,
  2. Patrick G Bray*,2,
  3. Dominik Verdier‐Pinard1,
  4. David J Johnson1,
  5. Paul Horrocks3,
  6. Rebecca A Muhle1,
  7. George E Alakpa1,
  8. Ruth H Hughes2,
  9. Steve A Ward2,
  10. Donald J Krogstad4,
  11. Amar Bir Singh Sidhu1 and
  12. David A Fidock*,1
  1. 1 Department of Microbiology and Immunology, Albert Einstein College of Medicine, Bronx, NY, USA
  2. 2 Molecular and Biochemical Parasitology Group, Liverpool School of Tropical Medicine, Liverpool, UK
  3. 3 Weatherall Institute of Molecular Medicine, University of Oxford, UK
  4. 4 Department of Tropical Medicine, Tulane School of Public Health, New Orleans, LA, USA
  1. *Corresponding authors: Molecular and Biochemical Parasitology Group, Liverpool School of Tropical Medicine, Liverpool L3 5QA, UK. Tel.: +44 151 705 3119; Fax: +44 151 708 9007; E‐mail: p.g.bray{at}liv.ac.ukDepartment of Microbiology and Immunology, Albert Einstein College of Medicine, Forchheimer 403, 1300 Morris Park Avenue, Bronx, NY 10461, USA. Tel.: +1 718 430 3759; Fax: +1 718 430 8711; E‐mail: dfidock{at}aecom.yu.edu

Abstract

Chloroquine resistance (CQR) in Plasmodium falciparum is associated with mutations in the digestive vacuole transmembrane protein PfCRT. However, the contribution of individual pfcrt mutations has not been clarified and other genes have been postulated to play a substantial role. Using allelic exchange, we show that removal of the single PfCRT amino‐acid change K76T from resistant strains leads to wild‐type levels of CQ susceptibility, increased binding of CQ to its target ferriprotoporphyrin IX in the digestive vacuole and loss of verapamil reversibility of CQ and quinine resistance. Our data also indicate that PfCRT mutations preceding residue 76 modulate the degree of verapamil reversibility in CQ‐resistant lines. The K76T mutation accounts for earlier observations that CQR can be overcome by subtly altering the CQ side‐chain length. Together, these findings establish PfCRT K76T as a critical component of CQR and suggest that CQ access to ferriprotoporphyrin IX is determined by drug–protein interactions involving this mutant residue.

Introduction

Chloroquine (CQ) was for decades the gold standard for prevention and treatment of uncomplicated Plasmodium falciparum malaria. However, the emergence and spread of CQ resistance (CQR), first detected in Columbia and Thailand and moving later into Africa, has contributed to a deteriorating malaria situation worldwide (Trape et al, 2002).

Key to the mode of CQ action is the process of hemoglobin degradation, which liberates toxic ferriprotoporphyrin IX (FP) moieties in the intracellular parasite's digestive vacuole (DV). These moieties are detoxified via incorporation into inert hemozoin polymers consisting of FP dimers (Pagola et al, 2000). CQ, a weak base, inhibits detoxification by accumulating in the DV (as CQ2+) and binding to FP, causing parasite death presumably as a consequence of increased membrane permeability and lipid peroxidation (Waller et al, 2004).

CQ‐resistant parasites exhibit elevated CQ IC50 values, reduced CQ accumulation and partial reversibility of resistance by the calcium channel blocker verapamil (VP) (Krogstad et al, 1987; Martin et al, 1987). Yet the biochemical mechanism of CQR has remained enigmatic, with theories espousing reduced CQ access to FP via leak of charged drug, carrier‐mediated drug efflux, reduced influx, altered CQ partitioning or heme turnover rates resulting from pH gradient changes, heme binding proteins or altered glutathione levels (Wellems and Plowe, 2001; Ursos and Roepe, 2002; Johnson et al, 2004; Sanchez et al, 2004).

A genetic cross between a CQ‐resistant line (Dd2, Indochina) and a CQ‐sensitive line (HB3, Honduras) earlier revealed a tight association between inheritance of VP‐reversible CQR and polymorphisms in the DV transmembrane protein PfCRT (Plasmodium falciparum chloroquine resistance transporter) (Fidock et al, 2000). Variant pfcrt alleles, encoding distinct haplotypes yet sharing a common K76T mutation, have been associated with in vitro CQR in lines from Asia, Africa and South America (reviewed in Bray et al, 2005). This correlation however did not extend to all field‐based studies (Thomas et al, 2002; Lim et al, 2003). This might reflect the involvement of additional genes potentially including pfmdr1 (Foote et al, 1989; Wilson et al, 1989; Reed et al, 2000), undetected pfcrt polymorphisms or technical caveats associated with performing one‐time, field‐based drug susceptibility assays on non‐culture‐adapted, frequently polyclonal patient isolates. A key role for pfcrt was reported by Sidhu et al (2002), who demonstrated acquisition of in vitro CQR by an originally CQ‐sensitive line (GC03) engineered to express mutant pfcrt (from the CQ‐resistant lines 7G8 (Brazil) and Dd2).

In vivo studies have also reported a significantly increased risk of CQ treatment failure in patients with P. falciparum infections carrying pfcrt K76T alleles (Wellems and Plowe, 2001; Basco et al, 2002; Ochong et al, 2003). This risk was highest in very young children, in whom acquired immunity is thought to have little influence on clinical outcome (Djimdé et al, 2001). Some patients carrying pfcrt K76T infections however were observed to respond clinically to CQ, fueling debate on the relative contributions of immunity and pfcrt (Chen et al, 2002; Happi et al, 2003). Here, we have further investigated the contribution of PfCRT K76T by transfection and characterization of recombinant parasites.

Results

Targeting pfcrt codon 76 by allelic exchange

Replacement of the mutant pfcrt T76 codon with the wild‐type K76 codon, in CQ‐resistant parasites harboring pfcrt alleles representative of Old World (Dd2) or New World (7G8) origins, was achieved by allelic exchange. The strategy was based on homologous recombination with the transfection plasmid pK76, effectively replacing endogenous pfcrt with a recombinant locus consisting of a downstream, full‐length functional pfcrt and an upstream, truncated nonfunctional pfcrt remnant (Figure 1A). Crossover events downstream of codon 76 introduced a T76K substitution in the full‐length functional pfcrt, generating a ‘back‐mutant’ line (Table I and Figure 1A). Crossover events upstream of codon 76 generated the same recombinant locus without altering the haplotype, thus providing recombinant controls.

Figure 1.

Allelic exchange strategy and molecular characterization of recombinant clones. (A) Schematic depicting integration of the pK76 plasmid into the endogenous pfcrt locus by homologous recombination and single‐site crossover. The recombinant downstream locus contained a full‐length functional pfcrt gene, transcriptionally controlled by a shortened pfcrt promoter and the endogenous 3′UTR. A truncated pfcrt remnant (with the first 5 of 13 exons), the bsd selectable marker and pBluescript (pBS) sequence were located upstream. Square brackets delineate the plasmid sequence that could integrate as multiple tandem copies (n). Fragments obtained upon restriction digestion, and pfcrt or bsd probe locations, are indicated. B, BglII; S, StuI; X, XbaI. (B) PCR detection of recombinant functional (downstream) pfcrt with primers p3/p4 (top panel), the truncated (upstream) remnant with primers p5/p6 (middle panel) or endogenous pfcrt with primers p5/p4 (bottom panel). Lanes: 1: Dd2; 2: T76K‐1Dd2; 3: C‐1Dd2; 4: 7G8; 5: T76K‐17G8; 6: C‐17G8; 7: TMD1‐1; M: 1 kb DNA ladder. Lane identities are maintained throughout the figure. (C, D) Southern hybridization of genomic DNA digested with BglII/StuI/XbaI (C) or BglII/StuI (D), and probed with pfcrt or bsd. (E) RT–PCR assays with primers p7/p9 revealed a 1.1 kb transcription product from the functional downstream locus (recombinant lines) and the endogenous locus (parental lines). Primers p7/p6 gave a 1.0 kb transcription product from the upstream truncated pfcrt remnant (in the recombinant but not parental lines). (F) Northern hybridization of total RNA probed with pfcrt or ef‐1α. The RNA gel used for blotting shows equivalent loading. Data shown are representative of four separate Northern analyses performed on synchronized parasites. (G) Western blot of protein samples from recombinant and parental lines probed with antibodies to PfCRT or the Golgi marker PfERD2. Total protein amounts were loaded in two‐fold dilutions, with three sets of dilutions per line (see Supplementary data).

View this table:
Table 1. PfCRT haplotype of recombinant and parental lines

Minimal promoter elements were first defined that could drive transcription of recombinant full‐length pfcrt. For this, we amplified the 3.0 kb region separating pfcrt from the upstream cg3 gene. Unidirectional digestion produced sequences stretching 1.6, 1.2 and 1.0 kb upstream from the pfcrt start codon. These elements gave mean±s.e.m. luciferase activities of 84±12, 60±13 and 3±2%, respectively, relative to the 3.0 kb element (n=4 assays). Our allelic exchange strategy employed the 1.6 kb element, which gave acceptable transcription activity while retaining favorable odds that a recombination event would occur downstream of codon 76.

To generate the desired recombinant clones, we transfected Dd2 and 7G8 parasites with pK76 (Figure 1A). Parasite lines that underwent integration into pfcrt were identified by PCR and sequencing, and cloned. Recombinant back‐mutants were named T76K‐1Dd2, T76K‐2Dd2, T76K‐17G8 and T76K‐27G8, based on their genetic background (Table I). Recombinant controls were named C‐1Dd2, C‐2Dd2, C‐17G8 and C‐27G8. PCR with primers p3/p4 and p5/p6 yielded, for all recombinant lines, 3.4 and 3.3 kb bands corresponding to functional and truncated pfcrt sequences, respectively (Figure 1B). Sequencing of nested PCR products confirmed their PfCRT haplotypes. Primers p5/p4, specific for endogenous pfcrt, produced a 3.4 kb band from parental (nontransfected) lines and episomally transfected cultures but not from the cloned recombinants (Figure 1B; data not shown).

Allelic replacement was confirmed by Southern hybridization. Upon BglII/StuI/XbaI digestion, all recombinant lines displayed 7.8, 7.4 and 4.8 kb bands when hybridized with a pfcrt probe (exon 2–intron 3), and 7.8 and 7.4 kb bands with a blasticidin S‐deaminase (bsd) probe (Figure 1C). Parental lines produced a 5.2 kb band. StuI/BglII digestion, which cleaves sites flanking pfcrt (Figure 1A), revealed tandem integration of two or more plasmid copies in all recombinant lines (Figure 1D).

Analysis of pfcrt expression in cloned recombinant lines

RT–PCR with primers p7/p9 showed transcription from the full‐length pfcrt locus in all lines (Figure 1E). In the recombinants, transcription was also detected with primers p7/p6 from the upstream remnant, which retained the endogenous promoter but was truncated midway through pfcrt (Figure 1E). Northern blot hybridizations revealed a 3.5 kb transcript emanating from the functional locus in all recombinant lines, compared to a 4.2 kb transcript from parental Dd2 and 7G8 (Figure 1F). Based on earlier mapping (Waller et al, 2003), this indicated that recombinant downstream pfcrt transcripts originated close to the start of the 1.6 kb element. Hybridizations with ef‐1α revealed similar loading in all lanes. Densitometry predicted a 65–75% decrease in steady‐state transcript levels of recombinant functional pfcrt compared to the parental locus. Western blots of highly synchronized parasite protein extracts, probed with antibodies to either PfCRT or PfERD2 (loading control), predicted a 40–55% decrease in expression levels in back‐mutant and control recombinants, compared to parental lines (Figure 1G). This greater reduction in mRNA compared to protein levels upon pfcrt allelic exchange was also observed previously (Waller et al, 2003).

Removal of the PfCRT K76T mutation abolishes CQR and VP reversibility of CQ and quinine resistance

Generating the back‐mutant lines enabled us to determine the contribution of K76T to P. falciparum susceptibility to antimalarial agents. Assays were performed against CQ and its in vivo metabolite mono‐desethylchloroquine (m‐dCQ), quinine (QN) and its diastereoisomer quinidine (QD), amodiaquine (ADQ), mefloquine (MFQ), artemisinin (ART) and amantadine (AMT). CQ, m‐dCQ and QN were all tested with or without VP (Figures 2 and 3). For each line, 3–14 independent assays were performed in duplicate (detailed after in Supplementary Table I).

Figure 2.

Antimalarial susceptibility profiles of pfcrt‐modified lines on the Dd2 genetic background, showing a graphical representation of IC50 values (mean±s.e.m.) (values indicated in Supplementary Table I). Note that the m‐dCQ graphs have a two‐segment Y‐axis to adequately represent the range of values. For graphs with left and right Y‐axes (with different scales), the left side (blacks bars) corresponds to QD or ADQ, whereas the right side (gray bars) corresponds to MFQ or AMT. Values are expressed in nM for all drugs, except AMT for which the unit is μM. VP was included at 0.8 μM. Each mean value was calculated from 3–14 assays performed in duplicate. Determinations of statistical significance used unpaired two‐tailed t‐tests, with the P‐value reporting the lesser of the significant values obtained when comparing back‐mutant lines against each of the two control recombinant lines. *P<0.05; **P<0.01; ***P<0.001. The haplotypes of all lines are listed in Table I.

Figure 3.

Antimalarial susceptibility profiles of pfcrt‐modified lines on the 7G8 genetic background. Data are presented as described in Figure 2 legend.

For the Dd2 lineage, the T76K substitution resulted in a total loss of CQR. Mean CQ IC50 values were 22.1 and 25.7 nM in the back‐mutants T76K‐1Dd2 and T76K‐2Dd2, a degree of sensitivity equivalent to the reference CQ‐sensitive line GC03 (22.8 nM; Supplementary Table I). These values were six‐ to seven‐fold lower than those of C‐1Dd2 and C‐2Dd2 controls (P<0.01) and 11‐ to 13‐fold lower than Dd2. Loss of resistance was even more dramatic with m‐dCQ, with T76K‐1Dd2 and T76K‐2Dd2 demonstrating IC50 values of 23.8 and 30.8 nM, directly comparable to GC03 (35.3 nM). These back‐mutant m‐dCQ values were 24‐ to 38‐fold lower than recombinant controls (P<0.001) and 48‐ to 62‐fold lower than Dd2. We note that the control lines C‐1Dd2 and C‐2Dd2 (which underexpress PfCRT by about half; Figure 1G) had CQ and m‐dCQ IC50 values that were approximately two‐fold lower than Dd2. This confirmed an earlier observation that reduced PfCRT expression levels can lower the degree of CQR (Waller et al, 2003).

Interestingly, there was a statistically significant, approximately 40% decrease in IC50 values for QN in the Dd2 recombinant control and back‐mutant lines (which all gave similar IC50 values), compared to Dd2 (Figure 2). Smaller decreases in IC50 values were observed for QD, MFQ and ADQ in the Dd2 back‐mutants; however, these were not significant.

We also assayed AMT, in view of recent reports that CQ‐resistant parasites are hypersensitive to this influenza A M2 ion channel inhibitor (Johnson et al, 2004). Dd2 and recombinant control lines showed mean IC50 values of 20–29 μM (Figure 2). In comparison, the CQ‐sensitive Dd2 back‐mutants became 11‐ to 17‐fold less susceptible (P<0.001; mean IC50 values of 309–331 μM). Thus, in Dd2 parasites, the PfCRT K76T mutation appears to significantly influence AMT susceptibility. We note that in a previous study, in vitro AMT pressure of the K1 (Thailand) CQ‐resistant line resulted in an AMT‐resistant mutant that had also lost VP‐reversible CQR and had acquired two novel pfcrt point mutations (T152A and S163R) (Johnson et al, 2004). These data suggest that various mutations in Dd2‐like pfcrt alleles can confer parasite resistance to AMT.

For the 7G8 lineage, a total loss of CQR was also observed in the back‐mutants. CQ IC50 values were 23.9 and 25.5 nM for T76K‐17G8 and T76K‐27G8, respectively (Supplementary Table I), which were four‐ to five‐fold lower than C‐17G8 and C‐27G8 (P<0.001), and 10‐ to 11‐fold lower than 7G8 (Figure 3). The CQ‐sensitive line 3D7 gave a CQ IC50 value of 25.2 nM. Again, the loss was greater with m‐dCQ, with T76K‐17G8 and T76K‐27G8 showing IC50 values comparable to 3D7 (29–42 nM), 11‐ to 13‐fold lower than C‐17G8 and C‐27G8 (P<0.01) and 28‐ to 29‐fold lower than 7G8.

QN assays revealed a statistically significant two‐fold decrease in IC50 values in the 7G8 back‐mutant lines compared to the controls (P<0.001; Figure 3). Thus, for 7G8 (but not Dd2), K76T appeared to contribute to QN resistance. These different QN results with Dd2 and 7G8 support the idea that pfcrt is but one component of a multifactorial basis of QN resistance (Reed et al, 2000; Sidhu et al, 2002; Ferdig et al, 2004), and that the extent of its contribution differs between strains. For our 7G8 lines, no significant differences were observed for QD, ADQ or MFQ. For AMT, all 7G8 lines displayed high IC50 values (129–213 μM), irrespective of pfcrt codon 76. Thus for 7G8 (in contrast to Dd2), the AMT response appears to be governed either by pfcrt polymorphisms not tested herein or by separate genes.

These assays also revealed a total loss of VP reversibility of CQ and m‐dCQ resistance in all Dd2 and 7G8 back‐mutant lines (P<0.01; Figures 2 and 3 and Supplementary Table I), implicating residue 76 as a key component of the VP‐reversible CQR phenotype in these two geographically distinct parasites. Recombinant control lines retained similar degrees of VP reversibility as parental, nontransfected lines.

Arguably, loss of CQR would inevitably lead to loss of VP reversibility, if there were no longer a mechanism to reverse in CQ‐sensitive parasites. Thus, to separate VP reversibility from drug resistance per se, we took advantage of the highly VP‐reversible, QN resistance phenotype observed in the Dd2 back‐mutants. VP reversibility was quantified using the ‘Response Modification Index’ (RMI: defined as the ratio of the IC50 in the presence of VP to that in VP's absence) (Mehlotra et al, 2001). For Dd2 and the controls C‐1Dd2 and C‐2Dd2, the QN RMI was 0.31–0.42 (equivalent to a QN resistance reversal of 69–58%). This was in sharp contrast to T76K‐1Dd2 and T76K‐2Dd2, for which the QN RMI increased to 0.89 and 0.91 (P<0.01; see Supplementary Figure 1). Thus, the loss of T76 abolished VP reversibility of both CQ and QN resistance in Dd2 parasites.

PfCRT transmembrane domain 1 mutations preceding K76T determine the degree of VP reversibility

Dd2 and 7G8 are known to differ in their degree of VP reversibility (Mehlotra et al, 2001; Figures 2 and 3), and differ at seven positions in PfCRT, including residues 72, 74 and 75 in transmembrane domain 1 (TMD1) that precede K76T (Table I). To assess whether the preceding TMD1 mutations might be responsible for these differences in VP reversibility, we replaced 7G8‐type TMD1 with the corresponding Dd2 sequence, while retaining all downstream 7G8 mutations. Recombinant parasites expressing this chimeric gene were identified and two clones, TMD1‐1 and TMD1‐2 (Table I), were characterized. Considering amino acids 72–76 as a haplotype, these TMD1 mutants were CVIET, versus CVIEK for the back‐mutants (T76K‐17G8 and T76K‐27G8) and SVMNT for the recombinant controls (C‐17G8 and C‐27G8). Molecular analyses confirmed their recombinant nature (Figure 1; data not shown).

Drug assays (4–7 for each line, performed in duplicate) produced CQ and m‐dCQ RMI values, respectively, of 0.37 and 0.43 for TMD1‐1 and 0.36 and 0.39 for TMD1‐2. In comparison, CQ and m‐dCQ RMI values were 0.35 and 0.37 for Dd2 and 0.68 and 0.64 for 7G8 (consistent with reduced VP reversibility in 7G8). CQ RMI values were significantly different between TMD1 lines and 7G8 (P<0.05; Supplementary Figure 1). Yet TMD1 and recombinant controls maintained very similar CQ and m‐dCQ IC50 values (Supplementary Table I). TMD1 lines also displayed a statistically significant reduction in QN+VP IC50 values compared to recombinant controls (P<0.05; Figure 3). These data suggest that while residue 76 can largely determine the presence or absence of VP reversibility of CQ or QN resistance, the preceding TMD1 mutations influence the degree of reversibility.

CQR depends on precise chemical specificity of mutant PfCRT for CQ

Recombinant lines were assayed (4–5 times in duplicate) for their susceptibility to diaminoalkanes containing the same quinoline ring as CQ yet differing in their side‐chain length. Compounds AQ13, AQ26, AQ33 and AQ40 contained, respectively, three, four, six or 12 CH2 groups, compared to CQ that has a five‐carbon side chain (De et al, 1996).

Assays with Dd2 indicated a pronounced bell‐shaped curve with highest resistance to CQ (Figure 4). Crossresistance was clearly evident with the analogs that varied by only a single CH2 group (i.e. AQ26 and AQ33), yet was absent when two CH2 groups were removed (AQ13) or six were added (AQ40). Results with 7G8 were suggestive of a slightly higher degree of crossresistance to AQ26 and AQ33 (also see Supplementary Table II). Compared to parental lines, recombinant controls displayed lower IC50 values, yet maintained similar crossresistance patterns. Strikingly, removal of K76T ablated this bell‐shaped profile in both Dd2 and 7G8, such that resistance was lost to CQ, AQ26 and AQ33.

Figure 4.

Effect of PfCRT mutations on parasite susceptibility to CQ side‐chain analogs. Shown are the mean±s.e.m. IC50 values (in nM) for recombinant and nontransfected lines tested with CQ and the side‐chain diaminoalkane analogs AQ13, AQ26, AQ33 and AQ40 (De et al, 1996). For each line, statistical comparisons between individual analogs and CQ were performed using unpaired, two‐tailed t‐tests (see Supplementary Table II). *P<0.05; **P<0.01; ***P<0.001.

These analogs were also tested on recombinant GC03 parasites engineered to express Dd2 or 7G8 forms of pfcrt in the place of the wild‐type allele (Sidhu et al, 2002). C4Dd2 and C67G8 clearly displayed crossresistance to the most closely related CQ analogs AQ26 and AQ33, closely resembling the respective Dd2 and 7G8 profiles (Figure 4). This pattern was not observed with C2GC03. Thus, pfcrt mutations appear to largely account for patterns of crossresistance to CQ analogs.

PfCRT controls the degree of saturable CQ accumulation at equilibrium

We leveraged the availability of these recombinant lines to further investigate the CQR mechanism, beginning with measurements in infected red blood cells (iRBCs) of saturable CQ accumulation at equilibrium. These measurements can be used to extrapolate apparent affinities of saturable CQ binding (apparent Kd values), which were previously observed to correlate with CQ's antimalarial activity (Bray et al, 1998). Results with parental lines (GC03, Dd2 and 7G8) confirmed a close correlation between apparent Kd (Figure 5A–C) and CQ IC50 values (Supplementary Table I). We then determined whether changes in equilibrium CQ accumulation could be related to PfCRT polymorphisms. Remarkably, the back‐mutants displayed a dramatic increase in equilibrium CQ accumulation (resulting in apparent Kd values of 10.0 and 9.2 nM for T76K‐1Dd2 and T76K‐17G8, as compared to apparent Kd values of 175.6 and 128.3 nM for Dd2 and 7G8, respectively; Figure 5A and B). This paralleled the ability of T76K to ablate CQR in both genetic backgrounds. Recombinant controls displayed CQ accumulation levels that were slightly lower than parental lines.

Figure 5.

CQ accumulation at equilibrium in recombinant and parental lines. For each line, the curve of best fit is shown for the saturable uptake of [3H]CQ in infected erythrocytes ([CQ]int), expressed as a function of varying extracellular concentrations of unlabeled CQ. Mean±s.e.m. data (from 6 independent experiments) and apparent Kd values are shown for (A) Dd2, (B) 7G8 and (C) GC03 sets of recombinant and parental lines. (D) Apparent Kd values were plotted against CQ IC50 values. Log10‐transformed data, line of best fit and correlation coefficient are indicated.

We also examined the GC03 recombinant lines expressing different pfcrt alleles (Sidhu et al, 2002). Here, the control CQ‐sensitive line C2GC03 displayed CQ accumulation levels equivalent to GC03 (apparent Kd=15.4 and 12.0 nM, respectively). Similarly, the CQ‐resistant mutant lines C4Dd2 and C67G8 displayed levels of CQ accumulation (apparent Kd=143.7 and 170.1 nM, respectively) comparable to Dd2 and 7G8 (Figure 5C). Apparent Kd values were highly correlated with CQ IC50 values for these 10 lines (r2=0.91; Figure 5D). These data confirm an association between saturable CQ accumulation at equilibrium and sensitivity to this drug and imply a key role for the PfCRT K76T mutation.

PfCRT controls the amount of CQ bound to FP

Saturable equilibrium CQ accumulation in iRBCs has been attributed to drug binding to FP (most likely in its dimeric β‐hematin state) (Bray et al, 1998). To investigate whether PfCRT might therefore regulate CQ access to FP, we measured [3H]CQ binding to this intracellular target. Results demonstrated a five‐ to eight‐fold greater CQ binding to FP in GC03 as compared to Dd2 and 7G8 (Figure 6A–C). Similarly, T76K‐1Dd2 and T76K‐17G8, compared with their respective recombinant controls, demonstrated a five‐fold increase in CQ–FP binding (P<0.001; Figure 6A and B). Comparing C4Dd2 or C67G8 with C2GC03, the CQ‐resistant mutants displayed a six‐fold reduction (P<0.001; Figure 6C). These data suggest that pfcrt mutations largely dictate the amount of CQ–FP binding in iRBCs.

Figure 6.

CQ–FP binding in recombinant and parental lines. Levels of [3H]CQ binding to its FP target, expressed as picomoles of CQ per micromole of FP, are shown for (A) Dd2, (B) 7G8 and (C) GC03 sets of recombinant and parental lines. Data represent means±s.e.m., calculated from 6–8 independent experiments. (D) CQ–FP binding values were plotted against CQ IC50 values calculated in the absence or presence of VP. Log10‐transformed data, line of best fit and correlation coefficient are indicated.

Upon addition of VP, all CQ‐resistant parental and recombinant control lines displayed a stimulation of CQ–FP binding (by 2.1‐ to 2.6‐fold and 1.2‐ to 1.4‐fold for lines expressing Dd2 or 7G8 pfcrt alleles, respectively, comparable to changes in IC50 values observed with VP; Supplementary Table I). In contrast, no change was observed with VP for the CQ‐sensitive lines T76K‐1Dd2, T76K‐17G8 and C2GC03 (Figure 6A–C). This implied that the ability of VP to increase CQ–FP binding was dependent on the presence of PfCRT K76T.

These studies also revealed increased CQ–FP binding in C‐1Dd2 compared to Dd2 (tested±VP), closely mirroring differences between these lines in their CQ±VP IC50 values. However, similar CQ–FP binding was observed for C‐17G8 and 7G8, despite differences in expression levels and CQ±VP IC50 values. One explanation, aside from possible experimental variability, could be that changes in PfCRT expression levels have a greater impact on CQ–FP binding in Dd2 than in 7G8. Nevertheless, for both genetic backgrounds, K76T appeared equally important in restricting CQ–FP binding (Figure 6A and B). Comparisons for all 10 lines, tested±VP, revealed a strong inverse relationship between CQ–FP binding and CQ IC50 values (r2=0.93; Figure 6D).

Discussion

The allelic exchange studies reported herein demonstrate a critical role for the PfCRT K76T mutation in maintaining P. falciparum CQR. Replacement of mutant T76 with wild‐type K76 in Old and New World parasites caused a total loss of CQR, with significant reductions in CQ IC50 values, loss of VP reversibility and increased saturable CQ accumulation at equilibrium. These studies, combined with earlier work (Sidhu et al, 2002), provide evidence that pfcrt mutations are necessary and sufficient for CQR, which in turn appears to be dependent on the presence of K76T.

CQ‐resistant parasites, however, differ in their degree of in vitro resistance, presumably due to a secondary contribution of other genes (Chen et al, 2002). One candidate is pfmdr1, which like pfcrt encodes a DV transmembrane protein, and which contains point mutations found to affect the degree of in vitro CQR in 7G8 parasites (Reed et al, 2000). Notably, pfmdr1 mutations have been associated with in vitro CQR in several but not all studies (Uhlemann et al, 2005). In contrast, most studies on pfcrt mutations, notably K76T, indicate an excellent association with in vitro CQR (reviewed in Bray et al, 2005). Some studies on patient blood samples nevertheless suggest that in vitro CQ‐resistant isolates might sometimes lack K76T (Thomas et al, 2002; Lim et al, 2003). Those data would suggest that P. falciparum could acquire CQR independently of pfcrt, although it must be pointed out that to date, this has not been demonstrated with culture‐adapted, non‐drug‐pressured, individual parasite lines.

Additional evidence that mutant pfcrt is the primary CQR determinant comes from Wootton et al (2002), who identified this as the genomic locus of greatest recent sequence evolution, consistent with the tremendous global impact of CQ pressure on parasite populations and the selective sweep for mutant pfcrt‐containing CQ‐resistant strains emanating from a few geographic origins (Chen et al, 2003). Recent studies from The Gambia also indicate that mutant pfcrt parasites can be more efficiently transmitted to the mosquito vector, providing a potentially significant selective advantage under CQ pressure (Sutherland et al, 2002). Yet studies from Malawi reported a significantly decreased frequency of mutant pfcrt since CQ use was discontinued a decade ago, implying that mutant pfcrt parasites carry a fitness disadvantage in the absence of CQ pressure (Kublin et al, 2003; Mita et al, 2003).

While our data indicate a key role for K76T in CQR, it is notable that no fewer than four pfcrt mutations have ever been found in a CQ‐resistant isolate (Chen et al, 2003). These other mutations may have been selected to compensate for loss/alteration of endogenous function associated with acquisition of K76T, or may themselves directly contribute to resistance to CQ or other antimalarial drugs. To test whether K76T might itself be sufficient to confer VP‐reversible CQR in vitro (which presumably is more favorable than in vivo semi‐immune conditions), we employed allelic exchange to introduce solely this mutation into wild‐type pfcrt (in GC03). From multiple episomally transfected lines, one showed evidence of K76T substitution in the recombinant, full‐length pfcrt locus (data not shown). However, these mutant parasites failed to expand in the bulk culture and could not be cloned, despite numerous attempts. These results suggest reduced parasite viability resulting from K76T in the absence of other pfcrt mutations. This situation is not reciprocal however, in that parasites harboring all the other mutations except for K76T (illustrated by our back‐mutants) show no signs of reduced viability in culture.

Multiple in vivo studies have also shown a strong association between K76T and increased risk of CQ treatment failure (defined either as persistence or reappearance of parasitemia or clinical symptoms) (Wellems and Plowe, 2001; Jelinek et al, 2002; Nagesha et al, 2003), with some exceptions (Ariey et al, 2002; Happi et al, 2003). Many studies report 100% sensitivity of the K76T mutation (i.e. 100% of treatment failure cases harbor this mutation), yet a lower specificity (i.e. not all infections harboring K76T result in treatment failure). Evidence that age‐dependent immunity can contribute to clearance of K76T infections came from elegant studies in Mali by Djimdé et al (2001, 2003). Some associations have also been reported between CQ treatment failure and the pfmdr1 N86Y mutation (Wellems and Plowe, 2001; Tinto et al, 2003). Interestingly, reports have also shown in vivo evidence that CQ treatment of mixed infections with mutant and wild‐type pfcrt resulted in treatment failures harboring solely mutant pfcrt (Basco et al, 2002; Schneider et al, 2002). The combined data provide powerful support for the use of K76T as a sensitive molecular marker of CQ‐resistant P. falciparum malaria in areas where this mutation has not attained 100% prevalence.

The earlier massive reliance on CQ as a first‐line antimalarial and the devastating impact of CQR on malaria mortality and morbidity rates have stimulated intense efforts to elucidate CQ's mode of action and the CQR mechanism. CQ accumulation in the acidic DV is thought to result from a proton trapping mechanism as well as from high‐affinity CQ binding to FP, producing lethal concentrations of FP or FP–drug complexes (Sullivan et al, 1998; Bray et al, 2005). CQ‐resistant P. falciparum strains circumvent this toxicity by accumulating lesser drug in the DV (Saliba et al, 1998). Interestingly, both sensitive and resistant strains of P. falciparum reportedly have similar capacities of CQ–FP binding and total FP quantities; yet, resistant parasites exhibit markedly reduced CQ accumulation at equilibrium (Bray et al, 1998; Zhang et al, 1999). These findings implicate restricted CQ access to FP as a central component of CQR.

Here we present compelling evidence that CQ access to FP is determined by mutations in pfcrt. Remarkably, reduced CQ access to FP appears to be dependent on the status of PfCRT position 76. T76K back‐mutants displayed five‐fold increases in CQ accumulation at equilibrium, rendering them CQ sensitive. Furthermore, replacement of wild type with mutant pfcrt (conferring CQR; Sidhu et al, 2002) produced six‐fold reductions. These data were verified by two independent methods. First, measurements of the binding of [3H]CQ to FP (in the form of hemozoin) established a direct relationship between parasite susceptibility to CQ, saturable CQ accumulation at equilibrium and CQ–FP binding, which in turn depended on the PfCRT haplotype. Second, the use of protease inhibitors (including Ro 40‐4388) to block hemoglobin digestion and prevent FP release indicated that the apparent affinity of CQ binding observed in the pfcrt‐modified lines was dependent on FP availability (data not shown).

These data can be useful in evaluating models proposed to account for reduced CQ access to FP, which include (1) leakage of charged drug, which results in its extrusion from the DV; (2) carrier‐mediated, energy‐dependent CQ efflux; (3) altered partitioning or heme turnover rates resulting from DV pH changes; (4) reduced activity of a putative CQ importer; or (5) the possible involvement of heme binding proteins (Ursos and Roepe, 2002; Sanchez et al, 2004; Waller et al, 2004). Little evidence currently supports the two latter models. Studies on DV pH changes remain a subject of intense debate, with earlier predictions that a less acidic DV would cause reduced CQ accumulation in CQ‐resistant parasites (Ginsburg and Stein, 1991). The opposite conclusion, that is, CQ‐resistant parasites appear to have a more acidic DV, was obtained by Roepe and colleagues using single‐cell fluorophotometric studies (Ursos and Roepe, 2002; Bennett et al, 2004). This was postulated to cause CQR by altering heme aggregation rates, effectively reducing amounts of free FP available for CQ binding. These authors, however, recently provided evidence that this might be secondary to a role of pfcrt in binding to CQ and altering its accumulation (Zhang et al, 2004). Furthermore, a primary role for pH in CQR has been challenged on experimental grounds (Bray et al, 2002; Wissing et al, 2002; Sanchez et al, 2003).

This leaves charged drug leak and carrier‐mediated CQ efflux as two leading models at present. The former predicts that CQ2+ can leak, through mutant PfCRT, out of the DV along a massive concentration gradient, made possible in part by the loss of the positively charged K76, predicted to lie on the lumenal side of the DV membrane (Warhurst et al, 2002; Martin and Kirk, 2004). This could reduce the DV concentration of CQ, resulting in reduced accumulation of saturable CQ at equilibrium and reduced CQ–FP binding. Additional support for the drug leak hypothesis came from a recent study involving halofantrine or AMT pressuring of a CQ‐resistant line (Johnson et al, 2004). This produced resistant lines that had become CQ sensitive and had acquired pfcrt mutations, including S163R in TMD4 that may have restored charge to a transmembrane conformation and prevented CQ2+ efflux, despite the presence of K76T in TMD1. Our results could also agree with the carrier‐mediated efflux model, which posits that CQ‐resistant parasites have an ATP‐ and temperature‐dependent inhibitable CQ efflux carrier (Sanchez et al, 2003, 2004). Supporting data demonstrated that, in CQ‐sensitive lines, external CQ appeared to compete with [3H]CQ for carrier binding sites, leading to reduced [3H]CQ accumulation with increasing external CQ concentrations. In contrast, in CQ‐resistant parasites, accumulation of [3H]CQ first increased at low external CQ concentrations and then decreased. This led to the proposal that CQ‐resistant parasites have an active CQ efflux carrier, with ‘trans’ CQ competing at low concentrations for carrier binding sites, leading to increased [3H]CQ accumulation, and at higher concentrations saturating the receptor sites and out‐competing [3H]CQ accumulation (Sanchez et al, 2003). These assays are clearly warranted with the new mutant lines reported herein.

Both the charged drug leak and the active efflux models are consistent with substrate specificity. Interestingly, results presented herein indicate that the ability of mutant PfCRT to confer CQR is precisely configured for CQ. Resistance was rapidly lost following subtle structural modifications of the basic diethylamino side chain linked to the 4‐aminoquinoline ring structure, an encouraging result with respect to the possibility of developing CQ analogs effective against drug‐resistant P. falciparum (De et al, 1996). These findings add to a growing body of evidence in support of PfCRT directly contributing to CQ transport, including recent bioinformatic analyses that places this protein in the drug/metabolite transporter superfamily and pfcrt heterologous expression studies in yeast and Dictyostelium discoideum (Martin and Kirk, 2004; Tran and Saier, 2004; Zhang et al, 2004; Naude et al, 2005).

Our data also suggest that protonated VP may physically interact with mutant PfCRT, possibly interfering with CQ (and perhaps QN) transport out of the DV. We observed that VP partially restored CQ–FP binding in parasites expressing mutant pfcrt and found that the Dd2 T76K back‐mutants had lost VP reversibility of CQ and QN resistance. These data also revealed a direct relationship between K76T and VP chemosensitization of QN resistance, consistent with indications from quantitative trait loci analysis that although QN resistance appeared to be multifactorial, its reversibility by VP was tightly linked to pfcrt (Ferdig et al, 2004). Interestingly, Cooper et al (2002) reported the selection of a mutant line harboring K76I, which paradoxically had an increased QN IC50 value with VP. Our TMD1 mutant lines also confirmed an earlier postulate that VP reversibility could be influenced by mutations preceding K76T, which differ between South American/Pacific and Asian/African parasites (Mehlotra et al, 2001; Warhurst, 2003). Novel pfcrt mutations have also been identified downstream of TMD1 that may also affect VP reversibility, although this has yet to be confirmed by allelic exchange (Chen et al, 2003; Johnson et al, 2004).

In conclusion, our data establish that the PfCRT K76T mutation plays a key role in determining CQ susceptibility in P. falciparum strains of New and Old World origins, is an important determinant of VP reversibility of CQ and QN resistance and can largely explain patterns of crossresistance to CQ side‐chain analogs. These data provide a compelling argument that direct interactions between CQ and this mutant residue, leading to reduced CQ–FP interactions and drug extrusion from the DV, are key to CQR.

Materials and methods

Plasmid constructs

For allelic exchange, a 2.9 kb pfcrt fragment containing a 1.6 kb promoter element and 1.3 kb of the gene (exons 1–5) was PCR amplified from 106/1 (Sudan) or Dd2 genomic DNA using primers p1 (ACGGATCCGGTACCTTAGAACCCTAAGAATATCAGCTC; pfcrt 5′UTR‐specific; BamHI and KpnI sites underlined) and p2 (TTGCGGCCGCATGCATGTCATGTTTGAAAAGCATACAGGC; pfcrt exon 5′‐specific; NotI and NsiI sites underlined). Sequence‐verified products were subcloned into BamHI–PstI‐digested pMini‐BSD, which expresses the bsd selectable marker that is under the transcriptional control of calmodulin 5′UTR and hrp2 3′UTR sequences. The resulting constructs harboring the 106/1 and Dd2 pfcrt sequences were termed pK76 and pT76, respectively. Luciferase constructs and assays are described in Supplementary data.

Parasite propagation, transfection, cloning and drug susceptibility assays

P. falciparum culturing, stable transfection leading to allelic exchange, limiting dilution cloning and 72 h [3H]hypoxanthine incorporation assays (to determine drug IC50 values) were performed as described (Waller et al, 2003). Complete drug response curves obtained from a representative assay are included in Supplementary Figure 2.

Nucleic acid and protein analyses

The functional downstream recombinant pfcrt locus was detected by PCR using primers p3 (CAATTAACCCTCACTAAAGGG; pBluescript‐specific) and p4 (CCCAAGAATAAACATGCGAAACC; pfcrt exon 7‐specific) (Figure 1A). The truncated upstream recombinant pfcrt fragment was detected using primers p5 (CTTCAATTCTCATATTTCAATATATTCC; pfcrt 5′UTR‐specific) and p6 (GATAGCGATTTTTTTTACTGTCTG; hrp2 3′UTR‐specific) (Figure 1A). PCR products were nested using pfcrt primers p7 (AATTCAAGCAAAAATGACGAGCG; exon 1‐specific) and p8 (ACTGAACAGGCATCTAACATGG; exon 3‐specific). For RT–PCR, cDNA was amplified using primers p7 and p9 (TTCCTACACGGTAAATTATAGAACC; pfcrt exon 12‐specific) for the functional gene and primers p7 and p6 for the upstream remnant. RT–PCR products were sequenced across codons 72–76. For Northern blotting, total RNA was resolved, blotted and hybridized with an exon 5–13 cDNA probe generated with primers p10 (CATTTACCATATAATGAAATATGGAC) and p11 (GTTAATTCTCCTTCGGAATCTTCATTTTCTTCAT). For Western blotting, parasites were doubly synchronized, harvested as early‐mid trophozoites and protein samples normalized using parasitemia, hematocrit and densitometry (also see Supplementary data).

Equilibrium CQ accumulation assays

These were performed as described (Bray et al, 1998). Briefly, synchronized trophozoites were incubated in triplicate for 1 h at 37°C in bicarbonate‐free RPMI with 10 mM HEPES, pH 7.4, 1 nM [3H]CQ and 5–250 nM unlabeled CQ. Counts corresponding to nonsaturable CQ uptake into iRBCs (calculated using 100 μM external CQ), as well as CQ uptake by uninfected RBCs, were subtracted from the average total counts to yield saturable CQ uptake at equilibrium. Data were analyzed by nonlinear regression (Marquart method). Standard errors were calculated using matrix inversion (Erithacus Software Ltd, UK).

CQ–FP binding assays

CQ–FP binding was measured by assessing the incorporation of sublethal concentrations of [3H]CQ into hemozoin crystals (Sullivan et al, 1996). Briefly, synchronized P. falciparum cultures were incubated for 48 h with 1 nM [3H]CQ±0.8 μM VP. Parasites were recovered from saponin‐lysed RBCs, washed, resuspended in hypotonic buffer and sonicated. Hemozoin was purified by sucrose centrifugation, washed with SDS, dissolved with NaOH into its monomeric FP form, and FP and [3H]CQ concentrations were determined (see Supplementary data). Earlier studies showed that free FP is required for CQ–hemozoin binding; therefore, these measurements can be used to extrapolate CQ–FP binding (Sullivan et al, 1996, 1998).

Supplementary data

Supplementary data are available at The EMBO Journal Online.

Supplementary Information

Supplementary Information [emboj7600681-sup-0001.pdf]

Acknowledgements

We thank Myles Akabas (AECOM), Scott Bohle (McGill University), Choukri Ben Mamoun (U Conn. Health Center), Paul Stocks (Liverpool School of Tropical Medicine) and MR4 (ATCC, Virginia) for helpful discussions, reagents or experimental assistance. Funding was provided by NIAID/NIH (R01 AI50234) and awards from the Speaker's Fund for Biomedical Research and the Ellison Medical Foundation Scholars in Global Infectious Diseases program (DF), and the Wellcome Trust, MRC and BBSRC (PGB and SAW).

References