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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Nature. Author manuscript; available in PMC 2011 February 1.
Published in final edited form as:
PMCID: PMC2917841

This article has been retractedRetraction in: Nature. 2015 March 17; 497(7451): 652    See also: PMC Retraction Policy

Branched Tricarboxylic Acid Metabolism in Plasmodium falciparum


A central hub of carbon metabolism is the tricarboxylic acid (TCA) cycle1, which serves to connect the processes of glycolysis, gluconeogenesis, respiration, amino acid synthesis and other biosynthetic pathways. The protozoan intracellular malaria parasites (Plasmodium spp.), however, have long been suspected of possessing a significantly streamlined carbon metabolic network in which TCA metabolism plays a minor role2. Blood-stage Plasmodium parasites rely almost entirely on glucose fermentation for energy and consume minimal amounts of oxygen3, yet the parasite genome encodes all of the enzymes necessary for a complete TCA cycle4. By tracing 13C-labeled compounds using mass spectrometry5 we show that TCA metabolism in the human malaria parasite P. falciparum is largely disconnected from glycolysis and is organized along a fundamentally different architecture than the canonical textbook pathway. We find that this pathway is not cyclic but rather a branched structure in which the major carbon sources are the amino acids glutamate and glutamine. As a consequence of this branched architecture, several reactions must run in the reverse of the standard direction thereby generating two-carbon units in the form of acetyl-coenzyme A (acetyl-CoA). We further show that glutamine-derived acetyl-CoA is used for histone acetylation while glucose-derived acetyl-CoA is used to acetylate aminosugars. Thus the parasite has evolved two independent acetyl-CoA-production mechanisms with different biological functions. These results significantly clarify our understanding of the Plasmodium metabolic network and highlight the ability of altered variants of central carbon metabolism to arise in response to unique environments.

The mitochondrion of Plasmodium falciparum contains the smallest genome sequenced to date and appears to have evolved reduced functional roles compared to other eukaryotic organisms6. Moreover, the limited number of mitochondrial cristae, minimal oxygen consumption and rapid fermentation of glucose into lactate observed in intraerythrocytic human malaria parasites suggest that oxidative phosphorylation is not a significant source of ATP generation during the blood stage6. Blood-stage Plasmodium spp. have also dispensed with several of the functions often associated with the mitochondrial TCA cycle, such as de novo amino acid biosynthesis. While the parasite possesses a functional electron transport chain and mitochondrial membrane potential is required for survival, we have recently shown that the critical metabolic function of electron transport during blood stage growth is the regeneration of ubiquinone in order to supply pyrimidine biosynthesis7.

Several lines of evidence, however, suggest that TCA metabolism plays an active role in the metabolism of the parasite. The parasite genome encodes orthologues for all TCA cycle enzymes, which are all transcribed during the blood stage8. The citrate synthase orthologue (PF10_0218), aconitase (PF13_0229) and isocitrate dehydrogenase (PfIDH, PF13_0242, see Supplementary Discussion) have been localized to the mitochondrion9,10, and PfIDH, aconitase and succinate dehydrogenase complex (PFL0630w, PF10_0334) have been biochemically characterized1012, suggesting an active mitochondrial pathway. The presence of an essential de novo heme biosynthesis pathway in P. falciparum2 further implies that succinyl-CoA must be generated in the mitochondrion. We recently found that the intracellular levels of several TCA metabolites oscillate over the parasite growth cycle roughly in phase with the expression profiles of cognate enzymes13. Therefore, TCA metabolites are actively synthesized by the parasite. However, it has recently been demonstrated that the P. falciparum pyruvate dehydrogenase (PDH) complex localizes not to the mitochondrion but the apicoplast, a non-photosynthetic plastid-like organelle14. Thus, instead of its canonical role of feeding glucose-derived carbon into the TCA cycle, the suggested role of PDH is solely to produce acetyl-CoA for fatty acid elongation14.

In addition to glucose, major TCA cycle carbon sources in many organisms are the amino acids aspartate, asparagine, glutamate, and glutamine, which can be deaminated to yield oxaloacetate or 2-oxoglutarate (α-ketoglutarate). In order to elucidate the role of the TCA cycle in parasite metabolism we have determined the major carbon source contributing to the accumulation of TCA intermediates. By culturing synchronized parasite-infected red blood cells (RBCs) in medium supplemented with either U-13C-glucose, U-13C-15N-aspartate or U-13C-15N-glutamine we measured intracellular metabolite isotope-labeling patterns throughout the 48-hour parasite cell cycle using a liquid chromatography-mass spectrometry platform capable of detecting most central carbon metabolites.

As expected, in parasites grown on U-13C-glucose the pools of all glycolytic intermediates were rapidly and uniformly labeled (data not shown). We observed limited labeling of carboxylic acid pools, with moderate amounts of +3 13C-malate and +3 13C-fumarate being formed, consistent with phosphoenolpyruvate (PEP) carboxylation incorporating unlabeled carbonate from the gaseous environment15 (Fig. 1a). The absence of labeling into other TCA intermediates suggests that these labeled dicarboxylic acids derive from cytosolic pathways independent of mitochondrial TCA metabolism (Supplementary Fig. 1a). Similarly, growth on U-13C-15N-aspartate results only in the generation of +4 13C-malate and +4 13C-fumarate (Supplementary Fig. 2), which can also occur in the cytosol (Supplementary Fig. 1b).

Figure 1
Glutamine drives reverse flux through the TCA cycle

When feeding U-13C-glucose, PDH complex activity yields acetyl-labeled 13C-acetyl-CoA (Fig 2a). Surprisingly, feeding of labeled glucose results in labeling of only a small fraction of the total acetyl-CoA pool, suggesting the presence of additional sources for two-carbon units. U-13C-glucose feeding also results in small but measurable amounts of both +2 and +5 13C-citrate (Fig 1a), which derive from the condensation of acetyl-labeled 13C-acetyl-CoA with either unlabeled or +3 13C-oxaloacetate, respectively. These labeled forms account for only a minor fraction of citrate and the labeling does not propagate to other intermediates downstream in the TCA cycle. These data raised the possibility that glucose- and aspartate-derived metabolites are disconnected from mitochondrial TCA metabolism.

Figure 2
Acetyl groups deriving from glucose and glutamine are functionally distinct

Consistent with the TCA cycle being fed instead from glutamine, we find significant labeling of all TCA compounds in parasites grown in the presence of U-13C-15N-glutamine (Fig. 1a). Extracellular glutamine is rapidly taken up by parasitized RBCs16 and deamidated to glutamate, which can donate its carbon skeleton to TCA metabolism through conversion to 2-oxoglutarate. Although the growth medium contains only labeled glutamine, the intracellular glutamine/glutamate pools are incompletely labeled due to the generation of unlabeled amino acids by hemoglobin catabolism17. Consistent with this glutamine-driven reaction pathway, 2-oxoglutarate is labeled at all five carbons (Fig. 1a). Similarly, we observe the +4 13C-labeled forms of the four-carbon (C4) compounds succinate, fumarate and malate, expected from the canonical TCA cycle reactions occurring in the standard clockwise direction (Fig. 1b).

Surprisingly, we detect only +5 13C forms of the C6 metabolite citrate. This labeling is inconsistent with the TCA cycle turning in the standard clockwise direction but is characteristic of the reductive carboxylation of 2-oxoglutarate to isocitrate, followed by isomerization to citrate18, in the reverse of standard TCA cycle directionality (Fig. 1c). We also observe +3 13C labeled forms of both malate and fumarate which are generated with temporal profiles similar to those of +5 13C-citrate (Fig 1a). Such malate labeling is consistent with +5 13C-citrate being cleaved into +2 13C- acetate or acetyl-CoA and +3 13C-oxaloacetate, which is then reduced to +3 13C-malate (Fig 1c). We also observe +2 13C acetyl-CoA during growth on U-13C-15N-glutamine (Fig 2a). Thus several TCA cycle reactions are running with net flux in the reverse direction, in the process generating C2 units from 2-oxoglutarate via citrate.

To further dissect the biological role of this reverse TCA branch we investigated the major metabolic fates for C2 units: fatty acid synthesis, protein modification and small molecule acetylation. We profiled carbon-13 labeling of parasite lipids during growth on U-13C-glucose or U-13C-15N-glutamine by gas chromatography-mass spectrometry but were unable to detect labeling under either condition, which is consistent with recent reports that the parasite's de novo fatty acid synthesis pathway is not required during the blood stages19,20. One of the major protein acetylation targets in eukaryotes are the lysine residues within the N-terminal tail of histones. When parasites are cultured in medium containing either U-13C-glucose or U-13C-15N-glutamine we observe robust labeling of the acetyl groups in histone tails only in the U-13C-15N-glutamine-fed cultures (Fig 2b, Supplementary Fig. 3). The acetyl-labeled histones comprise approximately 56% of the total acetylated histone pool, a proportion similar to the fractional labeling of the 2-oxoglutarate pool. However, UDP-N-acetyl-glucosamine (UDP-GlcNAc), a nucleotide aminosugar acetylated in the endoplasmic reticulum during the biosynthesis of glycosylphosphatidylinositol-anchored proteins associated with malaria pathogenesis21, is labeled at the acetyl group only during growth on U-13C-glucose (Fig 2c). Thus it appears that the malaria parasite has evolved two independent pathways that produce acetyl-CoA for different metabolic functions. How glucose- and glutamine-derived C2 units are maintained as functionally distinct pools and transported from their respective organelles to different sites of acetylation remains to be investigated.

Our metabolic labeling data suggest a branched architecture for mitochondrial carbon metabolism in which both arms produce malate. In order to achieve a net flux through these pathways it would be necessary to remove this terminal product, either by conversion or excretion. When we analyzed the liquid culture medium from cultures grown on labeled nutrients, we find that malate, 2-oxoglutarate and, to a lesser extent, fumarate are excreted from infected RBCs at a significant rate (Fig. 3 and Supplementary Fig. 4). Cytosolic fumarate is a byproduct of the parasite's purine salvage pathway22, while 2-oxoglutarate is produced by glutamate dehydrogenase. Our data imply that these metabolites, as well as malate derived from both cytosolic and mitochondrial pathways, are effluxed as waste products.

Figure 3
Malate excretion by P. falciparum-infected RBC cultures

Based on these results we propose a new model for central carbon metabolism in blood stage Plasmodium spp. (Fig. 4). In this pathway the ultimate carbon source for mitochondrial carboxylic acid pools are the amino acids glutamine and glutamate, and carbon flux in the mitochondrion is organized into two independent linear branches. Branch 1 (red in Fig 4) begins with the reductive carboxylation of 2-oxoglutarate to isocitrate, which is then isomerized to citrate. This citrate is cleaved into a C2 compound and oxaloacetate, which is reduced to malate. Branch 2 (blue in Fig. 4) comprises the standard clockwise turning of the TCA cycle to oxidize 2-oxoglutarate to malate, in the process generating reducing power and succinyl-CoA, an essential precursor for heme biosynthesis. The fact that two labeled forms are observed for malate and fumarate but no other TCA intermediates during growth on U-13C-15N-glutamine suggests that both branches converge at these metabolites and they are the terminal products of each. Based on current evidence, our model depicts these pathways as mitochondrial, although the localization of some enzymatic steps and details regarding transport have yet to be fully established (see Supplementary Discussion and Supplementary Figures 5–8).

Figure 4
An integrated model for central carbon metabolism in P. falciparum

This model for branched TCA metabolism is fundamentally different from any yet described. Reductive flux from 2-oxoglutarate has been demonstrated in human brown adipose cell cultures18, in which this pathway was shown to be a source of lipogenic C2 units18. However, these cells appear capable of running both a complete TCA cycle and this reductive pathway simultaneously. This was proposed to be due to the presence of two mitochondrial isoforms of IDH in the human cells: IDH3, the canonical TCA cycle enzyme which uses NAD(H) as a cofactor, and IDH2, which is specific for NADP(H) and may run in the reductive direction due to a mitochondrial NADP+:NADPH ratio favoring the reverse reaction18. Intriguingly, the P. falciparum genome encodes only an NADP(H)-specific, mitochondrial IDH11, suggesting that it may have entirely lost the ability to run a textbook TCA cycle and is effectively locked into this branched architecture. We propose that the mitochondrial NADPH required by this reductive pathway may be generated by the parasite's NADP(H)-specific glutamate dehydrogenase (PF14_0164), and glutamate oxidation has been detected in isolated P. falciparum mitochondria23.

This branched TCA pathway can be understood as an evolutionary trade-off in which metabolic flexibility is lost in order to optimize growth within the specific environment of the host cell. Within the human bloodstream, an abundant and homeostatic supply of glucose ensures a constant supply of energy, while the high levels of plasma glutamine (~0.5 mM) represent a ready source of C5 carbon skeletons to drive the mitochondrial production of reduced ubiquinone, succinyl-CoA and C2 acetyl units. In human cells it has been demonstrated that production of nuclear acetyl-CoA from mitochondrially-derived citrate is a major determinant of the acetylation state of histones24, and acetylation of metabolic enzymes is gaining recognition as a major post-translational modification involved in sensing and regulating responses to nutrient availability in diverse organisms25,26. It is possible that flux through this reductive TCA pathway in P. falciparum serves as a nutrient sensor regulating enzymatic activities and transcriptional responses via protein acetylation. Also, recent studies have found TCA cycle enzymes up-regulated in a subset of patient-derived blood stage parasite isolates27 as well as in salivary gland sporozoites27,28. Our results suggest that under these glucose-limited conditions, reductive TCA flux might compensate for reduced synthesis of C2 units from glucose. Whether the pathway architecture described in our model is maintained within other tissues invaded during the parasite life cycle, such as the mosquito midgut and salivary gland or the human liver, merits further study as the nutrient availabilities and metabolic demands in these environments vary substantially.

Our results highlight the growing role metabolomic technologies play in elucidating the architecture of metabolic pathways, particularly in such divergent pathogens as Apicomplexan parasites. Genomic reconstructions4, which generally map metabolic networks onto those of well-studied model organisms, must be informed by direct experimental evidence or run the risk of failing to identify the pathways that represent the best candidates for drug targets. This study clarifies our understanding of the metabolism underlying mitochondrial electron flow, heme biosynthesis and histone acetylation, all of which are current or suggested targets for pharmaceutical intervention29,30. In addition, it presents a clear case in which a fundamental metabolic pathway has undergone significant evolutionary adaptation towards a particular environmental niche.

Methods Summary

P. falciparum culturing and metabolomics were performed essentially as described in Olszewski et al.13. For further details, and for descriptions of cloning, fluorescent imaging, mitochondrial isolation, enzyme assays, histone extraction, proteomics and GC-MS analysis, see the Supplementary Methods.


P. falciparum culturing and metabolite extraction

P. falciparum cultures were maintained and synchronized by standard methods31,32. Briefly, P. falciparum (3D7 strain) infected red blood cells (RBCs) were grown in RPMI 1640 culture medium supplemented with sodium carbonate (2 mg/mL), hypoxanthine (100 μM), Albumax II (0.25%) and gentamycin (50 μg/mL) in a humidified incubator at 5% CO2, 6% O2 and 37° C. Human RBCs used for culturing were harvested two days prior to use in tubes supplemented with sodium heparin instead of standard citrate-containing anticoagulants in order to avoid contaminating citrate.

For metabolic labeling experiments, growth medium was formulated according to the standard nutrient concentrations of RPMI 1640 medium33. Vitamins were supplied using RPMI 100× Vitamins Solution (Sigma-Aldrich Corporation, St. Louis, MO); inorganic salts and all other nutrients were purchased individually from Sigma-Aldrich and were the highest purity available. This reformulated media was found by standard growth assays to be indistinguishable from commercial RPMI 1640 mixes in supporting P. falciparum growth. U-13C-glucose, U-13C-15N-aspartate and U-13C-15N-glutamine were purchased from Cambridge Isotope Laboratories (Andover, MA) and used to replace the replace the unlabeled nutrient at the normal concentrations of each (glucose, 2 g/L; aspartate, 20 mg/L; glutamine, 300 mg/L).

Metabolic labeling experiments were carried out as follows: a highly synchronized P. falciparum culture in the late schizont stage was inspected hourly by microscope until host cell lysis and reinvasion was complete. This culture was then adjusted to 6% parasitemia using cultured RBCs and diluted to 0.4% hematocrit in fresh, prewarmed (37° C) media containing one of the labeled nutrients and returned to the incubator. The cultures were allowed to equilibrate for 2 hours and then infected RBCs and culture medium were harvested for the t=0 timepoint and at 8-hour intervals thereafter. An uninfected RBC culture was treated similarly and extracted at t=0 to use for normalization.

Metabolite extraction was carried out using a modified version of our previous protocol13. Briefly, RBCs were pelleted from liquid cultures by centrifugation for 5 minutes at 500×g. Media samples were collected from the supernatant by dilution in 4 volumes of 100% methanol at −70° C, then the supernatant was removed by aspiration. The cell pellet was quickly flash-quenched in 4 volumes of 100% methanol at −70° C and incubated on dry ice for 15 minutes with vortexing every 5 minutes. The lysate was centrifuged 5 minutes at 500×g and the supernatant collected. The pellet was re-extracted in 10 volumes of 80:10 methanol:water at 4° C and sonicated on ice in a water-bath sonicator for 15 minutes. This was then centrifuged 5 minutes at 16,000×g and the supernatant collected and pooled with the previous extract. The pooled extract was centrifuged 10 minutes at 16,000×g to precipitate denatured protein, and then the supernatant transferred to a fresh tube and dried under nitrogen flow. Dried extracts were stored at −70° C until LC-MS analysis (less than 96 hours). For analysis, the dried extracts were resuspended in 200 μL of chromatographic buffer A (97:3 water:methanol, 10 mM tributylamine, 15 mM acetic acid). Media extracts were also prepared as described above. To quantitate TCA intermediates in media and cell extracts, a parasite culture was maintained in label-free media and extracted at 24 hours post invasion into methanol containing isotope-labeled internal standards at the following concentrations: U-13C-malic acid, 5 μg/mL; U-13C-fumaric acid, 0.5 μg/mL; 1,4-13C-succinic acid, 0.5 μg/mL; 2,4-13C-citric acid, 1 μg/mL; 1,2,3,4-13C-ketoglutaric acid disodium, 5 μg/mL (Cambridge Isotope Laboratories, Andover, MA). The extraction was otherwise performed as above.

LC-MS instrumentation

LC-MS analyses were performed on Exactive Orbitrap mass spectrometer, coupled with Accela U-HPLC system (Thermo Fisher Scientific, San Jose, CA) and HTC PAL autosampler (CTC Analytics AG, Zwingen, Switzerland). Liquid chromatography separation was achieved on a Synergy Hydro-RP column (100 × 2 mm, 2.5 μ particle size, Phenomenex), and the gradient is a modified version from our previous method34: 0 min, 0% B; 2.5 min, 0% B; 5 min, 20% B;7.5 min, 20% B; 13 min, 55% B; 15.5 min, 95% B; 18.5 min, 95% B; 19 min, 0% B; 25 min, 0% B. Solvent A is 97:3 water:methanol with 10 mM tributylamine and 15 mM acetic acid; solvent B is methanol. Other LC parameters are: autosampler temperature 4 °C; injection volume 10 μl; column temperature 25° C.

The Exactive Orbitrap mass spectrometer was operated in negative mode, scanning m/z 85–1000. Other instrumental parameters are: resolution 100,000 at 1 Hz (1 scan per second); AGC (automatic gain control) target 3E6; maximum injection time 100 μS; sheath gas flow rate 25 (arbitrary unit); aux gas flow rate 8 (arbitrary unit); sweep gas flow rate 3 (arbitrary unit); spray voltage 3 kV; capillary temperature 270 °C; capillary voltage −50 V; tube lens voltage −100 V.

LC-MS data extraction and analysis

Thermo Fisher mass spectrometry RAW files were converted from profile mode into centroid mode using the ReAdW program35. Centroided files were loaded into mzROLL, an in-house analysis program, and aligned with a non-linear regression using a high degree polynomial to the retention times of the highest intensity mass-charge ratio (m/z) measurements from each sample to construct a median reference. Extracted ion chromatograms (EICs) were extracted using a 5 ppm window centered around the expected m/z of each compound and smoothed by applying a Gaussian filter to the intensity signal. Within each EIC, peaks were detected and the quality of all peaks was evaluated by a Random Forest based classification model that takes into account peak height, peak width, peak area, the signal to noise ratio, and the peak shape36. Peaks were grouped across samples and matched to the expected retention time of each compound. The peak closest to the expected retention with a quality score >0.5 was used for quantitation. All peaks used for quantitation were hand-checked after automated extraction and assignment. The isotopically labeled forms of compounds were extracted in a similar manner. Quantitation of labeled forms was based on the highest intensity peak within a 5 ppm window around the expected m/z of the carbon-13 and/or nitrogen-15 labeled form of the compound. The specifics of the mzROLL program will be described in a forthcoming publication.

Metabolites were identified based both on their match to the expected m/z ratio and chromatographic retention times determined previously for standard solutions. Isotope-labeled forms were identified using the expected mass shifts given by carbon-13 and nitrogen-15. Where ambiguous, the positional labeling of specific atoms within the molecule was determined by LC-MS/MS analysis as described previously13. The height of the extracted peak for each compound was used as the signal. The raw signals for each labeled form are corrected to account for the naturally occurring isotope distribution as calculated by the Qual Browser included in the Xcalibur software suite (Thermo Fisher Scientific, San Jose, CA). The signal due to naturally occurring isotopes was added to the signal of the unlabeled form. Similarly, signals that were due to incomplete labeling of the isotope-labeled nutrients (which are generally 98–99% fully-labeled) were discounted. For quantitation experiments, the signal ratio of the unlabeled metabolite to its isotope-labeled internal standard was determined and used to calculate the concentration of the metabolite in the extract at the 24-hour timepoint. The relative signal in the other timepoint samples was used to calculate the concentrations over the temporal profile.

For display plots, the data for all labeled and unlabeled forms of a given metabolite are expressed as the ratio of the signal for that form at that timepoint to the signal of the unlabeled form in the t=0 timepoint of the 13C-glucose sample. Signals from the uninfected RBC sample are treated as the background level in host cells and were subtracted from every timepoint; where this reduced signals to less than 1000 counts the signal was set to 1000 counts, the approximate limit of quantitation for the instrument. Each plotted point shows the average of n= 3 biological replicates; error bars show the standard deviation. Labeled forms are only plotted if the average signal in at least one timepoint is greater than 1000 counts and the signal represents at least 1% of the signal of the unlabeled form.

Fatty acid extraction and analysis

For lipid-labeling experiments, parasites were cultured as above in labeled nutrient-supplemented RPMI for 96 hours (encompassing two growth cycles) harvested at the trophozoite stage. These experiments were conducted using both normally formulated RPMI and a minimal fatty acid formulation (lacking Albumax II, supplemented with 60 μM lipid-free BSA and 30 μM each of myristic, stearic and oleic acids) prepared according to an earlier report that growth in this media resulted in enhanced elongation of preformed fatty acids by the parasite37. Infected RBCs were lysed by treatment with 100 pellet volumes of 0.1% saponin in phosphate-buffered saline (PBS) buffer and the freed parasite cells collected by centrifugation at 2500×g, 4° C for 10 minutes. After washing the parasite pellet in PBS, lipids were extracted and derivatized to fatty acid methyl esters using the protocol of Yoo et al.18.

The fatty acid methyl esters were investigated by GC-MS, using an Agilent 7890A GC coupled to a 5975C inert MSD with a Rtx-5Sil MS (30m length, 0.25mmID, 0.25μm film) column. The thermal gradient used was: start at 70° C, holding for 2 min; ramping to 230° C at 20° C/min. The mass spectra of the molecular ions of the palmitic and stearic acid methyl esters (monoisotopic m/z = 270.45 and m/z = 298.51) were examined for deviation from the naturally occurring isotope distribution indicating 13C incorporation. No incorporation was detected in either the normal or minimal fatty acid RPMI cultures.

Histone extraction and analysis

For histone labeling experiments, parasites were cultured as above in labeled nutrient-supplemented RPMI for 96 hours (encompassing two growth cycles) and harvesting at the trophozoite stage. Infected RBC cultures (50 mL total volume, 2% hematocrit, 10% parasitemia) were lysed by treatment with 100 pellet volumes of 0.1% saponin in phosphate-buffered saline (PBS) buffer and the freed parasite cells collected by centrifugation at 2500×g, 4° C for 10 minutes. After washing the parasite pellet in PBS, histones were acid-extracted38 and purified39 according to modified versions of the published protocols. Briefly, the parasite pellet was resuspended in 800 μL of acid extraction buffer (0.2 M HCl, 1 mM dithiothreitol, 1 mM sodium orthovanadate, 10 mM sodium butyrate, 1 Roche EDTA-free protease inhibitor tablet per 50 mL). The suspension was gently agitated at 4° C for 2 hours, and then centrifuged at 16,000×g for 1 minute at 4° C. The supernatant was collected and 246 μL of 100% trichloroacetic acid was slowly added. This was mixed by inversion and incubated on ice overnight, and then centrifuged at 16,000×g for 10 minutes at 4° C. The supernatant was discarded and the pellet washed twice with cold acetone at 4° C. The acetone was allowed to evaporate and the pellet resuspended in deionized water for analysis.

Histone extracts were fractionated by RP-HPLC on a C18 column using a 30–60% B in 100 minute gradient (buffer A = 5% acetonitrile in 0.2% TFA, buffer B = 90% acetonitrile in 0.188 TFA). Fractions containing histone H4 were pooled and propionylated as previously published40. Propionylated histone H4 was digested with trypsin at a 20:1 protein:enzyme ratio for 7 hours at 37° C. Digested histone H4 was subjected to MS analysis on an Orbitrap mass spectrometer operated by obtaining a full mass spectrum at 30,000 resolution in the Orbitrap followed by 7 data-dependent MS/MS spectra acquired in the ion trap. All mass spectra were manually interpreted. Quantitation of carbon-13 labeling was performed using the natural isotope distribution of the propionylated acetyl-H4 peptide calculated by the Qual Browser included in the Xcalibur software suite (Thermo Fisher Scientific, San Jose, CA).

Preparation of P. falciparum mitochondria

P. falciparum cultures were synchronized twice by treatment with sorbitol as described, expanded and harvested at 8% parasitemia in the early trophozoite stage. Fractions substantially enriched in mitochondria were prepared using a procedure modified from the method of Takashima et al41. Parasitized erythrocytes were harvested by centrifugation, washed in AIM (120 mM KCl, 20 mM NaCl, 20 mM glucose; 6 mM HEPES, 6 mM MOPS, 1 mM MgCl2, 0.1 mM EGTA, pH 7.0) and lysed with 0.05% (w/v) saponin in AIM. After washing 3 times with AIM and once with MSEH (225 mM mannitol, 75 mM sucrose, 4.3 mM MgCl2, 0.25 mM EGTA, 10 mM HEPES [Tris], 5 mM HEPES [KOH]; pH 7.4), the parasites were disrupted by N2 cavitation (using a 4639 Cell Disruption Bomb, Parr, USA) at 1000 psi for 20 min at 4°C in deaerated MSEH buffer containing 5 mM glucose and mitochondrial substrates (5 mM α-glycerophosphate and 2.5 mM dihydroorotate) in the presence of 1 mM PMSF and 1 μL of fungal protease inhibitor cocktail (Sigma-Aldrich, Inc., St. Louis, MO, USA) per ml. After drop-wise release from the N2 bomb, another aliquot of protease inhibitors was mixed into the disrupted parasite sample. The unbroken cells and cell debris were removed by centrifugation at 900×g for 6 min at 4°C. The low speed supernatant was passed slowly through a MACS CS column prewashed with MSEH in a Vario MACS magnetic separation apparatus (Miltenyi Biotec, Auburn, CA, USA) to remove most of the hemozoin from the preparation. The mitochondria were then recovered as a pellet by centrifugation at 23000×g for 20 min at 4°C. The pellet was suspended in a minimal volume of MSEH containing 1 mM dihydroorotate and 1 mg/ml fatty acid-free BSA and stored at −80°C.

Mitochondrial IDH assay

Assays for mitochondrial reductive isocitrate dehydrogenase activity were performed using a modified version of the protocol of Kornberg and Pricer42. Briefly, purified mitochondrial preparations were thawed on ice and diluted into 9 volumes of assay buffer (final concentration: 50 mM Na2HPO4, 0.5 mM MgCl2, 5 mM NaHCO3, 0.2 mM NADPH, 10 mM 1,2,3,4-13C-2-oxoglutarate, pH 7.0) in a sealed tube. The reaction mixture was incubated in a water bath at 37 °C and samples removed at the specified times. The reaction was quenched by dilution into 9 volumes of methanol at −70 °C. These samples were centrifuged for 15 minutes at 16,000×g, 4 °C, to precipitate protein and biological material, then the supernatant was diluted into 9 volumes of water and subjected to HPLC-MS analysis as described.

Citrate lyase assays

Assays for ATP:citrate lyase and ATP-independent citrate lyase were performed on lysates of uninfected erythrocytes, infected erythrocytes (trophozoite-stage, 10% parasitemia), host cell-free parasites (trophozoite-stage) and isolated mitochondria. Host cell-free parasites were prepared by standard saponin treatment: briefly, blood cultures were harvested by centrifugation (500 RCF, 5 minutes); washed once with PBS (500 RCF, 5 minutes); resuspended at 2% hematocrit in 0.1% w/v saponin in PBS and incubated 2 minutes at room temperature; pelleted by centrifugation (2000 RCF, 10 minutes, 4° C); and washed once more in PBS (2000 RCF, 10 minutes, 4° C).

ATP:citrate lyase assays were performed using a protocol modified slightly from Ma et al.43. Erythrocyte and parasite cell samples were diluted into 9 volumes of lysis buffer [50 mM Tris-HCl (pH 8.0), 50 mM NaCl, 2 mM DTT, 1 mM MgCl2, 1 EDTA-free protease inhibitor cocktail tablet per 10 mL (Roche complete Mini)], lysed by three rounds of freeze-thaw cycles (1 minute in liquid nitrogen, 5 minutes in 37° C water bath) and centrifuged (16,000 RCF, 10 minutes, 4° C) to clear the supernatant. Mitochondrial preparations were diluted into the same buffer but with 0.05% dodecyl maltoside to permeabilize the membranes and were not lysed. 2.5 μL of these samples were diluted into 17.5 μL of reaction cocktail with final concentrations of: 87 mM Tris-HCl (pH 8.0), 20 μM MgCl2, 10 mM KCl, 10 mM DTT, 100 μM coenzyme A, 150 μM 2,4-13C-citric acid, with or without 400 μM ATP. The reaction mix was incubated for 15 minutes at 37° C and terminated by adding 80 μL of methanol and cooling to −70° C on dry ice for 15 minutes. The samples were centrifuged (16,000 RCF, 10 minutes, 4° C) and the supernatant diluted 1:10 in water and analyzed by LC-MS for the production of +1 13C-acetyl-CoA.

ATP-independent citrate lyase assays were performed using a protocol modified from Bergmeyer et al44. The samples were prepared as described above except in a buffer with the formulation: 10 mM triethanolamine (pH 7.6), 0.3 mM ZnCl2, 454 mM ammonium sulfate, 10 mM DTT and 1 EDTA-free protease inhibitor cocktail tablet per 10 mL (Roche complete Mini). 1 μL of sample was diluted into 29 μL of a reaction cocktail to final concentrations: 96 mM triethanolamine (pH 7.6), 0.5 mM ZnCl2, 0.23 mM β-NADH, 0.67 mM 2,4-13C-citric acid, 15 mM ammonium sulfate, 100 unites L-lactate dehydrogenase, 50 units malate dehydrogenase. The reaction mix was incubated for 2 hours at 37° C and terminated by adding 120 μL of methanol and cooling to −70° C on dry ice for 15 minutes. The samples were centrifuged (16,000 RCF, 10 minutes, 4° C) and the supernatant diluted 1:10 in water and analyzed by LC-MS for the enzyme-linked production of +1 13C-malate and +1 13C-lactate.

IDH leader-GFP localization

The GFP gene was amplified from pHDGFP45 with added 5' Xho I and 3' Sal I sites. The internal BstBI site was eliminated using site directed mutagenesis. The modified GFP was sub-cloned into pHHMC*/3R0.546 digested with Xho I, producing the plasmid pHHGFP19. The 5' 204 bp of the Pf IDH gene (PF13_0242), corresponding to the initial 68 amino acids (MGKHIRILKNQYLQFMSKRCIQSKAAFNICGKINVENPIVELDGDEMTRIIWKDIKEKLILPYVNLKI), were amplified from P. falciparum 3D7 DNA with primers adding a 5' BstB I site and a 3' Xho I site. The product was inserted into pHHGFP19 digested with BstB I and Xho I to produce pHHIDHldrGFP. The cloned DNA sequences were all confirmed by sequencing. P. falciparum transfection was performed using standard methods47 and parasites were selected with the drug WR99210.

Primers used:



PCR prod 806 bp





Fluorescence microscopy

Live infected erythrocytes were suspended in RPMI medium containing mitotracker Red CM-H2XROS (Molecular Probes) at 50 nM and Hoechst 33342 dye (Sigma) at 1 μg/ml and incubated at 37°C for ~25 minutes. The stained erythrocytes were immobilized on a microscope coverslip in RPMI using a fibrin clot procedure modified from Forer and Pickett-Heaps48. Images were captured with an Olympus BX60 microsope equipped with a SPOT RT Slider digital camera and software system (Diagnostic Instruments).

Supplementary Material


We are grateful to G. McFadden and I. Sherman for critical discussions and scrutiny of the manuscript; B. Bennett, T. Campbell, E. De Silva, J. O'Hara, and H. Painter for critical reading of the manuscript; I. Ying for assistance with histone extraction; T. Spurck and C. Tonkin for the modified erythrocyte immobilization procedure for microscopy; M. Clasquin and W. Lu for developing the LC-MS methodology; E. Melamud for LC-MS data extraction and analysis; J. Groves and H. Cooper for GC-MS analysis. M.L. is funded by the Burroughs Wellcome Fund and an NIH Director's New Innovators award (1DP2OD001315-01). J.D.R. is funded by a Beckman Young Investigators award, an NSF CAREER award, and NIH R01 AI078063. M.L and J.D.R. receive support from the Center for Quantitative Biology (P50 GM071508). B.A.G. receives support from NSF grant CBET-0941143. K.O. is funded by an NSF Graduate Research Fellowship. J.M.M., M.W.M, and A.B.V. were supported by a grant AI028398 from NIAID, NIH.


Supplementary Information is linked to the online version of the paper at


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