Malaria parasites contain a relict plastid, the apicoplast, which is considered an excellent drug target due to its bacterial-like ancestry. Numerous parasiticidals have been proposed to target the apicoplast, but few have had their actual targets substantiated. Isopentenyl pyrophosphate (IPP) production is the sole required function of the apicoplast in the blood stage of the parasite life cycle, and IPP supplementation rescues parasites from apicoplast-perturbing drugs. Hence, any drug that kills parasites when IPP is supplied in culture must have a nonapicoplast target. Here, we use IPP supplementation to discriminate whether 23 purported apicoplast-targeting drugs are on- or off-target. We demonstrate that a prokaryotic DNA replication inhibitor (ciprofloxacin), several prokaryotic translation inhibitors (chloramphenicol, doxycycline, tetracycline, clindamycin, azithromycin, erythromycin, and clarithromycin), a tRNA synthase inhibitor (mupirocin), and two IPP synthesis pathway inhibitors (fosmidomycin and FR900098) have apicoplast targets. Intriguingly, fosmidomycin and FR900098 leave the apicoplast intact, whereas the others eventually result in apicoplast loss. Actinonin, an inhibitor of bacterial posttranslational modification, does not produce a typical delayed-death response but is rescued with IPP, thereby confirming its apicoplast target. Parasites treated with putative apicoplast fatty acid pathway inhibitors could not be rescued, demonstrating that these drugs have their primary targets outside the apicoplast, which agrees with the dispensability of the apicoplast fatty acid synthesis pathways in the blood stage of malaria parasites. IPP supplementation provides a simple test of whether a compound has a target in the apicoplast and can be used to screen novel compounds for mode of action.
Malaria is caused by the protozoan parasite Plasmodium, and six species of Plasmodium infect humans. In 2015, 3.2 billion people in ∼100 countries were at risk for malaria, and there were 212 million infections and 429,000 deaths (1). Malaria also causes economic losses of billions of dollars in parts of the world that cannot afford it (2). Drugs are a major element of malaria control, but the specter of drug resistance is a constant worry and provides an ongoing impetus to identify new drug leads in order to stay one step ahead of the parasites.
Identification of a relict plastid (apicoplast) in Plasmodium parasites provided a new set of potential drug targets for the battle against malaria. Plastids, which ultimately derive from endosymbiotic bacteria, maintain a small genome (∼35 kb in the case of malaria parasites) that is separate from the nucleus and is prokaryotic in its structure and mode of expression (3–7). The apicoplast was acquired by secondary endosymbiosis prior to the separation of phylum Apicomplexa (intracellular parasites) from chromerids and dinoflagellates (photosynthetic algae) around 450 million years ago (8–11). The apicoplast genome encodes large subunit and small subunit rRNAs, a complete set of tRNAs, 18 ribosomal proteins, three subunits of RNA polymerase, a protein implicated in DNA replication, a translation elongation factor Tu, and a subunit of Clp protease (3–7, 12). Similar to other plastids, the majority of the original apicoplast DNA has undergone endosymbiotic gene transfer to the nucleus, which encodes ∼450 proteins that are targeted to the apicoplast (13).
The Plasmodium apicoplast lacks enzymes or pigments required for photosynthesis; however, it retains a number of other anabolic pathways that are indispensable at some stage of the life cycle for parasite growth or viability. These include isoprenoid precursor biosynthesis, fatty acid biosynthesis, Fe-S cluster assembly, and heme biosynthesis (13–16). The apicoplast is thus indispensable, and either genetic or pharmaceutical perturbation of its activities kills parasites, making the apicoplast a valid drug target (17–30).
Over the course of 20 years of Plasmodium apicoplast research, numerous apicoplast inhibitors have been postulated. Although these compounds kill parasites, few have been validated as actually perturbing the apicoplast. For instance, apicoplast type II fatty acid biosynthesis (FASII) is dispensable in the malaria parasite blood stage, which casts doubt on the mode of activity of a range of parasiticidal compounds alleged to have targets in fatty acid biosynthesis (31). Similarly, heme biosynthesis also appears to be dispensable in blood stages, yet the heme biosynthesis inhibitor succinylacetone still kills parasites (32). It is also unclear whether all the tested protein translation inhibitors postulated to inhibit apicoplast protein synthesis have sole targets in the apicoplast or whether they also inhibit translation in other compartments, such as the mitochondrion or cytosol. Furthermore, some drugs with bona fide apicoplast targets, such as the translation inhibitor azithromycin (33), also have secondary targets, namely, the blocking of host cell invasion by the merozoite stage of the parasite (34). There is thus a rather patchy understanding of how these drugs actually kill parasites, even though several are in clinical use (35, 36).
An ideal way to approach this dilemma would be to examine the effect of these compounds on parasites not dependent on their apicoplast. Yeh and DeRisi (37) showed that isopentenyl pyrophosphate (IPP) production is the sole required function of the apicoplast in the asexual blood stage, and the apicoplast could be lost by Plasmodium falciparum as long as IPP is supplied in the culture medium. This unique feature allows us to separate out the effect of any drug on the apicoplast from effects on any alternative target during the asexual blood stage of P. falciparum. Here, we used a chemical supplementation approach (37), a technique now routinely in use (30, 38–42), to test the target of 23 presumed apicoplast drugs, including compounds with confirmed apicoplast targets, confirmed nonapicoplast targets, and compounds with putative apicoplast targets.
IPP supplementation rescues parasites only from validated apicoplast-targeting drugs.
Although IPP is now used increasingly to discriminate whether or not a drug has a target in the apicoplast, we performed IPP supplementation trials on parasites exposed to two definite nonapicoplast drugs (chloroquine and atovaquone) and two validated apicoplast drugs (azithromycin and fosmidomycin) to establish a comprehensive set of assays focused on apicoplast target validation (Table 1 and Fig. 1) (30, 37–42). Chloroquine targets the food vacuole in which the parasite digests ingested hemoglobin (43, 44), and atovaquone targets the cytochrome bc1 complex in the mitochondrion (45, 46). For bona fide apicoplast-targeting drugs, we used azithromycin (which inhibits apicoplast protein synthesis by binding to the large subunit of apicoplast ribosomes) (33) and fosmidomycin, which inhibits 1-deoxy-d-xylulose 5-phosphate reductoisomerase (DXR), an enzyme of the nonmevalonate apicoplast IPP synthesis pathway (19, 47–50). Our growth assays across two cycles (Table 1) confirm previous reports of immediate death (no difference between 1st- and 2nd-cycle half-maximal inhibitory concentrations [IC50s]) for chloroquine, atovaquone, and fosmidomycin, in contrast to delayed death for azithromycin, for which the IC50 is more than 100-fold lower in the second cycle (51).
A drug targeting apicoplast housekeeping functions (azithromycin) has a substantially lower IC50 in the 2nd asexual erythrocytic growth cycle. Drugs with targets outside the apicoplast (chloroquine and atovaquone) or targeting apicoplast metabolism (fosmidomycin) show little change with prolonged exposure to the drug. Each drug concentration was assayed in triplicate.
We tracked parasite growth for four red blood cell asexual cycles (4 × 48 h/cycle) in the presence of our four test drugs, with and without IPP (Fig. 1A). IPP supplementation clearly rescues parasites from the confirmed apicoplast drugs azithromycin and fosmidomycin but not from chloroquine or atovaquone (nonapicoplast drugs), thus validating the use of IPP to confirm apicoplast targets in growth assays (Fig. 1B and see Fig. S1A and B in the supplemental material) (37). Drug-free controls with and without IPP supplementation demonstrate that 200 μM IPP does not significantly inhibit growth (Fig. 1).
Parasites rescued with IPP lose their apicoplast genome when targeted with the housekeeping inhibitor azithromycin.
We used quantitative PCR to amplify single-copy genes on the apicoplast, mitochondrial, and nuclear genomes to monitor the ratio of organellar to nuclear genomes during the course of drug treatment and IPP rescue (37). Figure 1B demonstrates a marked decline in the apicoplast-to-nuclear genome ratio after the 2nd cycle in azithromycin-treated parasites, regardless of supplementation with IPP. At the end of the fourth cycle, the apicoplast-to-nuclear genome ratio was reduced almost 100-fold by azithromycin treatment (Fig. 1B). We conclude that the disruption of apicoplast protein synthesis by azithromycin results in a loss of the organelle DNA, similar to the effects of the protein synthesis inhibitor doxycycline (37). The mitochondrial genome-to-nuclear genome ratio did not decline (Fig. 1B), which strongly suggests that azithromycin does not target housekeeping functions in the parasite mitochondrion. In contrast, there was no change in the apicoplast-to-nuclear genome ratio (Fig. 1B) or the mitochondrial genome-to-nuclear genome ratio (Fig. 1B) in fosmidomycin-treated parasites rescued with IPP, confirming previous findings (37, 38, 52).
Parasites rescued with IPP have impaired protein import capacity when targeted with the housekeeping inhibitor azithromycin.
Apicoplast transit peptide processing is a marker of successful protein import into the apicoplast (53, 54). In the D10 ACPLGFP parasite line, the 33-kDa transit peptide/green fluorescent protein (GFP) precursor protein is cleaved to produce a 30-kDa mature (processed) protein as a consequence of being transported into apicoplast (53). Canonically, the amount of unprocessed protein is minimal, often being just barely detectable (53, 54), and our nondrugged parasites (with or without IPP supplementation) contain predominantly processed GFP (Fig. 1C). In contrast, azithromycin-treated, IPP-rescued parasites contain predominantly the unprocessed 33-kDa transit peptide/GFP precursor (Fig. 1C). Fosmidomycin-treated, IPP-rescued parasites exhibit normal protein processing equivalent to untreated parasites (Fig. 1C), which is consistent with them retaining their apicoplast genomes (Fig. 1B).
Parasites rescued with IPP have disrupted apicoplasts when targeted with the housekeeping inhibitor azithromycin.
Visualization of the apicoplast during the 4th cycle after azithromycin treatment in conjunction with IPP rescue reveals substantial disruption of apicoplast integrity (Fig. 1D). Nondrugged cells (either with or without IPP) have a single punctate apicoplast at the late-ring/early trophozoite stage (Fig. 1D). However, parasites treated with azithromycin and rescued with IPP have very diffuse GFP throughout the parasite (Fig. 1D). Conversely, fosmidomycin treatment in conjunction with IPP rescue produced parasites with healthy-looking apicoplasts (Fig. 1D), again in agreement with the other assays (38). Combined with our previous data showing the loss of the apicoplast genome (Fig. 1B) and impairment of apicoplast-specific protein processing (Fig. 1C) following azithromycin treatment, the loss of an identifiable apicoplast demonstrates that the apicoplast translation inhibitor azithromycin disrupts apicoplast integrity.
Our assays with four test compounds demonstrate the utility of IPP supplementation for discriminating between compounds that perturb the apicoplast (azithromycin and fosmidomycin) and those that do not (chloroquine and atovaquone). Tracking the survival of apicoplast DNA, assaying apicoplast protein import, and visualizing the compound's effect on GFP-labeled apicoplasts permit a detailed investigation of how a compound perturbs this vital organelle (Fig. 1). We next set out to screen a suite of compounds with alleged targets in the apicoplast to explore whether or not they indeed impact the apicoplast.
Effect of bacterial housekeeping inhibitors on the apicoplast.
The apicoplast genome must be copied, transcribed, and translated to generate the ∼30 proteins it encodes (4). Ciprofloxacin inhibits bacterial DNA gyrase A and perturbs apicoplast genome duplication (18). We observe delayed death with this drug (Table 2), confirming previous reports (18, 55). IPP rescued growth in the presence of ciprofloxacin (Table 2), and similar to azithromycin, the apicoplast genome was gradually lost (Fig. 2A), as were apicoplast protein import (Fig. 2B) and apicoplast integrity (Fig. 2C). We conclude that ciprofloxacin exclusively targets the apicoplast, likely the nucleus-encoded apicoplast-targeted DNA gyrase A (13, 56).
TABLE 2 List of drugs assayed, IC50s, IPP rescue, and impact on the apicoplasta
Rifampin is a bacterial transcription inhibitor proposed to act by perturbing apicoplast transcription (57, 58). We confirm a previous report that rifampin causes immediate death (51) and find that IPP supplementation does not rescue parasites from rifampin (Table 2). These data strongly imply that rifampin has a nonapicoplast target in P. falciparum. Although rifampin may interfere with apicoplast transcription, there appears to be a more immediate mode of action that effectively masks any apicoplast-specific inhibition. Because IPP cannot rescue parasites from rifampin (Table 2 and Fig. S1C), we were unable to analyze apicoplast genome perseverance, apicoplast protein import, or apicoplast morphological integrity.
Doxycycline is widely prescribed as a malaria prophylactic and causes delayed death in P. falciparum, presumably by binding to the apicoplast small subunit rRNA and blocking protein synthesis, as it does in bacteria (55). Here, we confirm previous reports (37, 38, 52) of rescue by IPP from doxycycline (Table 2), associated loss of the apicoplast genome (Fig. 2A), loss of apicoplast protein import (Fig. 2B), and apicoplast integrity (Fig. 2C), and we extend these same observations to the related compound tetracycline (Table 2 and Fig. 2).
Similar responses were seen with five other drugs that putatively target apicoplast translation, regardless of their mechanism of action. Chloramphenicol is bacteriostatic and inhibits the peptidyl transferase activity of the bacterial ribosome by binding to the small subunit rRNA, preventing peptide bond formation. Clindamycin prevents protein synthesis by binding to the 23S subunit of prokaryotic rRNA, thus inhibiting prokaryotic ribosomal translocation. Erythromycin and its derivative clarithromycin also bind to the 23S rRNA but interfere with aminoacyl translocation. Growth during treatment with any of these drugs could be rescued by IPP supplementation (Table 2). Long-term treatment with these drugs in the presence of IPP resulted in a gradual loss of the apicoplast genome (Fig. 2A), a reduction in apicoplast protein import (Fig. 2B), and a loss of apicoplast integrity (Fig. 2C). We conclude that all these bacterial translation inhibitors have their primary target in the P. falciparum apicoplast, most likely in the components of the apicoplast ribosome that are also the targets in bacteria.
Fusidic acid is a fungus-derived prokaryotic protein synthesis inhibitor that prevents the turnover of elongation factor G (EF-G) from the bacterial ribosome. We previously showed immediate death with fusidic acid (59), and Gupta et al. (60) provided evidence that it inhibits both apicoplast ribosomes and mitochondrial ribosomes. Here, we corroborate immediate death, and we were unable to rescue parasites with IPP (Table 2 and Fig. S1D), which confirms that the drug has a target beyond the apicoplast, likely the mitochondrion. IPP-supplemented rescue showed a negative impact on parasite growth with fusidic acid, although the difference is not significant (Fig. S1D). Given that 200 μM IPP supplementation is nontoxic for the parasite (Fig. 1), one explanation is that fusidic acid somehow sensitizes the parasite to IPP, but further investigation will be required to draw any firm conclusion. Immediate death from a target outside the apicoplast prevents any insight as to whether fusidic acid targets the apicoplast. Because any such inhibition would presumably cause delayed death, it is effectively masked by the immediate death, which we suspect is caused by an inhibition of mitochondrial protein synthesis, as previously inferred (60).
Mupirocin binds reversibly to the bacterial isoleucyl t-RNA synthetase preventing protein synthesis. Istvan et al. (61) identified mutations and/or copy number variations of an apicoplast-targeted isoleucyl t-RNA synthetase as conferring resistance to mupirocin. We confirm a previous report of delayed death on P. falciparum (61), and we show rescue by IPP (Table 2), loss of apicoplast DNA (Fig. 2A), impairment of apicoplast protein import (Fig. 2B), and corruption of apicoplast integrity by mupirocin (Fig. 2C), thereby reaffirming an apicoplast target.
Borrelidin is also a tRNA synthetase inhibitor, but it targets threonine tRNA synthetases. Borrelidin causes immediate death in P. falciparum, and IPP supplementation could not rescue parasites exposed to borrelidin (Table 2 and Fig. S1E). Indeed, borrelidin resembles fusidic acid in sensitizing the parasite to IPP (Fig. S1E). The data are thus consistent with a cytosolic target for borrelidin. If borrelidin impacts the apicoplast threonine tRNA synthesis, the effect could be masked by immediate death, likely through the inhibition of cytosolic translation (27).
IPP rescues parasites from the immediate death inhibitor actinonin.
Actinonin is an antibacterial that inhibits the activity of peptide deformylase, an enzyme that removes the formyl group from the formyl methionine used to initiate many bacterial proteins (62–65). Actinonin also inhibits a human mitochondrial peptide deformylase and is being assessed as an antitumor agent (66). Apicoplasts harbor a peptide deformylase (13, 67), but whether or not it is the target of actinonin in malaria parasites is unclear. We confirm previous reports of antimalarial activity for actinonin (68, 69), with an IC50 in the low micromolar range (Table 2). Importantly, we demonstrate that while actinonin does not exhibit delayed death, the growth of actinonin-treated parasites can be rescued by IPP supplementation (Table 2 and Fig. 3A). We also show that actinonin-treated, IPP-rescued parasites suffer a loss of apicoplast DNA (Fig. 3B), have impaired apicoplast protein import (Fig. 3C), and have aberrant apicoplast structure (Fig. 3D). Actinonin is thus starkly different from the collection of bona fide antiapicoplast compounds described above, all of which exhibit delayed death, apparently by perturbing apicoplast housekeeping activities. We previously showed that actinonin specifically retards growth and division of the apicoplast but does not impact growth and division of the mitochondrion or nucleus (69), a finding confirmed here by our quantitative PCR (qPCR) genome analyses (Fig. 3B). Since IPP rescued malaria parasites exposed to actinonin, we tentatively conclude that actinonin primarily targets the P. falciparum apicoplast and not the parasite mitochondrion. Given that actinonin exhibits a different death kinetic to all the established apicoplast housekeeping inhibitors, it seems unlikely that it inhibits apicoplast housekeeping peptide deformylase (67). Rather, recent data demonstrate that actinonin inhibits apicoplast biogenesis in Toxoplasma gondii, and probably P. falciparum, by targeting the apicoplast membrane protein FtsH1 (70), and our results are concordant.
IPP does not rescue parasites from a photosynthesis inhibitor.
Having explored IPP rescue as a way to verify apicoplast targets for various antibacterials with housekeeping targets, we next turned our attention to apicoplast anabolism targets. Toltrazuril, an herbicide that also kills Plasmodium parasites, is postulated to interact with the D1 (PsbA) protein of a relict photosynthetic reaction center in the apicoplast (71). Toltrazuril is also parasiticidal against the apicomplexan parasites Eimeria and Toxoplasma gondii, and in vivo against intestinal and hepatic coccidiosis in rabbits (72–74). However, no psbA gene candidates have been identified in any apicomplexan genome, and there is no evidence for photosynthesis in these parasites. IPP supplementation did not rescue parasites from toltrazuril, which in any case does not exert delayed death and requires high concentrations to kill parasites (Table 2 and Fig. S1F). Our data demonstrate that this herbicide does not target the apicoplast in P. falciparum and confirm that photosynthesis is not a valid malaria drug target.
IPP supplementation rescues parasites from isoprenoid biosynthesis inhibitors but not fatty acid biosynthesis inhibitors.
As described earlier (37, 38, 41) and confirmed above, fosmidomycin inhibition could be rescued by IPP (Fig. 1A). Here, we extend these observations to the fosmidomycin analogue FR900098 (Table 2), which is also posited to target IPP synthesis in the apicoplast (19, 39, 49). Both fosmidomycin and FR900098 exhibit immediate death, and IPP supplementation restored growth (Table 2 and Fig. 4). FR900098 mimics fosmidomycin in that treated parasites rescued by IPP suffer no loss of apicoplast DNA (Fig. 2A), they maintain apicoplast protein import (Fig. 2B), and they preserve apicoplast integrity (Fig. 2C). We conclude that inhibition of the anabolic IPP synthesis pathway, either by fosmidomycin or FR900098, is an apicoplast-specific effect but results in immediate death.
We next turned our attention to apicoplast fatty acid biosynthesis. We confirm that the presumed fatty acid biosynthesis (FASII) inhibitors triclosan (75), cerulenin (76, 77), and hexachlorophene (78, 79) are parasiticidal (Table 2). However, IPP supplementation could not recue the parasites from these compounds (Table 2 and Fig. 4), strongly implying that the primary target of these three compounds is outside the apicoplast. Since genetic knockdown of various FASII enzymes (31) has shown that fatty acid biosynthesis is dispensable in the asexual blood stage of P. falciparum, Plasmodium berghei, and Plasmodium yoelii, our results further confirm that these compounds are off-target.
Heme biosynthesis and protein transport inhibitors.
Gene knockouts indicate that the Plasmodium apicoplast/mitochondrion heme synthesis pathway is also dispensable at the blood stage (32, 80). Unfortunately, the only known inhibitor of heme biosynthesis, succinylacetone, requires extremely high concentrations (>600 μM) to kill P. falciparum parasites in vitro, which obviated rigorous testing for IPP rescue (Table 2 and Fig. S2).
The fungal metabolite brefeldin A (BFA) disrupts protein trafficking from the endoplasmic reticulum (ER) to the Golgi in many organisms (81, 82). It has been debated whether or not protein targeting to the apicoplast is routed via the Golgi in P. falciparum (83, 84). We observe immediate death with BFA, and IPP failed to rescue the parasites (Table 2 and Fig. S1G). We conclude that BFA impacts other nonapicoplast processes, as previously reported, namely, disruption of the Golgi, distension of the nuclear envelope, and export of proteins (85), and this immediate effect masks any perturbation of apicoplast protein targeting.
The goal of a safe, effective, and cheap malaria vaccine is yet to be realized, which places chemotherapy as our primary defense against the disease. Parasite drug resistance has been a chronic issue with antimalarials, and it is clear that a larger arsenal of compounds would allow more flexibility in managing resistance (86, 87). Huge investments are required to find new drugs and identify their targets, so a better understanding of the mode of action of existing antimalarials is an effective use of resources. The use of antibacterial drugs against malaria started long before the apicoplast was identified in the malaria parasite (88–95). Nowadays, we appreciate that the bacteria-like housekeeping machinery of the apicoplast, such as DNA replication, transcription, translation, and posttranslational modification, as well as metabolic pathways for synthesis of fatty acids, isoprenoid precursors, heme, and iron-sulfur complexes in the organelle make it an attractive drug target (17–30, 51, 96–102). However, an important facet of this strategy is knowing when particular pathways are essential to the parasite and being certain that the compounds in question do indeed have an apicoplast target. Moreover, it is strategic to understand the kinetics of drug effect in considering their use. Our approach of screening parasiticidal compounds with IPP rescue, along with a suite of apicoplast viability assays, unequivocally discriminates between compounds whose primary target is in the apicoplast and those that target other aspects of the parasite. These assays further define the mode of action and extend our understanding of drug-target interactions to optimize and prioritize a collection of existing clinically used antimalarials and sundry leads (Fig. 5).
Our data confirm that drugs disrupting the basic housekeeping functions in the apicoplast result in delayed-death drug kinetics (Fig. 5). This is not surprising, as two bacterial protein synthesis inhibitors, azithromycin and doxycycline, have confirmed apicoplast targets and delayed death (33). Inhibiting apicoplast genome replication with drugs specifically targeting prokaryotic DNA gyrase activity, such as the fluoroquinolone ciprofloxacin (103, 104), or protein translation with any of four classes of antibacterials (chloramphenicol, tetracyclines, lincosamides, and macrolides) (12, 18, 33, 55, 94, 105–107) or a tRNA synthesis inhibitor (mupirocin) all cause characteristic delayed death and were confirmed to primarily target the apicoplast in our assays (Table 2 and Fig. 2 and 5) (37). Until now, rifampin has presented us with something of a conundrum. Rifampin inhibits transcription from the 35-kb apicoplast genome by targeting the plastid-encoded RNA polymerase (57) but causes immediate death that is not rescued by IPP (Table 2). Rifampin apparently has another target outside the apicoplast, which prevents us from drawing specific conclusions about the drug response kinetics of specifically inhibiting apicoplast transcription. Similarly, our inability to rescue parasites from the effects of the steroid fusidic acid imply a strong inhibition of a nonapicoplast target, likely the mitochondrion, which can also be targeted by this drug (59, 108).
Encouragingly, our IPP rescue assays successfully discriminate between aminoacyl-tRNA synthetase inhibitors, like mupirocin (an isoleucyl-tRNA synthetase inhibitor that specifically blocks apicoplast division [109, 110]) and borrelidin (a noncompetitive selective inhibitor first isolated from Streptomyces spp.) that binds to a unique hydrophobic cluster near the active sites of some bacterial and eukaryotic threonyl-tRNA synthetases. Plasmodium possesses only one threonyl-tRNA synthetase gene with isoforms trafficked to both the apicoplast and cytosol (111–114). IPP rescue indicates an exclusively apicoplast target for mupirocin but a target outside the apicoplast for borrelidin. This indicates that the cytosolic isoform of threonyl-tRNA synthetase is the primary target of borrelidin. The apicoplast isoform may also be inhibited, but the drug concentrations required for this inhibition are not significantly lower than those producing immediate death induced by targeting of cytosolic threonyl-tRNA synthetase isoform, so apicoplast-specific activity may be masked.
Actinonin clearly targets the apicoplast, based on our IPP rescue and previous data that showed an interruption of apicoplast development with exposure to this drug (69). Curiously, actinonin, whose primary target in bacteria is a posttranslational modification enzyme involved in protein synthesis, induces immediate death, whereas the other bona fide apicoplast housekeeping inhibitors all induce delayed death (Fig. 5). Actinonin resistance in Toxoplasma gondii is conferred by a point mutation in the apicoplast membrane-associated protein FtsH1, and an homologous target is inferred for P. falciparum with protease inhibition rather than posttranslational modification proposed as the mode of action (70).
Immediate-death apicoplast drugs, such as actinonin, are of particular therapeutic interest, as they circumvent the delayed action that limits the usefulness of many apicoplast drugs in clinical applications. Given that actinonin exhibits a different death kinetic from all the established apicoplast housekeeping inhibitors (Fig. 5), it is unlikely that it inhibits apicoplast housekeeping. Rather, treatment with actinonin appears to immediately impact apicoplast development and division, suggesting that it may target apicoplast biogenesis. Confirmation that the target of actinonin in malaria parasites is an apicoplast FtsH homologue (70) will be an important next step in dissecting the mode of action of a potentially significant new class of antimalarials with immediate death and a bona fide apicoplast target.
The IPP rescue assay facilitates the definitive identification of off-target compounds. We show clearly that IPP cannot rescue parasites from the effects of putative FASII inhibitors but can rescue parasites from inhibitors of the IPP synthesis pathway (Fig. 5). This is entirely consistent with the asexual blood stage essentiality of isoprenoid precursor synthesis (37) and the dispensability of apicoplast fatty acid biosynthesis at this life stage (31, 115–117), and it definitively shows that apicoplast fatty acid synthesis is not a drug target during the parasite asexual blood stages. IPP rescue assays thus present an efficient means of screening new compounds and will hopefully eliminate the misplaced effort in drug development that has occurred previously.
In sum, we show that simple IPP supplementation of in vitro-grown asexual P. falciparum readily identifies whether a compound has an exclusive target in the apicoplast, thus providing further intelligence about drugs already in use (35, 36) or in clinical trials (36, 118) in malaria patients. IPP supplementation can also inform us about new compounds, allowing deeper characterization of their mode of action and readily defining if they target the apicoplast.
MATERIALS AND METHODS
The drugs screened for apicoplast targets and controls are listed in Table 1.
P. falciparum cultures and chemical rescue assay.
P. falciparum D10 ACPLGFP parasites (53) were grown in human red blood cells at 2% hematocrit in RPMI 1640 supplemented with 0.5% AlbuMAX (GibcoBRL) and 0.1 μM pyrimethamine (Sigma) and gassed with 5% CO2 and 1% O2 in N2 at 37°C (53, 119). The apicoplasts of the parasite D10 ACPLGFP line express green fluorescent protein (GFP), which allowed us to visualize any disruption of the apicoplast by drug treatment. The half-maximal inhibitory concentration (IC50) of each drug was measured using tightly synchronized parasites and a standard SYBR green assay to determine immediate (48 h, 1st cycle) and any possible delayed (120 h, after 2nd cycle) death effects (120–122). The IC50 determined at 120 h of parasite growth inhibition was used in the IPP supplementation assays. The culture medium was supplemented with 200 μM IPP to assess whether parasite growth could be rescued from the inhibitors used (37). Supplementation continued for four (48-h) parasite life cycles, and samples were collected after each life cycle (48, 96, 144, and 192 h). Cultures otherwise received equivalent medium changes and addition of blood cells. Data were analyzed and graphs were generated using the GraphPad Prism software.
Parasites from 200 μl of culture were used to isolate genomic DNA (gDNA; Bioline Isolate II genomic DNA kit). We used primer pairs to specifically amplify apicoplast genes (PF3D7_API02900, elongation factor Tu), mitochondrial genes (mal_mito_3, cytochrome b), and nuclear genes (PF3D7_1252200, chitinase; PF3D7_0717700, serine-tRNA ligase [putative]; and PF3D7_1462800, glyceraldehyde-3-phosphate dehydrogenase) to determine the presence or absence of organelle genomes (Table 3) (37, 52). PCR mixtures contained template gDNA, 0.15 μM each primer, and 1× SensiMix (SYBR No-ROX kit). The reactions were performed at 56°C for primer annealing and 65°C for template extension for 39 cycles on a Bio-Rad qPCR machine. Relative quantification of target genes was determined using the comparative method (threshold cycle [2−ΔΔCT]) (37, 52, 123).
TABLE 3 Primer sequences used for quantitative real-time PCR
Parasites (D10 ACPLGFP) from 200 μl of culture were isolated by 0.15% saponin lysis, washed twice with phosphate-buffered saline (PBS) (pH 7.4), and resuspended in NuPAGE LDS sample buffer (Invitrogen). Parasite proteins were separated by SDS-PAGE and transferred to a nitrocellulose membrane (Amersham), and then the membrane was blocked in 5% skim milk powder in Tris buffer solution with 0.05% Tween 20 for at least 1 h at room temperature or overnight at 4°C. The membrane was probed with mouse anti-GFP (1:500; Roche) primary antibody, followed by goat anti-mouse-horseradish peroxidase (1:1,000; Thermo Scientific) to allow for detection with enhanced chemiluminescence Western blotting substrate (Pierce). Imaging was performed with a ChemiDoc (Bio-Rad).
Parasites (D10 ACPLGFP) were incubated in 2 μg/ml Hoechst 33342 stain for 10 min at 37°C. Cells in culture media were allowed to settle onto a glass slide (MakTek) coated with 1% polyethylenimine (Sigma), and they were then mounted with coverslips using fluorescence mounting medium (Dako) and imaged using a Leica DMIRB-TCS SP2 inverted confocal laser scanning microscope with a PL APO 63×/1.4 λBL oil immersion objective. Images were merged and global contrast adjusted where necessary using the ImageJ software package (124).
We thank the Australian Red Cross Blood Services, Melbourne, Australia, for supplying human erythrocytes. We also thank Stuart Ralph, University of Melbourne, for supplying borrelidin and Danny Wilson, University of Adelaide, for supplying erythromycin.
This work was supported by an Australian National Health and Medical Research Project Grant and an Australian Research Council Discovery Project Grant and Laureate Fellowship.
Gardner MJ, Hall N, Fung E, White O, Berriman M, Hyman RW, Carlton JM, Pain A, Nelson KE, Bowman S, Paulsen IT, James K, Eisen JA, Rutherford K, Salzberg SL, Craig A, Kyes S, Chan M-S, Nene V, Shallom SJ, Suh B, Peterson J, Angiuoli S, Pertea M, Allen J, Selengut J, Haft D, Mather MW, Vaidya AB, Martin DMA, Fairlamb AH, Fraunholz MJ, Roos DS, Ralph SA, McFadden GI, Cummings LM, Subramanian GM, Mungall C, Venter JC, Carucci DJ, Hoffman SL, Newbold C, Davis RW, Fraser CM, Barrell B. 2002. Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419:498–511.
Wilson IRJM, Denny PW, Preiser PR, Rangachari K, Roberts K, Roy A, Whyte A, Strath M, Moore DJ, Moore PW, Williamson DH. 1996. Complete gene map of the plastid-like DNA of the malaria parasite Plasmodium falciparum. J Mol Biol 261:155–172.
Gardner MJ, Bates PA, Ling IT, Moore DJ, McCready S, Gunasekera MBR, Wilson RJM, Williamson DH. 1988. Mitochondrial DNA of the human malarial parasite Plasmodium falciparum. Mol Biochem Parasitol 31:11–17.
Williamson DH, Gardner MJ, Preiser P, Moore DJ, Rangachari K, Wilson RJM. 1994. The evolutionary origin of the 35 kb circular DNA of Plasmodium falciparum: new evidence supports a possible rhodophyte ancestry. Mol Gen Genet 243:249–252.
Waller RF, Keeling PJ, Donald RG, Striepen B, Handman E, Lang-Unnasch N, Cowman AF, Besra GS, Roos DS, McFadden GI. 1998. Nuclear-encoded proteins target to the plastid in Toxoplasma gondii and Plasmodium falciparum. Proc Natl Acad Sci U S A 95:12352–12357.
Jomaa H, Wiesner J, Sanderbrand S, Altincicek B, Weidemeyer C, Hintaz M, Turbachova I, Eberl M, Zeidler J, Lichtenthaler HK, Soldati D, Beck E. 1999. Inhibitors of the nonmevalonate pathway of isoprenoid biosynthesis as antimalarial drugs. Science 285:1573–1576.
Hoen R, Novoa EM, López A, Camacho N, Cubells L, Vieira P, Santos M, Marín-García P, Bautista JM, Cortés A, Ribas de Pouplana L, Royo M. 2013. Selective inhibition of an apicoplastic aminoacyl-tRNA synthetase from Plasmodium falciparum. Chembiochem 14:499–509.
Ke H, Sigala PA, Miura K, Morrisey JM, Mather MW, Crowley JR, Henderson JP, Goldberg DE, Long CA, Vaidya AB. 2014. The heme biosynthesis pathway is essential for Plasmodium falciparum development in mosquito stage but not in blood stages. J Biol Chem 289:34827–34837.
Sidhu ABS, Sun Q, Nkrumah LJ, Dunne MW, Sacchettini JC, Fidock DA. 2007. In vitro efficacy, resistance selection, and structural modeling studies implicate the malarial parasite apicoplast as the target of azithromycin. J Biol Chem 282:2494–2504.
Bowman JD, Merino EF, Brooks CF, Striepen B, Carlier PR, Cassera MB. 2014. Antiapicoplast and gametocytocidal screening to identify the mechanisms of action of compounds within the malaria box. Antimicrob Agents Chemother 58:811–819.
Yao Z-K, Krai PM, Merino EF, Simpson ME, Slebodnick C, Cassera MB, Carlier PR. 2015. Determination of the active stereoisomer of the MEP pathway-targeting antimalarial agent MMV008138, and initial structure-activity studies. Bioorganic Med Chem Lett 25:1515–1519.
Korsinczky M, Chen N, Kotecka B, Saul A, Rieckmann K, Cheng Q. 2000. Mutations in Plasmodium falciparum cytochrome b that are associated with atovaquone resistance are located at a putative drug-binding site. Antimicrob Agents Chemother 44:2100–2108.
Zeidler J, Schwender J, Müller C, Wiesner J, Weidemeyer C, Beck E, Jomaa H, Lichtenthaler HK. 1998. Inhibition of the non-mevalonate 1-deoxy-d-xylulose-5-phosphate pathway of plant isoprenoid biosynthesis by fosmidomycin. Zeitschrift für Naturforschung C 53:980–986.
Kuzuyama T, Shimizu T, Takahashi S, Seto H. 1998. Fosmidomycin, a specific inhibitor of 1-deoxy-d-xylulose 5-phosphate reductoisomerase in the nonmevalonate pathway for terpenoid biosynthesis. Tetrahedron Lett 39:7913–7916.
Steinbacher S, Kaiser J, Eisenreich W, Huber R, Bacher A, Rohdich F. 2003. Structural basis of fosmidomycin action revealed by the complex with 2-C-methyl-d-erythritol 4-phosphate synthase (IspC). Implications for the catalytic mechanism and anti-malaria drug development. J Biol Chem 278:18401–18407.
Zhang B, Watts KM, Hodge D, Kemp LM, Hunstad DA, Hicks LM, Odom AR. 2011. A second target of the antimalarial and antibacterial agent fosmidomycin revealed by cellular metabolic profiling. Biochemistry 50:3570–3577.
Wiley JD, Merino EF, Krai PM, McLean KJ, Tripathi AK, Vega-Rodríguez J, Jacobs-Lorena M, Klemba M, Cassera MB. 2015. Isoprenoid precursor biosynthesis is the essential metabolic role of the apicoplast during gametocytogenesis in Plasmodium falciparum. Eukaryot Cell 14:128–139.
van Dooren GG, Su V, D'Ombrain MC, McFadden GI. 2002. Processing of an apicoplast leader sequence in Plasmodium falciparum and the identification of a putative leader cleavage enzyme. J Biol Chem 277:23612–23619.
Dar MA, Sharma A, Mondal N, Dhar SK. 2007. Molecular cloning of apicoplast-targeted Plasmodium falciparum DNA gyrase genes: unique intrinsic ATPase activity and ATP-independent dimerization of PfGyrB subunit. Eukaryot Cell 6:398–412.
Johnson RA, McFadden GI, Goodman CD. 2011. Characterization of two malaria parasite organelle translation elongation factor G proteins: the likely targets of the anti-malarial fusidic acid. PLoS One 6:e20633.
Yoon HJ, Kim HL, Lee SK, Kim HW, Kim HW, Lee JY, Mikami B, Suh SW. 2004. Crystal structure of peptide deformylase from Staphylococcus aureus in complex with actinonin, a naturally occurring antibacterial agent. Proteins 57:639–642.
Tonkin CJ, van Dooren GG, Spurck TP, Struck NS, Good RT, Handman E, Cowman AF, McFadden GI. 2004. Localization of organellar proteins in Plasmodium falciparum using a novel set of transfection vectors and a new immunofluorescence fixation method. Mol Biochem Parasitol 137:13–21.
Goodman CD, McFadden GI. 2014. Ycf93 (Orf105), a small apicoplast-encoded membrane protein in the relict plastid of the malaria parasite Plasmodium falciparum that is conserved in Apicomplexa. PLoS One 9:e91178.
Tasdemir D, Kaiser M, Brun R, Yardley V, Schmidt TJ, Tosun F, Ruedi P. 2006. Antitrypanosomal and antileishmanial activities of flavonoids and their analogues: in vitro, in vivo, structure-activity relationship, and quantitative structure-activity relationship studies. Antimicrob Agents Chemother 50:1352–1364.
Tasdemir D, Lack G, Brun R, Ruedi P, Scapozza L, Perozzo R. 2006. Inhibition of Plasmodium falciparum fatty acid biosynthesis: evaluation of FabG, FabZ, and FabI as drug targets for flavonoids. J Med Chem 49:3345–3353.
Wickramasinghe SR, Inglis KA, Urch JE, Müller S, van Aalten DMF, Fairlamb AH. 2006. Kinetic, inhibition and structural studies on 3-oxoacyl-ACP reductase from Plasmodium falciparum, a key enzyme in fatty acid biosynthesis. Biochem J 393:447–457.
Tarun AS, Peng X, Dumpit RF, Ogata Y, Silva-Rivera H, Camargo N, Daly TM, Bergman LW, Kappe SHI. 2008. A combined transcriptome and proteome survey of malaria parasite liver stages. Proc Natl Acad Sci U S A 105:305–310.
Tonkin CJ, Struck NS, Mullin KA, Stimmler LM, McFadden GI. 2006. Evidence for Golgi-independent transport from the early secretory pathway to the plastid in malaria parasites. Mol Microbiol 61:614–630.
Hager KM, Striepen B, Tilney LG, Roos DS. 1999. The nuclear envelope serves as an intermediary between the ER and Golgi complex in the intracellular parasite Toxoplasma gondii. J Cell Sci 112:2631–2638.
Ramya TNC, Mishra S, Karmodiya K, Surolia N, Surolia A. 2007. Inhibitors of nonhousekeeping functions of the apicoplast defy delayed death in Plasmodium falciparum. Antimicrob Agents Chemother 51:307–316.
Beckers CJ, Roos DS, Donald RG, Luft BJ, Schwab JC, Cao Y, Joiner KA. 1995. Inhibition of cytoplasmic and organellar protein synthesis in Toxoplasma gondii. Implications for the target of macrolide antibiotics. J Clin Invest 95:367–376.
Burkhardt D, Wiesner J, Stoesser N, Ramharter M, Uhlemann A-C, Issifou S, Jomaa H, Krishna S, Kremsner PG, Borrmann S. 2007. Delayed parasite elimination in human infections treated with clindamycin parallels “delayed death” of Plasmodium falciparum in vitro. Int J Parasitol 37:777–785.
Gupta A, Mir SS, Jackson KE, Lim EE, Shah P, Sinha A, Siddiqi MI, Ralph SA, Habib S. 2013. Recycling factors for ribosome disassembly in the apicoplast and mitochondrion of Plasmodium falciparum. Mol Microbiol 88:891–905.
Khan S, Sharma A, Jamwal A, Sharma V, Pole AK, Thakur KK, Sharma A. 2011. Uneven spread of cis- and trans-editing aminoacyl-tRNA synthetase domains within translational compartments of P. falciparum. Sci Rep 1:188.
Jackson KE, Pham JS, Kwek M, De Silva NS, Allen SM, Goodman CD, McFadden GI, Ribas de Pouplana L, Ralph SA. 2012. Dual targeting of aminoacyl-tRNA synthetases to the apicoplast and cytosol in Plasmodium falciparum. Int J Parasitol 42:177–186.
Ruan B, Bovee ML, Sacher M, Stathopoulos C, Poralla K, Francklyn CS, Soll D. 2004. A unique hydrophobic cluster near the active site contributes to differences in borrelidin inhibition among threonyl-tRNA synthetases. J Biol Chem 280:571–577.
Shears MJ, MacRae JI, Mollard V, Goodman CD, Sturm A, Orchard LM, Llinás M, McConville MJ, Botté CY, McFadden GI. 2017. Characterization of the Plasmodium falciparum and P. berghei glycerol 3-phosphate acyltransferase involved in FASII fatty acid utilization in the malaria parasite apicoplast. Cell Microbiol 19:e12633.
van Schaijk BCL, Kumar TRS, Vos MW, Richman A, van Gemert GJ, Li T, Eappen AG, Williamson KC, Morahan BJ, Fishbaugher M, Kennedy M, Camargo N, Khan SM, Janse CJ, Sim KL, Hoffman SL, Kappe SHI, Sauerwein RW, Fidock DA, Vaughan AM. 2014. Type II fatty acid biosynthesis is essential for Plasmodium falciparum sporozoite development in the midgut of anopheles mosquitoes. Eukaryot Cell 13:550–559.
Vaughan AM, O'Neill MT, Tarun AS, Camargo N, Phuong TM, Aly ASI, Cowman AF, Kappe SHI. 2009. Type II fatty acid synthesis is essential only for malaria parasite late liver stage development. Cell Microbiol 11:506–520.
Smilkstein M, Sriwilaijaroen N, Kelly JX, Wilairat P, Riscoe M. 2004. Simple and inexpensive fluorescence-based technique for high-throughput antimalarial drug screening. Antimicrob Agents Chemother 48:1803–1806.
Gunjan S, Singh SK, Chauhan BS, Pandey SK, Ahmad H, Dwivedi AK, Tripathi R. 2015. Clarithromycin enhances the antimalarial efficacy of mefloquine via its increased bioavailability and disrupting P. falciparum apicoplast. Life Sci 136:126–132.
Bracho C, Dunia I, Romano M, Benedetti EL, Perez HA. 2001. Plasmodium vivax and Plasmodium chabaudi: intraerythrocytic traffic of antigenically homologous proteins involves a brefeldin A-sensitive secretory pathway. Eur J Cell Biol 80:164–170.
If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download.