ABSTRACT
In the malaria parasite Plasmodium falciparum, synthesis of isoprenoids from glycolytic intermediates is essential for survival. The antimalarial fosmidomycin (FSM) inhibits isoprenoid synthesis. In P. falciparum, we identify a loss-of-function mutation in HAD2 (PF3D7_1226300) as necessary for FSM resistance. Enzymatic characterization reveals that HAD2, a member of the haloacid dehalogenase-like hydrolase (HAD) superfamily, is a phosphatase. Harnessing a growth defect in resistant parasites, we select for suppression of HAD2-mediated FSM resistance and uncover hypomorphic suppressor mutations in the locus encoding the glycolytic enzyme phosphofructokinase (PFK9). Metabolic profiling demonstrates that FSM resistance is achieved via increased steady-state levels of MEP pathway and glycolytic intermediates and confirms reduced PFK9 function in the suppressed strains. We identify HAD2 as a novel regulator of malaria parasite metabolism and drug sensitivity and uncover PFK9 as a novel site of genetic metabolic plasticity in the parasite. Our study informs the biological functions of an evolutionarily conserved family of metabolic regulators and reveals a previously undescribed strategy by which malaria parasites adapt to cellular metabolic dysregulation.
IMPORTANCE Unique and essential aspects of parasite metabolism are excellent targets for development of new antimalarials. An improved understanding of parasite metabolism and drug resistance mechanisms are urgently needed. The antibiotic fosmidomycin targets the synthesis of essential isoprenoid compounds from glucose and is a candidate for antimalarial development. Our study identifies a novel mechanism of drug resistance and further describes a family of metabolic regulators in the parasite. Using a novel forward genetic approach, we also uncover mutations that suppress drug resistance in the glycolytic enzyme PFK9. Thus, we identify an unexpected genetic mechanism of adaptation to metabolic insult that influences parasite fitness and tolerance to antimalarials.
Introduction
Malaria remains a global health threat, infecting 216 million people per year and causing nearly half a million deaths, mainly of pregnant women and young children (1). Resistance to current therapies has limited control efforts for malaria (2, 3). New drugs and a deeper understanding of drug resistance mechanisms are urgently needed.
Malaria is caused by infection with unicellular eukaryotic parasites of the genus Plasmodium. The species Plasmodium falciparum is responsible for most life-threatening malarial disease. As an obligate intracellular parasite of human erythrocytes, Plasmodium falciparum has unique metabolic features that may be exploited to discover new drug targets and develop new therapies. In the red blood cell niche, Plasmodium parasites are highly dependent on glucose metabolism. Infection with Plasmodium spp. results in a nearly 100-fold increase in glucose import in red blood cells (4–6). Despite these energy requirements, the parasite demonstrates little aerobic respiration via the TCA cycle. Instead, it relies on anaerobic glycolysis to produce ATP (7–10).
Besides ATP production, glucose also has a number of anabolic fates in P. falciparum. One such fate is the synthesis of isoprenoids. Isoprenoids are a large class of hydrocarbons with extensive structural and functional diversity (11). In the malaria parasite, isoprenoids perform several essential functions, including protein prenylation, dolichylation, and synthesis of GPI anchors (12–14). Despite this diversity, all isoprenoids are synthesized from a common five-carbon building block, isopentyl pyrophosphate (IPP). Evolution has produced two distinct routes for IPP synthesis: the mevalonate pathway, found in archaea, fungi, animals, and the cytoplasm of plants; and the methylerythritol phosphate (MEP) pathway, found in most eubacteria, plant chloroplasts, and apicomplexan parasites such as P. falciparum (15). Because it is both essential for the parasite and absent from the human host, the MEP pathway is a compelling target for antimalarial development. The antibiotic and antimalarial fosmidomycin (FSM) is a competitive inhibitor of the first committed enzymatic step of the MEP pathway, catalyzed by 1-deoxy-D-xylulose-5- phosphate reductoisomerase (DXR, E.C. 1.1.1.267) (16–18). FSM has been validated as a specific inhibitor of the MEP pathway in P. falciparum (19) and is a valuable chemical tool to study MEP pathway biology and essential metabolism in the parasite. In this study, we find that FSM is also a useful tool for probing glycolytic metabolism upstream of the essential MEP pathway.
Parasites are likely to control the proportion of glucose used for energy production versus production of secondary metabolites, such as isoprenoids. We previously used a screen for FSM resistance to identify HAD1, a metabolic regulator whose loss results in increased levels of MEP pathway intermediates and resistance to MEP pathway inhibition. HAD1 is a cytoplasmic sugar phosphatase that dephosphorylates a number of sugar phosphate intermediates upstream of the MEP pathway (20, 21). HAD1 belongs to the haloacid dehalogenase-like hydrolase (HAD) enzyme superfamily and more specifically, the IIB and Cof-like hydrolase subfamilies (22). While HADs are found in all kingdoms of life, HAD1 is most related to bacterial members of this superfamily (20, 23), which have been implicated in metabolic regulation, stress response, and phosphate homeostasis (24–28). However, most members of this superfamily remain uncharacterized.
In this study, we describe the discovery of HAD2, a second HAD family member in P. falciparum. We find that HAD2 is a cytosolic phosphatase required for metabolic homeostasis. Loss of HAD2 dysregulates glycolysis and misroutes metabolites toward the MEP pathway, conferring drug resistance. In our study, we harness a fitness defect in had2 parasite strains to employ an innovative screen for suppression of drug resistance in the parasite. Selection for suppression of drug resistance identifies mutations in PFK9, which encodes the canonical glycolytic regulatory enzyme, phosphofructokinase. Reduction in PFK9 activity rescues the metabolic dysregulation in our resistant mutants and restores FSM sensitivity. Our unique approach thus reveals PFK9 as a site of exceptional metabolic plasticity in the parasite and uncovers a novel genetic mechanism by which P. falciparum malaria parasites may adapt to metabolic stress and drug selective pressure.
This article was submitted to an online preprint archive (29).
RESULTS
A FSMR strain possesses a nonsense allele of HAD2, homolog of the MEP pathway regulator HAD1
The MEP pathway is responsible for the synthesis of the essential isoprenoid precursors isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP). This pathway is specifically inhibited by the antibiotic FSM (19, 30, 31). We previously generated P. falciparum strains resistant to FSM. Mutations in HAD1 (PF3D7_1033400) cause the resistance phenotype in a majority of these strains (20). However, the genetic and biochemical basis of FSM resistance in strain E2 remained unknown. As previously reported, we find that E2 is less sensitive to FSM than its wild-type 3D7 parental line (Fig. 1A) (20). Strain E2 has a FSM IC50 of 4.8 ± 1.2 μM, significantly greater than that of its parent strain (0.9 ± 0.06 μM) (p≤0.01, unpaired Student’s t-test).
We find that this resistance phenotype is not due to changes in expression of the genes encoding the first two, rate-limiting, steps of the MEP pathway, DXS and DXR (32–35). In addition, strain E2 does not have genetic changes in the known FSM resistance locus and MEP pathway regulator, HAD1, nor are there changes in HAD1 expression by immunoblot (Figs. S1A and S1B).
To identify new genetic changes that may result in FSM resistance, we performed whole genome sequencing on strain E2 and identified an A469T mutation in PF3D7_1226300 (PlasmoDB ID), hereafter referred to as HAD2 (36). Variant data for strains sequenced in this study can be found in Dataset S1. Sanger sequencing of the HAD2 locus in strain E2 confirmed the presence of the A469T SNP. The A469T single nucleotide polymorphism (SNP) yields a truncated (R157X) protein variant and therefore we expect HAD2 function is lost in strain E2. Interestingly, HAD2 is a close homolog of the known MEP pathway regulator, the sugar phosphatase HAD1 (20). Sequence homology places both proteins in the haloacid dehalogenase-like hydrolase (HAD) superfamily and further within the IIB and Cof-like hydrolase subfamilies (Interpro IPR006379 and IPR000150, respectively) (22). While no structural information exists for P. falciparum HAD2, the structure of the Plasmodium vivax HAD2 (PVX_123945, PvHAD2) has been solved (PDB ID 2B30). PvHAD2 (93% identical and 98% similar to PfHAD2) contains the common structural motifs found in other HADs, including a core and cap domain (Fig. 1B). HAD2 possesses the four conserved sequence motifs found in HAD proteins (Fig. 1C), which are involved in binding of the substrate, coordination of the phosphoryl group and Mg2+ ion, and hydrolysis of the substrate phosphate (37–39). Overall, HAD2 and HAD1 protein sequences share ~29% sequence identity and ~53% sequence similarity (Fig. 1C). We hypothesized that HAD2, like HAD1, regulates metabolism in P. falciparum, and that loss of HAD2-mediated metabolic control was responsible for FSM-resistance in malaria parasite strain E2.
HAD2 is a functional phosphometabolite phosphatase
We have previously established that P. falciparum HAD1 is a promiscuous sugar phosphatase, with activity against a wide range of phosphometabolites. Similarly, P. vivax HAD2 has been enzymatically characterized and found to possess phosphatase activity against various monophosphorylated substrates, including glycerol 2-phosphate (glc2P) and pyridoxal phosphate (PLP) (40). Recombinant PvHAD2 also utilizes additional monophosphorylated substrates, such as adenosine 5’-monophosphate (AMP) and glycerol 1-phosphate (glc1P), with moderate activity.
Based on the previous characterization of a close Plasmodium homolog, as well as sequence homology to HAD1 and other HAD proteins, we predicted that PfHAD2 would also function enzymatically as a phosphatase. We successfully purified recombinant PfHAD2 in E. coli and confirmed the phosphatase activity of recombinant PfHAD2 using para-nitrophenyl phosphate (pNPP), a promiscuous, chromogenic phospho-substrate (Fig. 2A) (23, 41). Because E. coli expresses a number of HAD-like phosphatases (23), we confirmed that the phosphatase activity was specific to purified PfHAD2 by expression and purification of a catalytically inactive mutant (HAD2D26A). The Asp26 residue was chosen for mutagenesis as the corresponding residue in PfHAD1 (Asp27) has been previously shown to be required for catalysis (21).
We also established the activity of PfHAD2 against a panel of phosphorylated substrates and determined that its substrate profile closely mirrors that of PvHAD2 (Fig. 2B). Overall, we find that PfHAD2 is a phosphatase with activity against small phosphosubstrates, such as pNPP and glc2P. These data suggest that, like HAD1 and related HADs in microbes and plants (23, 42–44), HAD2 is a promiscuous phosphatase with a preference for a variety of monophosphorylated phosphometabolites.
In vitro evolution of mutations suppressing FSM resistance
During routine culturing of E2 FSMR parasites, we observed that the E2 strain is growth-attenuated compared to its parental parasite strain. Surprisingly, during prolonged culture in the absence of FSM, this growth phenotype resolved, and improved growth rates correlated with a return to FSM sensitivity (Fig. 3A). From these observations, we hypothesized that had2R157X-mediated FSM resistance led to a fitness cost in cultured parasites. We sought to harness this fitness cost to drive in vitro evolution of a FSM-sensitive (FSMS) population, possessing additional, novel mutations that might suppress FSM resistance in had2R157X parasite strains.
FSM-resistant strain E2 was cultured through multiple passages in the absence of FSM selection. Through limiting dilution, we derived five E2-based clones in the absence of drug pressure (Fig. 3B). Three of the five clones remained FSMR (designated clones E2-R1, -R2, and -R3), but two of these were found to be FSMS (designated E2-S1 and -S2) (Fig. 3B and 4A). To validate our novel suppressor screen approach, we independently repeated this genetic selection with the three FSMR E2 clones by again culturing in the absence of FSM for >1 month (Fig. 3B). As before, these strains (E2-S3, -S4, and -S5) also lost their FSM resistance phenotype (Fig. 4A).
Consistent with our initial observation that our had2R157X FSM resistant strain grows poorly, we find that the FSMR clone E2-R1 grows at a significantly reduced rate compared to the parental strain, while two FSMS clones (E2-S1 and E2-S3) have restored growth rates, similar to that of wild-type (Fig. 4B).
Loss of FSM resistance might have occurred by reversion of the had2R157X mutation in E2-derived strains. Instead, we find that all E2-SX clones maintained loss of HAD2 via the had2R157X mutation. We hypothesized that the FSMS E2 clones, driven by a fitness advantage, had acquired new suppressor mutation(s) at an additional locus. We performed whole genome sequencing on the original five E2 clones to identify any genetic changes that segregated with FSM-sensitivity. Sequencing revealed that a new mutation (C3617T) in the locus encoding phosphofructokinase-9 (PFK9, PF3D7_0915400) is present in both suppressed (FSMS) E2 clones and none of the three FSMR E2 clones (Fig. 3B, Dataset S1). The C3617T mutation results in a PFK9T1206 protein variant. PFK9 contained the only SNP that segregated with the change in FSM tolerance. Two other loci had indels that also segregated with our FSM phenotype. These loci encode a tyrosine recombinase (MAL13P1.42) (45) and an erythrocyte surface protein (PIESP1, PFC0435w). Given their predicted functions and the presence of A/T indels in poly-A and poly-T tracts, we concluded that mutations in these loci were unlikely to result in our suppressed phenotype and prioritized PFK9 as the likely locus of our suppressor mutation.
To verify whether mutations in PFK9 were responsible for suppressing FSM resistance in all of our suppressed strains, we investigated HAD2 and PFK9 in the E2-S3, -S4, and -S5 strains, which were derived through independent evolution of the E2-R1, -R2, and -R3 populations in the absence of FSM. By Sanger sequencing, we find that, as before, all strains maintain the had2R157X mutation and acquire new, independent PFK9 mutations (Fig. 4A). The independent acquisition of four different PFK9 alleles during selection, each of which was associated with both improved growth and loss of FSM sensitivity, strongly indicates that loss of PFK9 function is responsible for these phenotypes in strains lacking HAD2.
Loss of HAD2 is necessary for FSM-resistance in had2R157X parasites
HAD2 was not the sole genetic change in FSMR strain E2. In addition, because intraerythrocytic P. falciparum parasites are haploid, we cannot distinguish recessive from dominant or gain-of-function mutations. Therefore, we sought to establish whether restoring HAD2 expression in trans in a had2R157X strain would restore FSM sensitivity. Using a previously described expression system enabled by the piggyBac transposase (20, 46), we expressed HAD2-GFP driven by the heat shock protein 110 (Hsp110) promoter (47). We confirmed that the transfected had2R157X E2-R2 clone maintains the had2R157X allele at the endogenous locus and successfully expresses HAD2-GFP (Fig. 5A). Expression of HAD2-GFP in had2R157X parasites results in restoration of FSM sensitivity (Fig. 5B), confirming that loss of HAD2 is necessary for FSM resistance in this strain. The resistant clone (E2-R2) has a FSM IC50 of 3.9 ± 0.2 μM, significantly higher than that of the wild-type parent strain (0.9 ± 0.06 μM, p≤0.001, one-way ANOVA and Sidak’s post-test). Expression of HAD2-GFP in E2-R2 results in an IC50 of 0.6 ± 0. 02 μM for FSM, significantly lower than the E2-R2 strain (p≤0.001) but not significantly different than the parental strain (p>0.5).
Using fluorescence microscopy, we also investigated the localization of HAD2-GFP in our E2-R2 Hsp110:HAD2-GFP strain. We observe that HAD2-GFP is diffusely present throughout the cytoplasm in asexual P. falciparum trophozoites and schizonts but excluded from the digestive food vacuole (Fig. S2). This finding is consistent with the lack of a predicted signal sequence for HAD2 using SignalP, PlasmoAP, and PlasMit algorithms (48–50).
PFK9 mutations in suppressed strains are hypomorphic
The PFK9 locus encodes the enzyme phosphofructokinase (PFK, E.C. 2.7.11), which catalyzes the first committed and canonically rate-limiting step of glycolysis, which is the conversion of fructose 6-phosphate to fructose 1,6-bisphosphate. PFK9 is comprised of two domains, alpha (α) and beta (β), which are typically encoded by independent genes in non-apicomplexans (51) (Fig. 6A). While in other systems the a domain is regulatory, previous work on P. falciparum PFK9 has demonstrated catalytic activity for both domains (51–55).
Of the four PFK9 variants identified in this study, three variants map to the α domain, while one variant (S335L) maps to the β domain (Fig. 6A). We projected our mutations on a three-dimensional model of PfPFK9 to reveal a possible structural basis for altered PFK function. Three variants (S335L, T1206I, and S1267L) align and model to currently available crystal structures of PFK, while a fourth allele (N1359Y) does not. While three out of four mutations map to the α domain of PfPFK9 (T1206I, S1267L, N1359Y), these mutations do not appear to cluster in any particular region. All mutations affect amino acid residues that are physically distant from the substrate-binding pocket of either domain and are not predicted to impact binding or specific catalytic residues.
Consistent with a previous study on PfPFK9 (51), attempts to purify recombinant full-length PFK9 were unsuccessful. To then understand the enzymatic impacts of our PFK9 variants, we quantified the native PFK specific activity in P. falciparum (51, 56) (Fig. 6B, Fig. S3). Lysates from strains possessing PFK9 mutations (E2-SX strains) have markedly reduced specific activity of PFK compared to the parental strain (Fig. 6B). Combined with the diverse mutation locations (Fig. 6A), the reduced lysate PFK activity in E2-SX strains suggests that a variety of genetic strategies to alter PFK function can lead to resistance suppression.
Metabolic profiling reveals mechanisms of resistance and suppression in HAD2 and PFK9 mutant parasites
Reduced activity of PFK9, which catalyzes the canonical rate-limiting step in glycolysis, is associated with restored FSM sensitivity to had2 mutant strains. Therefore, we anticipated that metabolic changes might underlie both resistance and suppression in our E2 clones. We performed targeted metabolic profiling on the parental parasite strain as well as E2 clones R1-R3 and S1-S2 (Fig. 7A, Table S1). We find that levels of the MEP pathway intermediate DOXP are significantly increased in FSMR (had2R157X, PFK9WT) strains (Fig. 7A, p≤0.05, one-way ANOVA and Sidak’s post-test). FSM is competitive with DOXP for inhibition of its target enzyme DXR. Therefore, our data indicate that the FSMR strains achieve resistance via increased levels of DOXP, which outcompetes FSM. We also observe a significant increase in the downstream MEP metabolite, MEcPP, in our FSMR strains (p≤0.05).
To understand the role of PFK9 in conferring and suppressing FSM resistance, we determined the steady-state levels of intermediates from glycolysis, metabolites of which feed into the MEP pathway (Fig. 7A, Table S1). Hierarchical clustering indicates that resistant clones are characterized by a metabolic signature of increased levels of FBP, DOXP, and MEcPP (Fig. 7A). We observe that the abundance of DOXP and MEcPP are tightly correlated with cellular levels of the PFK9 product, FBP (Fig. 7B, p<0.01), but not with the other upstream glycolytic metabolites, such as glu6P/fru6P (Fig. 7C, p>0.2).
Of note, the pfk9T1206l suppressor allele in strains S1 and S2 restores nearly parental levels of FBP and downstream MEP pathway intermediates (Fig. 7A), consistent with our finding that PFK activity is reduced in lysate from these strains (Fig. 6B).
DISCUSSION
Cells must control levels of critical metabolites in order to efficiently utilize carbon sources for energy and biosynthesis of essential molecules. Cells may regulate their metabolism via transcriptional, post-transcriptional, post-translational, allosteric, or enzymatic mechanisms that are necessary for growth (57–60). In the glucose-rich red blood cell niche, Plasmodium spp. malaria parasites display a unique dependence on glycolysis for energy and biosynthesis.
Using resistance to a metabolic inhibitor, we identify a phosphatase member of the HAD superfamily, HAD2, as a novel regulator of metabolism in P. falciparum. Importantly, HAD2 controls substrate availability to the parasite-specific MEP pathway for isoprenoid synthesis, which is a promising drug target for much needed new antimalarials. We find that tolerance to inhibitors such as FSM is a robust and sensitive readout of metabolic perturbation. HAD2 is necessary for metabolic homeostasis in malaria parasites. Cells lacking HAD2 exhibit marked dysregulation of central carbon metabolism, including altered steady-state levels of glycolytic intermediates and isoprenoid precursors. We find that mutations in phosphofructokinase (PFK9) restore wild-type growth rates and FSM sensitivity to our had2 mutant strains. Our study thus genetically connects the function of HAD2, a HAD superfamily member, to control of essential central carbon metabolism. In addition, our work reveals a previously undescribed strategy by which malaria parasites may respond to cellular metabolic dysregulation through mutation in the gene encoding the rate-limiting glycolytic enzyme PFK9.
HAD2 is a member of the HAD superfamily and a homolog of the previously described metabolic regulator HAD1. With our previous studies on HAD1 (20, 21), we define the cellular role of these proteins in P. falciparum and contribute to the greater understanding of the HADs, an evolutionarily conserved and widespread protein family. Both enzymes belong to the IIB (IPR006379) and Cof-like hydrolase (IPR000150) subfamilies (22). HAD enzymes display diverse substrate preferences (23, 27, 43, 44, 61–64), and their biological functions are largely unknown. Like other HAD homologs, including PfHAD1 (20), HAD2 appears to be a non-specific, cytoplasmic phosphatase with preference for small, monophosphorylated substrates. While the HAD superfamily is thought to be highly evolvable pool of enzymes with broad substrate specificity (37, 42), our work strongly suggests that, like HAD1 and HAD2, other members of this superfamily are likely to perform specific and biologically important cellular functions. As P. falciparum has a smaller repertoire of HADs than bacteria, and Plasmodium HADs influence easily quantified phenotypes (drug tolerance, growth, metabolite levels), the malaria parasite may be an attractive system for study of the molecular mechanisms by which HAD proteins control metabolic homeostasis and growth.
Metabolic profiling reveals that loss of HAD2 function leads to metabolic dysregulation, which is centered around the canonical rate-limiting step of glycolysis, catalyzed by PFK9. While the cellular abundance of the PFK9 product FBP is increased in had2 parasites, HAD2 does not directly utilize FBP as an enzymatic substrate, suggesting an indirect mechanism of HAD2-mediated metabolic regulation. However, the distinct metabolic signature of had2 parasites, characterized by increased levels of the MEP pathway metabolites DOXP and MEcPP and the key glycolytic metabolite FBP, suggests that MEP pathway metabolism is precisely linked to FBP production. In other microbial systems, FBP levels reflect metabolic flux and are cued to environmental perturbations (65). FBP-centered metabolic regulation is also important to the related apicomplexan Toxoplasma gondii, which constitutively expresses the fructose 1,6- bisphosphatase (FBPase) to fine-tune glucose metabolism (57). While P. falciparum does not appear to possess an FBPase, necessary for gluconeogenesis, the parasite may possess alternative FBP-sensing mechanisms to tune metabolism, perhaps via regulators such as HAD1 and HAD2.
The metabolic dysregulation we observe in the had2 mutant strains appears to come with a fitness disadvantage. Under FSM-selective pressure, the benefits of dysregulated metabolism outweigh the costs. However, in the absence of FSM, had2 parasites achieve metabolic relief through secondary mutation in PFK9. The improved growth of had2 pfk9 double mutant parasites, compared to parasites with a had2 single mutation, argues that both the growth and metabolic phenotypes are linked. However, our complementation studies cannot strictly discern whether restoring HAD2 directly improves the growth rate of the had2 strain, as transfection and complementation of HAD2 is inherently an additional selection for increased fitness. Of note, two recent essentiality screens in Plasmodium spp. have found that HAD2 is dispensable for growth, and loss of HAD2 was not found to be associated with any significant fitness defect (66, 67). However, it is unknown whether the mutant strains generated in these screens have also acquired additional suppressor mutations, such as polymorphisms in PFK9, that have facilitated their growth.
Likewise, PFK9 provides an additional case-study of the context dependence of gene essentiality in Plasmodium spp. As the canonical rate-limiting step in glycolysis, PFK9 is strongly predicted to be essential for asexual growth of malaria parasites (66, 67). In the context of a had2 mutation, our strains readily develop mutations in PFK9 that reduce function but are nonetheless associated with increased fitness. Indeed, it is surprising that parasites that are entirely dependent on glycolysis for ATP production tolerate such a significant loss of activity in this enzyme. Because we identify mutations across the length of PFK9 in our suppressed strains, our studies do not appear to point to a specific disrupted function, such as alterations in an allosteric binding pocket or a dimer interface. The observed mutability of PFK9 points to a remarkable and unexpected metabolic plasticity in the parasite. That is, even though the parasite inhabits a highly controlled intraerythrocytic niche, a wide range of metabolic states of P. falciparum growth are still permissive for parasite growth. This previously undescribed metabolic plasticity centered on PFK9 should be considered in future efforts to target essential metabolism in Plasmodium.
Combined with the above study, our work highlights the central role of the glycolytic enzyme PFK9. A recent kinetic model of parasite glycolysis confirms that PFK has a high flux-control coefficient, is sensitive to competitive inhibition, and can effectively reduce glycolytic flux (68, 69). Like HAD2, PFK9 is plant-like and evolutionarily divergent from its mammalian homologs (51). These differences may be exploited for PFK inhibitor design and may indicate that PFK9 can be specifically targeted for antimalarial development. However, our work cautions that the parasite has a surprising capacity to adapt to perturbations in central carbon metabolism, which may present challenges when targeting these pathways.
Finally, our approach demonstrates the power of forward genetics to uncover novel biology in a clinically relevant, non-model organism. We employ a previously described screen (20) to uncover a novel resistance locus and employ a second selection for parasite fitness to identify changes that suppress our resistance phenotype. Of the 19 strains in our original FSM resistance screen (20), we identify only one had2 mutant, likely due to the reduced fitness associated with resistance in this strain. While fitness costs associated with antimalarial resistance are well known (70–73), this study represents, to our knowledge, the first to harness this evolutionary trade-off to identify suppressor mutations in a non-target locus. Additional methods to identify low fitness resistant mutants have recently been recently described (73), and fitness assessment of resistance mutations may allow for suppressor screening for other antimalarials or other target pathways to reveal new aspects of biology and drug resistance in malaria parasites.
MATERIALS AND METHODS
Parasite strains and culture
Unless otherwise indicated, parasites were maintained at 37 °C in 5% O2/5% CO2/90% N2 in a 2% suspension of human erythrocytes in RPMI medium (Sigma Aldrich) modified with 27 mM NaHCO3, 11 mM glucose, 5 mM HEPES, 0.01 mM thymidine, 1 mM sodium pyruvate, 0.37 mM hypoxanthine, 10 μg/mL gentamycin, and 5 g/L Albumax (Thermo Fisher Scientific).
FSMR strain E2 was generated by selecting a clone of genome reference strain 3D7 (MRA-102, MR4, ATCC, Manassas, VA) under continuous treatment with 3 μM FSM, as previously described (20). Clones of strain E2 were isolated by limiting dilution.
Quantification of FSM resistance
Opaque 96-well plates were seeded with asynchronous cultures at 0.5-1.0% parasitemia (percent of infected red blood cells). After 3 days, media was removed and parasitemia was measured via Picogreen fluorescence on a POLARStar Omega spectrophotometer (BMG Labtech), as previously described (74). Half maximal inhibitory concentration (IC50) values were calculated using Graphpad Prism. Unless otherwise indicated, all IC50 data are representative of means from at least ≥3 independent experiments performed with technical replicates.
HAD2 structural alignment
Structures were aligned using the TM-align algorithm in Lasergene Protean 3D software (root mean square deviation (RMSD) of 1.9 Å).
Whole genome sequencing and variant analysis
Library preparation, sequencing, read alignment, and variant analyses were performed by the Washington University Genome Technology Access Center. One microgram of parasite genomic DNA was sheared, end repaired, and adapter ligated. Libraries were sequenced on an Illumina HiSeq 2500 in Rapid Run mode to generate 101 bp paired end reads. Reads were aligned to the P. falciparum 3D7 reference genome (PlasmoDB v7.2) using Novoalign (V2.08.02). Duplicate reads were removed. SNPs were called using samtools (quality score ≥20, read depth ≥5) and annotated using snpEff. Background variants were removed using previously sequenced genomes from parental and control strains (20). Mixed variant calls and variants in highly variable surface antigen loci (75, 76) were removed.
Sanger sequencing
The HAD2 (PlasmoDB PF3D7_1226300) A469T (R157X) SNP was amplified and sequenced using the HAD2_R157X_F and HAD2_R157X_R primers. The PFK9 locus was amplified using the PFK9_F and PFK9_R primers. PFK9 amplicons were sequenced using the PFK9_seq (1-8) primers. Primer sequences can be found in Table S2.
Generation of recombinant HAD2
The predicted coding sequence of HAD2 was amplified using the HAD2_LIC_F and HAD2_LIC_R primers (Table S2). A catalytic mutant (D26A) was also generated to use as a negative control in activity assays. The had2D26A allele was created using the HAD2_D26A_F and HAD2_D26A_R site-directed mutagenesis primers (Table S2).
Ligation-independent cloning was used to clone HAD2 and had2D26A into vector BG1861 (77), which introduces an N-terminal 6xHis fusion to the expressed protein. BG1861:6xHis-HAD2 was transformed into One Shot BL21(DE3)pLysS Escherichia coli cells (Thermo Fisher Scientific). Protein expression was induced for 3 hours with 1 mM isopropyl-β-D-thiogalactoside at mid-log phase (OD600 0.4-0.5). Cells were collected by centrifugation and stored at −20°C.
Cells were lysed in buffer containing 1 mg/mL lysozyme, 20 mM imidazole, 1 mM dithiothreitol, 1 mM MgCl2 10 mM Tris HCl (pH 7.5), 30 U benzonase (EMD Millipore), and EDTA-free protease inhibitor tablets (Roche). 6xHis-HAD2 was bound to nickel agarose beads (Gold Biotechnology), washed with 20 mM imidazole, 20 mM Tris HCl (pH 7.5), and 150 mM NaCl and eluted in 300 mM imidazole, 20 mM Tris HCl (pH 7.5), and 150 mM NaCl. This eluate was further purified by size-exclusion gel chromatography using a HiLoad 16/600 Superdex 200 pg column (GE Healthcare) equilibrated in 25 mM Tris HCl (pH 7.5), 250 mM NaCl, and 1 mM MgCl2. The elution fractions containing HAD2 were pooled and concentrated and glycerol was added to 10% (w/v). Protein solutions were immediately flash frozen and stored at −80°C.
HAD2 activity assays
The rate of para-nitrophenyl phosphate (pNPP; Sigma-Aldrich S0942) hydrolysis by HAD2 was determined by continuous measurement of absorbance at 405 nm. Assays were performed at 37°C in 50 μl with 50 mM Tris-HCl, pH 7.5, 5 mM MgCl2, 10 mM pNPP, and 1.2 μM enzyme.
Hydrolysis of other phosphorylated substrates by HAD2 was measured using the EnzChek phosphate assay kit (Life Technologies). Reaction buffer was modified to contain 50 mM Tris-HCl, 1 mM MgCl2, pH 7.5, 0.1 mM sodium azide, 0.2 mM 2-amino-6-mercapto-7-methylpurine riboside (MESG), and 1 U/mL purine nucleoside phosphorylase (PNP). Reactions contained 5 mM substrate and 730 nM enzyme. Activity was normalized to that obtained from catalytically inactive 6xHis-HAD2D26A. All data are means from ≥3 independent experiments performed with technical replicates.
P. falciparum growth assays
Asynchronous cultures were seeded at 1% parasitemia. Media (no drug) was exchanged daily. Samples were taken at indicated timepoints and fixed in PBS + 4% paraformaldehyde, 0.05% glutaraldehyde. Cells were stained with 0.01 mg/ml acridine orange and parasitemia was determined on a BD Biosciences LSRII flow cytometer (Thermo Fisher Scientific). All data are means from ≥3 independent experiments.
pTEOE110:HAD2 plasmid construction
The pTEOE110 construct contains the heat shock protein 110 (PF3D7_0708800) 5’ UTR and a C-terminal GFP tag (20). Human dihydrofolate reductase (hDHFR) is present as a selectable marker. Inverted terminal repeats are included for genome integration by a co-transfected piggyBac transposase (pHTH, MRA912, MR4, ATCC, Manassas, VA).
HAD2 was amplified with the HAD2_XhoI_F and HAD2_AvrII_R primers (Table S2) and cloned into AvrII and XhoI sites in the pTEOE110 plasmid.
Parasite transfections
Transfections were performed as previously described (20). Briefly, 50-100 μg of plasmid DNA was precipitated and resuspended in Cytomix (25 mM HEPES pH 7.6, 120 mM KCl, 0.15 mM CaCl2, 2 mM EGTA, 5 mM MgCl2, 10 mM K2HPO4).
A ring-stage P. falciparum culture was washed with Cytomix and resuspended in the DNA/Cytomix solution. Cells were electroporated using a BioRad Gene Pulser II electroporator at 950 μF and 0.31 kV. Electroporated cells were washed with media and returned to normal culture conditions. Parasites expressing the construct were selected by continuous treatment with 5 nM WR92210 (Jacobus Pharmaceuticals). Transfectants were cloned by limiting dilution and presence of the HAD2-GFP construct was verified by PCR using gene- and GFP-specific primers (HAD2_R157X_F and GFP_R, Table S2). Maintenance of the endogenous HAD2 and PFK9 genotypes were verified by Sanger sequencing.
Antiserum generation
Polyclonal anti-HAD2 antiserum was raised in rabbits against 6xHis-HAD2, with Titermax as an adjuvant (Cocalico Biologicals). Antiserum specificity was confirmed by immunoblot of lysate lacking HAD2. Polyclonal anti-HAD1 antiserum has been previously described (MRA-1256, MR4, ATCC) (20).
Immunoblotting
Lysates were separated on a polyacrylamide gel and transferred to a polyvinylidene difluoride membrane. Membranes were blocked in 5% non-fat dry milk, 0.1% Tween-20 in PBS. Rabbit polyclonal antisera were used at the following dilutions: 1:2,000-5,000 anti-HAD2 and 1:20,000 anti-HAD1 (20). For all blots, 1:20,000 HRP-conjugated goat anti-rabbit IgG antibody was used as a secondary antibody (ThermoFisher 65-6120). Blots were stripped with 200 mM glycine, 0.1% SDS, 1% Tween-20, pH 2.2 and reprobed with 1:5,000 rabbit anti-heat shock protein 70 (Hsp70) (AS08 371, Agrisera Antibodies) as a loading control. All blots shown are representative of ≥3 independent experiments. Minimal adjustments were applied equally to all blot images.
PfPFK model construction
PfPFK subunits were searched against the HHpred server for protein remote homology detection and 3D structure prediction using statistics as previously described (78–81). The Borellia burgdorferii PFK structure (PDB 1KZH) (82) returned the highest similarity for both PfPFK domains and was used to predict 3D structure for each domain using the program MODELLER. PFK product orientation in the active site of the model was predicted via the alignment tool, using PyMOL software against the E. coli PFK crystal structure (PDB 1PFK) (83). The α domain model encompasses amino acids 779-1347 and the β domain model encompasses amino acids 110-638.
Assay of native PFK9 activity
Sorbitol-synchronized trophozites were isolated using 0.1% saponin. Cells were washed in buffer containing 100 mM Tris-HCl (pH 7.5), 1 mM MgCl2, 1 mM DTT, 10% glycerol, and EDTA-free protease inhibitor tablets (Roche) and lysed by sonication at 4°C (Fisher Scientific Model 550 Sonic Dismembrator, amplitude 3.5), followed by centrifugation at 4°C (10,000 × g, 10 min). An “RBC carryover” control was comprised of the trace cellular material remaining after saponin lysis, centrifugation, and wash of uninfected erythrocytes.
Lysate PFK9 activity was monitored by linking it to the oxidation of NADH, as previously described (51, 56). Reactions contained 100 mM Tris-HCl (pH 7.5), 1 mM MgCl2, 1 mM DTT, 0.25 mM NADH, 1 mM ATP, 3 mM fructose 6-phosphate, and excess of linking enzymes aldolase (7.5 U), triose-phosphate isomerase (3.8 U), and glycerol 3-phosphate dehydrogenase (3.8 U). After adding fresh cell lysate (10-15 μg total protein), absorbance at 340 nm was measured at 37°C for 40 min. Activity was determined by linear regression using Graphpad Prism software. Unless otherwise indicated, data are means from ≥3 independent experiments.
Metabolite profiling
Approximately ~1 × 109 sorbitol-synchronized early trophozites were isolated using 0.1% saponin, washed with ice-cold PBS + 2 g/L glucose, and frozen at −80 °C. Samples were extracted in 600 μL of ice-cold extraction solvent [chloroform, methanol, and acetonitrile (2:1:1, v/v/v)] using two liquid-nitrogen cooled 3.2 mm stainless steel beads and homogenization in a Tissue-Lyser II instrument (Qiagen) at 20 Hz for 5 minutes in a cold sample rack. Ice-cold water was added and samples were homogenized for 5 min at 20 Hz. Samples were centrifuged at 14,000 rcf at 4°C for 5 min. The polar phase was lyophilized and re-dissolved in 100 μL water and analyzed by LC-MS/MS. LC-MS/MS was performed on a 4000QTRAP system (AB Sciex) in multiple-reaction monitoring mode using negative ioniziation and 10 mM tributylammonium acetate (pH 5.1-5.5) as the ion pair reagent. The specific parameters used for analysis of MEP pathway metabolites have been previously described (19). Liquid chromatography separation was performed using ion pair reverse-phase chromatography (84) with the following modifications: (1) RP-hydro 100 mm × 2.0 mm, 2.5 μm high performance liquid chromatography column (Phenomenex), (2) flow rate of 0.14 mL/min, (3) solvent A of 10 mM tributylammonium acetate in 5% methanol, (4) binary LC gradient (20% solvent B (100% methanol) from 0 to 2.5 min, 30% B for 12.5 min, 80% B for 5 min, and column equilibration at for 5 minutes), and (5) 20 μL autosampler injection volume.
Accession numbers
All genome data have been deposited in the NCBI BioProject database (PRJNA222697) and Sequence Read Archive (SRP038937).
SUPPLEMENTAL MATERIAL LEGENDS
Supplemental Figure 1. FSMR strain E2 does not have increased expression of DXS, DXR, or HAD1.
Supplemental Figure 2. HAD2 is cytosolic.
Supplemental Figure 3. Assay of PFK activity from P. falciparum lysate is linear and specific.
Supplemental Table 1. Relative metabolite levels in strains described in this study.
Supplementary Table 2. Primers used in this study.
ACKNOWLEDGEMENTS
We thank the Genome Technology Access Center in the Department of Genetics at Washington University School of Medicine for help with genomic analysis. The Center is partially supported by NCI Cancer Center Support Grant #P30 CA91842 to the Siteman Cancer Center and by ICTS/CTSA Grant# UL1TR002345 from the National Center for Research Resources (NCRR), a component of the National Institutes of Health (NIH), and NIH Roadmap for Medical Research. This publication is solely the responsibility of the authors and does not necessarily represent the official view of NCRR or NIH. We thank Sophie Alvarez and the Proteomics and Mass Spectrometry Facility at the Donald Danforth Plant Science Center for performing the LC-MS/MS sample prep and analysis. This work was supported by the National Science Foundation under grant number DBI-0521250 for acquisition of the QTRAP LC-MS/MS. We thank Daniel Goldberg (Washington University) for supplying the pTEOE110 plasmid. We thank the Malaria Research Reference and Reagent Resource (MR4) for providing reagents contributed by D.J. Carucci (MRA-102) and John Adams (MRA-912). We thank Katherine Andrews for sharing unpublished data relevant to this study.
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