ApicoplastandEndoplasmicReticulumCooperateinFatty ... · fatty acid biosynthetic pathways that...

Apicoplast and Endoplasmic Reticulum Cooperate in FattyAcid Biosynthesis in Apicomplexan Parasite Toxoplasmagondii*□S

Received for publication, October 4, 2011, and in revised form, December 15, 2011 Published, JBC Papers in Press, December 16, 2011, DOI 10.1074/jbc.M111.310144

Srinivasan Ramakrishnan‡1, Melissa D. Docampo‡, James I. MacRae§, François M. Pujol¶2, Carrie F. Brooks�,Giel G. van Dooren�3, J. Kalervo Hiltunen¶2, Alexander J. Kastaniotis¶2, Malcolm J. McConville§4,5,and Boris Striepen‡�5,6

From the ‡Department of Cellular Biology and �Center for Tropical and Emerging Global Diseases, University of Georgia, Athens,Georgia 30602, the §Department of Biochemistry and Molecular Biology, Bio21 Institute of Molecular Science and Biotechnology,University of Melbourne, Parkville, Victoria 3010, Australia, and the ¶Department of Biochemistry and Biocenter Oulu, University ofOulu, FIN-90014 Oulu, Finland

Background: Parasite fatty acid synthesis is an attractive drug target but complex and poorly understood.Results:We delineate the molecular activity of two pathways in Toxoplasma combining metabolomic and genetic analyses.Conclusion: The apicoplast is a significant source of fatty acids, and its products are further modified in the parasite endoplas-mic reticulum.Significance:We define the metabolic host-parasite relationship with molecular resolution in intracellular parasites.

Apicomplexan parasites are responsible for high impacthuman diseases such as malaria, toxoplasmosis, and cryptospo-ridiosis. These obligate intracellular pathogens are dependenton both de novo lipid biosynthesis as well as the uptake of hostlipids for biogenesis of parasite membranes. Genome annota-tions and biochemical studies indicate that apicomplexan para-sites can synthesize fatty acids via a number of different biosyn-thetic pathways that are differentially compartmentalized.However, the relative contribution of each of these biosyntheticpathways to total fatty acid composition of intracellular parasitestages remains poorly defined. Here, we use a combination ofgenetic, biochemical, andmetabolomic approaches to delineatethe contribution of fatty acid biosynthetic pathways in Toxo-plasma gondii. Metabolic labeling studies with [13C]glucoseshowed that intracellular tachyzoites synthesized a rangeof longand very long chain fatty acids (C14:0–26:1). Genetic disruptionof the apicoplast-localized type II fatty-acid synthase resulted ingreatly reduced synthesis of saturated fatty acids up to 18 car-bons long. Ablation of type II fatty-acid synthase activityresulted in reduced intracellular growth that was partially

restored by addition of long chain fatty acids. In contrast, syn-thesis of very long chain fatty acids was primarily dependent ona fatty acid elongation system comprising three elongases, tworeductases, and a dehydratase that were localized to the endo-plasmic reticulum. The function of these enzymes was con-firmed by heterologous expression in yeast. This elongase path-way appears to have a unique role in generating very longunsaturated fatty acids (C26:1) that cannot be salvaged from thehost.

Apicomplexans are a phylum of single-celled eukaryotic par-asites that includes the causative agents for important humandiseases such as malaria, toxoplasmosis, and cryptosporidiosisas well as numerous veterinary pathogens. Control of these dis-eases is challenging, in no small part due to a rapid emergenceof drug resistance that requires a continuous pipeline of newdrugs with novel targets and/or modes of action. Apicomplex-ans are obligate intracellular pathogens and deploy complexmechanisms of synthesis and salvage to meet their essentialnutritional requirements. The acquisition of lipids and fattyacids has emerged as a particularly important aspect of themetabolism of intracellular pathogens (1–3). Fatty acids are themain building blocks of membranes and also play a significantrole in the post-translational modification of numerousproteins.Apicomplexanswere believed to be incapable of synthesizing

their own fatty acids and to rely completely on host-derivedlipids. However, the discovery of the apicoplast, a plastid-likeorganelle challenged this view. As found for the chloroplasts ofplants and algae, the apicoplast contains a type II fatty-acidsynthesis (FASII)7 pathway (4). Humans lack plastids and the

* This work was supported, in whole or in part, by National Institutes of HealthGrants AI084414 and AI064671 (to B. S.). This work was also supported byNational Health and Medical Research Council Grant 1006024 (to M. J. M.).

□S This article contains supplemental Tables S1–S3.The nucleotide sequence(s) reported in this paper has been submitted to the Gen-

BankTM/EBI Data Bank with accession number(s) JN544918 and JN5449191 Supported by a predoctoral fellowship from the American Heart

Association.2 Recipient of funding from the Academy of Finland and the Sigrid Juselius

Foundation.3 Present address: Research School of Biology, Australian National University,

Canberra, ACT, Australia.4 National Health and Medical Research Council Principal Research Fellow.5 Both authors contributed equally to this work.6 Georgia Research Alliance Distinguished Investigator. To whom corre-

spondence should be addressed: Center for Tropical and Emerging GlobalDiseases, University of Georgia, 500 D.W. Brooks Dr., Athens, GA 30602. Tel.:706-583-0588; Fax: 706-542-3582; E-mail: [email protected].

7 The abbreviations used are: FASII, type II fatty-acid synthase; ER, endoplas-mic reticulum; ACP, acyl carrier protein; 5�FOA, 5�-fluoroorotic acid; ATc,anhydrotetracycline.

THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 287, NO. 7, pp. 4957–4971, February 10, 2012© 2012 by The American Society for Biochemistry and Molecular Biology, Inc. Published in the U.S.A.

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associated FASII, and instead they rely on type I fatty acid syn-thesis for the production of bulk long chain fatty acids. A FASII-type synthase is present in human mitochondria, but there arestructural differences between its components and those ofbacterial or apicomplexan FASII (5). Furthermore, apicoplastFASII also differs from the mammalian FASI pathway, therebymaking it a potential target for the treatment of apicomplexaninfections (6). Initial pharmacological studies on Plasmodiumfalciparum and Toxoplasma gondii supported this idea (7, 8).Subsequent genetic studies found the essentiality of this path-way to be more complicated. We constructed a conditionalmutant for the apicoplast acyl carrier protein (ACP) in T. gon-dii. ACP is a key molecule of the FASII pathway; fatty-acylintermediates are covalently bound to ACP as the enzymes ofthe pathway act on them. Loss of ACP in this conditionalmutant affects apicoplast biogenesis and results in the death ofthe parasite. Moreover, mice infected with a lethal dose of T.gondii can be cured by suppression of the FASII pathway, sug-gesting its importance in the growth and survival of the parasite(9). In contrast, this pathway is not essential for the blood-stream and mosquito stages of the malarial parasite Plasmo-dium, although it is essential for the liver stage that is requiredfor establishment of infection in the mammalian host (10, 11).Other apicomplexan parasites, such asCryptosporidium, Babe-sia, and Theileria appear to completely lack the FASII biosyn-thetic machinery suggesting that they are dependent on fattyacid salvage from the host (1).Although both T. gondii and P. falciparum have been shown

to take up radiolabeled fatty acids from the culturemedium (12,13), these parasites are predicted to contain a number of otherfatty acid biosynthetic pathways that could partly compensatefor the loss of FASII (1, 14). These alternative mechanismsinclude a FASI pathway (in T. gondii, Eimeria, and Cryptospo-ridium) and a putative fatty acid elongation pathway (in mostapicomplexans with the exception of Theileria and Babesia).FASI in apicomplexans so far has only been studied in Crypto-sporidium parvum (15, 16).Overall, these studies suggest that apicomplexan fatty acid

metabolism is complex and thatmultiple potentially redundantbiosynthetic pathways could contribute to the total cellularfatty acid pool. Identifying the specific biochemical activities ofindividual components and delineating their sequence andinteraction and their relative importance under differentgrowth conditions are critical for understanding host-parasiteinteraction and identifying potential drug targets. Here, wedevelop a combined genetic and metabolomic strategy to iden-tify the products of FASII and the elongation pathways for T.gondii. De novo fatty acid synthesis was monitored in wild typeparasites and in mutant parasite lines in which specific fattyacid biosynthetic enzymes were selectively ablated by labelingparasite-infected host cells with [U-13C]glucose and analysis ofparasite lipids by gas chromatography-mass spectrometry (GC-MS). The activity of specific fatty acid elongases was furtherexamined by heterologous expression and biochemical analy-ses. Our analyses suggest that the FASII and elongase pathwayshave nonredundant functions in intracellular tachyzoite stagesinmaintaining the fatty acid composition of bulk lipids and that

these de novo pathways cannot be by-passed by fatty acid sal-vage from the host cell.

EXPERIMENTAL PROCEDURES

Gene Identification and Gene Tagging—The proteinsequences of the yeast fatty acid elongation machinery(ELO1–3, ketoacyl-CoA reductase, dehydratase, and enoyl-CoA reductase; GenBankTM accession numbers NP_012339,NP_009963, NP_13476, AAS56194, NP_012438, NP_010269)were used as query sequences for BLAST searches against theT.gondii genome and GenBankTM databases. To evaluate thecomputational predictions for these genes, we conducted RT-PCRexperiments. The predictions forT. gondiiELO-A, ELO-B,ELO-C, andDEHwere confirmed. However, for ECR and KCR,the beginning and ending of the coding sequences were differ-ent and were subsequently established by RT-PCR, subcloning,and sequencing. The experimentally validated T. gondii genesdescribed in this study are ELO-A, ELO-B, ELO-C, KCR, DEH,and ECR (see supplemental Table S1 for GenBankTM accessionnumbers). The full-length coding sequences for all genes wereamplified from T. gondii cDNA using primers introducingflanking BglII and AvrII restriction sites, subcloned into plas-mid pCR2.1 (Invitrogen), and subsequently introduced into theequivalent sites of either plasmid pdt7s4H3 or pdt7s4M3 (17),placing them under the control of a tetracycline-regulatablepromoter and fusing a triple HA (pDT7S4H3) or a triple c-Mycepitope tag (pDT7S4M3), respectively, to the 3�-end. All prim-ers used are listed in supplemental Table S1.Parasite Culture and Construction of Mutants—Parasites

were cultured and genetically manipulated as described previ-ously (18). Conditional mutants were generated using the pre-viously described two-step strategy (9, 19). Plasmids containinga second copy of the target gene under the control of a regulat-able promoter (pdt7s4H3) were stably introduced into a T. gon-dii tetracycline transactivator strain (20). Stable clones forELO-A, -B, or -C were established in the presence of pyrimeth-amine.Wenext engineered a plasmid-based construct to deletethe ELO-C locus and cosmid-based targeting vectors forELO-A and -B. For ELO-C, sequences (�2 kb) upstream anddownstream of the ELO-C genomic locus were amplified byPCR and subcloned into the respective BamHI/KpnI andAatII/SpeI restriction sites of the PTCY vector flanking a chloram-phenicol resistance cassette (the vector also contains a YFP cas-sette for negative selection (9)). This construct was linearizedwith SpeI and transfected into the pDT7S4-ELO-C-H3 parasiteline. Transgenic parasites that were chloramphenicol-resistantand YFP-negative were isolated. Clones were tested by PCR forintegration, and gene disruption was confirmed by Southernblot analyses. (Plasmids used for the probes are listed in supple-mental Table S2.) Cosmids (TOXP929/ELO-B or TOXO789/ELO-A) were modified to replace the coding sequence with achloramphenicol resistance cassette by recombineering (19).Targeting cosmids were transfected into the respective para-site, and transgenic clones were selected in the presence ofchloramphenicol. Mutants were identified as described forELO-C.We conducted plaque assays in the absence or presenceof 0.5 �M ATc to evaluate the growth of mutants (18). Forchemical complementation assays, myristic and palmitic acids

Toxoplasma Fatty Acid Metabolism

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were dissolved in equal molar amounts in ethanol, dried into amicrocentrifuge tube, and complexed to fatty acid-free BSA inwater (Invitrogen, 1:3 molar ratio) with sonication prior toaddition to normal growth medium as indicated.Microscopy—Immunofluorescence microscopy was per-

formed as described previously (17). Infected coverslip cultureswere fixed with 3% paraformaldehyde in PBS, followed by per-meabilization in 0.25% Triton X-100 in PBS. After blocking, ratanti-HA antibodies (Roche Applied Science) were used at adilution of 1:100, rabbit anti-ACP (a kind gift from GeoffMcFadden, University ofMelbourne (4)) at 1:1000,mouse anti-GFP (Torrey Pines Biolabs) at 1:400, and rabbit anti-Myc(Roche Applied Science) at 1:100. Secondary antibodies wereused at a dilution of 1:200 and were anti-rat Alexa Fluor 546,anti-rabbit Alexa Fluor 546, anti-mouse Alexa Fluor 488, andanti-rat Alexa Fluor 488 (all from Invitrogen). Images weretaken using an Applied Precision Delta Visionmicroscope, andimageswere deconvoluted and adjusted for contrast using Soft-worx. Protein localization annotations have been submitted tothe ApiLoc data base of protein subcellular localization inApicomplexa.Western Blotting—Western blot analyses were performed as

described previously (17) and used rat anti-HA antibodies(Roche Applied Science) at 1:100 and anti-GRA8 antibodies at1:2000 (a kind gift from Gary Ward, University of Vermont(21)).[14C]Acetate Radiolabeling and Thin Layer Chromatography—

Parasites were grown in the absence and presence ATc for 48 h,and free tachyzoites were metabolically labeled as describedpreviously (9). Briefly, 108 tachyzoites were incubated with 10�Ci of sodium [14C]acetate (Movarek) in 1ml of DMEM for 4 hat 37 °C and 5% CO2. Total lipids were extracted with chloro-form/methanol (2:1). The extract was dried, and fatty acidmethyl esters were subjected to acidic methanolysis. Theresulting fatty acid methyl esters were extracted with hexaneand analyzed on RP-18 HPTLC plates developed in methanol/chloroform/water (75:25:5) and exposed to film for 1 week.Stable Isotope Labeling and Metabolomic Analyses—T. gon-

dii-infected fibroblasts were grown inDMEM in a T175 flask inthe absence or presence of ATc for 48 h. The medium wassupplemented with 8 mM [U-13C]glucose (final concentrationof [13C/12C]glucose was 16 mM) 24 h prior to egress. Free par-asites were separated from host cells by filtration through amembrane with a 3-�m pore size. Parasites were quenched byrapid chilling of the cell suspension in a dry ice/ethanol bath,and parasites were recovered by centrifugation (4000 � g, 25min, 0 °C). Cell pellets were washed three times with ice-coldPBS, and cell aliquots (2 � 108 cells) were transferred tomicro-centrifuge tubes and centrifuged (10,000� g, 30 s, 0 °C). Pelletswere suspended in chloroform (50 �l) and vortex-mixed thor-oughly prior to the addition of 200 �l of methanol/water (3:1).Samples were sonicated for 1 min at RT and then incubated at60 °C for 20 min. Extracts were centrifuged at 10,000 � g for 5min, and the supernatantwas subjected to biphasic partitioningby addition of 100 �l of water. The organic phase was trans-ferred to a partially sealed capillary formethanolysis, driedwithan internal standard of 1 nmol of scyllo-inositol, and subse-quently washed twice with methanol. Dried residues were dis-

solved in 0.5 N methanolic HCl (50 �l, Sigma), and the capillar-ies were sealed under vacuumand incubated at 95 °C overnight.Sampleswere neutralizedwith 10�l of pyridine (20min) beforebeing transferred to GC-MS vials, dried, and derivatized byaddition of 25 �l of N-methyl-N-(trimethylsilyl)-trifluoroacet-amide/trimethylchlorisane. Samples were analyzed on an Agi-lent 7890A-5975CGC-MS system. Splitless injection (injectiontemperature 280 °C) onto a 30 m � 10m � 0.25 mmDB-5MS �DG column (J&W, Agilent Technologies) was used, usinghelium as the carrier gas. The initial oven temperature was80 °C (2 min), followed by temperature gradients to 140 °C at30 °C/min, from 140 to 250 °C at 5 °C/min, and from 250 to265 °C at 15 °C/min. The final temperature was held for 10min.Data analysis was performed using Chemstation software(MSD Chemstation D.01.02.16, Agilent Technologies). Abun-dance and label incorporation was calculated as described pre-viously (22). Data shown are the average of three technical rep-licates and their standard deviation. They are representative oftwo biological experiments. Statistical significance was evalu-ated by Wilcoxon Rank Sum Testing with continuity correc-tions, using the R data analysis package (version 2.14.0), wherep values of �0.05 were considered significant.Yeast Media—Yeast strains were grown on YP (1% yeast

extract, 2% peptone) or yeast synthetic complete (SC) media(yeast synthetic dropoutmedium supplement (Sigma)), supple-mented with 0.67% Difco yeast nitrogen base (BD Biosciences),containing either 2% (w/v) D-galactose and 0.05% (w/v) D-glu-cose (GalGlu), 4% (w/v) D-glucose or 2% (w/v) raffinose as thecarbon source(s). For solid media, 2% agar was added. Selec-tions were carried out on plates lacking leucine (SCGal-Glu�Leu). Counter-selections against plasmids containing aURA3 marker were done on 5�-fluoroorotic acid plates (23)supplemented with GalGlu (5�FOAGalGlu). Liquid culturewith 4% (w/v) D-glucose was used to repress expression fromthe GAL1 promoter. Strains subjected to spotting assay weregrown overnight in YPGalGlu, kept for 24 h below an A600 �1,and then grown for 8 h in YPRaf. The cell density was normal-ized to an A600 � 0.5 before being spotted on YPGalGlu,SCGalUra GalGlu, and SC 5�FOAGalGlu.Yeast Strain Construction—The yeast strain Saccharomyces

cerevisiae W1536 �elo2�elo3/YCp33 GAL1 ScELO3 (URA3)(�ade2, �ade3, his3-11, his3-12, trp1-1, ura3-1, elo2::kanMX4,elo3::kanMX4) has been described before (24–26). The yeaststrain was transformed with either one of the following plas-mids (all LEU2): YCp111 GAL1-TgELO-B, YCp111 GAL1-TgELO-C, or YCpP111 GAL1-TgELO-A on SCGalGlu-Leu.(Primers used in the construction of these plasmids are listed insupplemental Table S3.) Candidate transformants on SCGal-Glu-Leu plates were picked and streaked on 5�-FOAGalGluplates to select against the presence of the YCp33 GAL1ScELO3 plasmid. Complementation of the �elo2�elo3 muta-tions was indicated by the production of a large amount ofFOA-resistant colonies. Presence of the plasmids carrying T.gondii constructs was verified by the ability of the FOA-resis-tant colonies to grow on SCGalGlu-Leu. These candidates wereused in further experiments. Strain W1536-5B �tsc13/YCplac33GAL1 ScTSC13 (MATa;�ade2,�ade3, his3-11, his3-12, trp1-1, ura3-1, tsc13::kanMX4) was transformed with



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YCp111GAL1-TgTSC13 on SCGalGlu-Leu, and colonies com-plemented by T. gondii TSC1were identified and isolated in ananalogous fashion as described above. Finally, yeast mutantscomplemented by T. gondii PHS1 were generated in a similarmanner, using strain W1536-5B �phs1/pYES2 PHS1 (MATa;�ade2, �ade3, his3-11, his3-12, trp1-1, ura3-1, phs1::kanMX4)transformed with YCp111 GAL1-TgPHS1.Preparation of ER Extracts fromYeast Cells—Yeast cells were

grown to exponential phase (2�106 cells�ml�1) and then har-vested by centrifuging for 15 min at 1300 � g, 4 °C. The pelletwas washed with sterile water and suspended in ice-cold lysisbuffer. The cells were disrupted with glass beads and centri-fuged for 15 min at 15,000 � g at 4 °C to remove the cell debris.The supernatants were centrifuged for 90 min at 85,000 � g ina Sorvall Ti70 rotor at 4 °C to generate pellet (P85) and super-natant (S85) fractions. The P85 was suspended in lysis bufferand used for the elongase assay. The protein concentrationswere determined by the method of Lowry using bovine serumalbumin as the standard. Equivalent amounts of total lysate,P85, and S85 were precipitated by adding trichloroacetic acid(TCA) to a 10% final concentration and processed forimmunoblotting.Yeast Fatty Acid Analysis—Yeast cultures were inoculated

from an overnight starter culture to anA600 of 0.05 in 100ml ofYPGalGlu, allowed to grow to anA600 of 0.5, and harvested by 5min centrifugation at 4300 � g. Cells were washed in 50 ml ofwater and harvested by centrifugation. The cell pellet was flash-frozen in liquid nitrogen and stored at �70 °C. Fatty acid anal-ysis was performed following methods published earlier (27)with minor modifications. Briefly, the cell pellets wereextracted with 1 ml of chloroform/methanol/formic acid (10:10:1), followed by a second extraction with 1 ml of chloroform/methanol/water (5:5:1 v/v). The combined organic phases werewashed with an equal volume of 1 M KCl in 0.2 M phosphoricacid and dried. Lipids were dissolved in 1 ml of toluene andtreated with 1ml ofmethanolic HCl (prepared by adding 0.5mlof acetyl chloride to 5 ml of cold dry methanol) overnight at50 °C. The solution was washed with 1 ml of 4 M KCl; theorganic phase was separated, and the aqueous phase wasextracted with 1ml of hexane. The organic phases were pooled,washed with 0.5 ml of 0.5 M ammonium bicarbonate, dried,redissolved in 100 �l of hexane, and subjected to gas chroma-tography on an HP6890 gas chromatograph with an HP5973mass analyzer and equipped with a J&W Scientific DB624 (30m � 0.32 mm, stationary phase 1.8 �m) column. 1 �l wasinjected splitless (detector block 250 °C) and subjected to atemperature program from 60 to 250 °C (20 °C/min) and a holdperiod of 35 min, flow 2 ml/min. Masses were scanned fromm/z 35 to 800, and a Wiley 275 spectral library was used foridentification.

RESULTS

Toxoplasma Is Capable of de Novo Fatty Acid Synthesis—Inapicomplexans, the enzymes of the FASII pathway are localizedto the apicoplast (4). In plants, the homologous chloroplastpathway is the sole source of de novo-synthesized fatty acids.We have previously constructed a T. gondiimutant with a con-ditional defect in this pathway. Although the growth of this

mutant was severely reduced upon inhibition of FASII, no dif-ference was observed in the rate of fatty acid biosynthesis asmeasured using [14C]acetate (9).We considered that FASIImaynot use acetyl-CoA generated from exogenous [14C]acetate. Totest this, we have established a new metabolic labeling proce-dure, in which the incorporation of 13C-derived from [U-13C]-glucose into the entire cellular pool of fatty acids is detected byGC-MS analysis (Fig. 1, A–C). When infected fibroblasts werelabeled with [U-13C]glucose, label was rapidly incorporatedinto tachyzoite lipids that were recovered from extracellularstages after host cell lysis and parasite egress. [U-13C]Glucoselabeling of extracellular tachyzoites resulted in negligible label-ing of parasite lipids indicating that de novo fatty acid synthesisoccurs predominantly in intracellular stages and decreases dra-matically following egress (data not shown). Host fibroblastcells are also capable of synthesizing fatty acids via a FASI sys-tem. In principal, 13C derived from exogenous [13C]glucosecould be incorporated into host fatty acids and subsequentlyscavenged by intracellular tachyzoites. To investigate theextent to which fibroblasts synthesize fatty acids, uninfectedfibroblasts weremetabolically labeledwith [U-13C]glucose, andincorporation of 13C into fatty acids was assessed byGC-MS. Incontrast to the intracellularly labeled tachyzoites, negligiblelabel was detected in the uninfected fibroblast lipids (Table 1).These experiments, together with the data presented below,indicate that 13C labeling of tachyzoite fatty acidswith [13C]glu-cose represents de novo fatty acid biosynthesis within the par-asite rather than uptake of labeled host lipids. In this context, itis notable that significant differences occur in the extent towhich different fatty acids are labeled. Although the long andvery long chain fatty acids, C14:0, C16:0, C16:1, C18:1, C20:1,and C26:1, were strongly labeled (60%) others, includingC18:0, C22:0, and C24:1, were labeled to less than 20%. Thesedifferences could reflect differences in the turnover andachievement of isotopic equilibrium in different fatty acidspools or, more likely, differences in the rate of salvage of “unla-beled” host fatty acids. Regardless, the results indicate that anumber of major fatty acids in intracellular stages are primarilysynthesized de novo rather than taken up from the host.Apicoplast FASII Is Required for Synthesis of Long Chain

Fatty Acids—The detection of multiple mass isotopomers ofthe long chain fatty acids, differing by 2 atomic mass units in[U-13C]glucose fed-parasites (Fig. 1B) indicated multiplerounds of incorporation of [13C]acetyl-CoA and [12C]acetyl-CoA into these chains, consistent with the operation of a FASsystem. To investigate the contribution of the apicoplast-lo-cated FASII complex to cellular fatty acid synthesis, fibroblastswere infected with the T. gondii �ACP/ACPi mutant thatexpresses components of the apicoplast FASII complex underthe control of a tetracycline-regulated promoter (9). When T.gondii �ACP/ACPi-infected fibroblasts were incubated with[U-13C]glucose in the absence of ATc, high levels of 13C incor-poration were observed in tachyzoite lipids (Fig. 1,D and E). Incontrast, when labelingwas done in the presence ofATc, result-ing in selective down-regulation of parasite FASII activity, thelabeling of tachyzoite fatty acids was strongly inhibited. In par-ticular, labeling of the long chain saturated and unsaturatedfatty acid C14:0 and C16:0 was reduced by 80% in the pres-



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ence of ATc (Fig. 1E). In a control experiment, ATc was foundto have no effect on the labeling of wild type tachyzoite fattyacids, indicating that ATc by itself does not inhibit [U-13C]glu-

cose uptake and catabolism to acetyl-CoA or fatty acid synthe-sis (Fig. 1C and Table 1). These experiments provide furtherdirect support for the conclusion that labeling of parasite fatty

FIGURE 1. Metabolic labeling of intracellular tachyzoites with [U-13C]glucose and analysis of de novo fatty acid biosynthesis. A, T. gondii-infected fibroblastswere precultured in the presence or absence of ATc for 24 h and then labeled with [U-13C]glucose 24 h prior to parasite egress. After host cell lysis, extracellulartachyzoites were metabolically quenched in a dry ice/ethanol bath and separated from host cell debris by filtration prior to metabolite extraction.Examples of differentlabeling patterns (13C indicated in gray) and their interpretation are schematically shown for myristate. B, lipids from [U-13C]glucose-labeled tachyzoites were sub-jected to methanolysis and trimethylsilyl derivatization and analyzed by GC-MS. A portion of the mass spectrum containing the molecular ions of the methyl ester ofmyristate is shown. Unlabeled myristoyl methyl ester (M0) has an m/z of 242 atomic mass units. Higher mass isotopomers (M1–12) contain between 1 and 12 13C atoms,corresponding to incorporation of 13C into the fatty acid (FA) biosynthetic pathways via [13C]acetyl-CoA. arb., arbitrary units. C, incorporation of 13C into the major fattyacids of intracellular wild type T. gondii tachyzoites (labeling is given as mol % of all labeled mass isotopomers (M1–12) relative to M0, after correction for naturalabundance). The fatty acid notation Cn:m indicates the length of the fatty acid (n, carbon number) and degree of unsaturation (m, number of double bonds). Note thattreatment of wild type parasites with ATc (black) does not result in any significant changes of the fatty acid labeling pattern when compared with untreated controls(white). D, intracellular tachyzoites of the T. gondii �ACP/ACPi mutant carrying an inducible copy of FASII ACP were labeled with [U-13C]glucose in the presence orabsence of ATc. Total lipids were extracted from the purified tachyzoites and released fatty acid methyl esters analyzed by GC-MS. The mass spectrum (molecular ionregion from m/z 240 to 250) of the myristate fatty acid methyl esters from unlabeled parasites, and from [13C]glucose-labeled parasites cultured in the absence orpresence of ATc is shown. Incorporation of multiple 13C2H4 units into myristate is observed in the absence of ATc (ACP active) and largely inhibited in the presence ofATc (ACP repressed). The spectrum is representative of three individual experiments. E, total 13C incorporation into fatty acids in �ACP/ACPi tachyzoites labeled in thepresence (black) or absence (white) of ATc. Error bars represent standard deviation where n � 3. Fatty acids for which significant changes in labeling were observed inthe presence and absence of ATc using the Wilcoxon rank sum test (p values less than 0.05) are indicated with an asterisk. The absolute amount of incorporation canvary from experiment to experiment depending on the stage of the culture (no ATc in C and E). We therefore always directly compare ATc-treated samples with anuntreated control from the same culture batch.



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acids with [U-13C]glucose reflects parasite de novo synthesisrather than salvage. They also suggest that the FASII pathway isdirectly responsible for the de novo synthesis of saturated longchain fatty acids (C14:0 and C16:0) that are utilized, at least inpart, for the synthesis of unsaturated and very long chain fattyacids.Growth Defects Due to Loss of FASII Pathway Are Only Par-

tially Restored by Chemical Complementation with Myristicand Palmitic Acid—We have previously shown that loss ofFASII activity leads to inhibition of parasite growth (9). As lossof FASII results in a deficiency in myristic and palmitic acidsynthesis, we investigated whether nutritional supplementa-tion of FASII-deficient parasites with these fatty acids wouldrestore growth. Parasite lines expressing red fluorescent pro-tein were cultured in 96-well plates in medium supplementedwith fatty acids complexed to BSA at concentrations rangingfrom 20 nM to 2 �M. These fatty acid concentrations have noeffect on the growth of the wild type parasites (data not shown).As expected, intracellular growth of the �ACP/ACPi mutantwas markedly reduced after 6 days in the presence of ATc (Fig.2A) (9). Growth inhibition in the presence of ATc was delayedby 2 days in the presence of a combination of 1 �Mmyristic and1 �M palmitic acid. However, fatty acid supplementation didnot restore wild type growth or prevent death over longer peri-ods, possibly reflecting inefficient transport to the parasite vac-uole, increased fatty acid demand at later time points, and/orthe involvement of FASII in other lipid biosynthetic pathways,such as the synthesis of lipoic acid.Parasites Express an Endoplasmic Reticulum-associated

Fatty Acid Elongation System—Several very long chain fattyacids were labeled with [U-13C]glucose in the absence of FASIIsuggesting that additional fatty acid biosynthetic pathways areactive during intracellular tachyzoite development. Bioinfor-matic analysis of the T. gondii genome indicates that these par-asites may have a functional fatty acid elongase system. Usingthe sequences of the components of the yeast elongation path-way as query in similarity searches, we found three candidategenes coding for fatty acid elongases and a single candidate foreach of the downstream reductases and the dehydratase in T.gondii (see supplemental Table S1).We evaluated the coding sequence for all six genes by cDNA

and rapid amplification of cDNA ends-PCR followed by sub-

cloning and sequencing. This analysis validated the computa-tionally predicted gene models for four genes: ELO-A–C andthe dehydratase, which is annotated as a hypothetical proteintyrosine phosphatase-like domain-containing protein in the T.gondii genome database. We established a diverging codingsequence for the presumptive enoyl-reductase and keto-acyl-CoA reductase (see “Experimental Procedures” and supple-mental Table S1 for details). The coding sequences were ampli-fied and introduced into a parasite expression vector thatappended a C-terminal epitope tag. These constructs weretransfected into parasites, and stable transgenic lines wereestablished through drug selection and cloning (see “Experi-mental Procedures” for details). The six epitope-tagged pro-teins localized to the same perinuclear structure in the parasite,as determined by immunofluorescence assays. For comparison,we transfected these lines with an expression plasmid for Der1-GFP (17), a marker for the T. gondii endoplasmic reticulum.Thismarker co-localizedwith the epitope-tagged proteins indi-cating that these enzymes localize to the endoplasmic reticu-lum (Fig. 3) (note that ELO-C had a more restricted pattern atthe apical face of the nuclear envelope reminiscent of the pre-viously characterized ER exit site (28)). These observations areconsistent with reports from other organisms where the com-ponents of the fatty acid elongation pathway have been found toform a multienzyme complex on the membrane of the endo-plasmic reticulum (29, 30).Candidate T. gondii Genes Rescue Lethal Yeast Fatty Acid

ElongationMutants—Wenext tested themolecular function ofeach of the six enzymes by heterologous expression in welldefined yeast mutants lacking equivalent enzymes. For thesephenotypic complementation experiments, we constructedyeast expression vectors under the control of the galactose-activated/glucose-repressed GAL1 promoter. Constructsencoding the putative T. gondii fatty acid elongases were trans-formed into S. cerevisiae strain W1536 �elo2�elo3/YCp33GAL1-ScELO3 (26). This mutant lacks the genes for both fattyacid elongase 2 and 3. This is a lethal double mutation (27), andthe strain is only viable due to episomal expression of elongase3 from aGAL1 promoter. Following transformation with para-site test plasmids, loss of the URA3-containing yeast plasmidwas achieved by counter-selection on media containing theURA3 activated toxic substrate 5�-fluoroorotic acid. T. gondiiELO-B and ELO-C but not ELO-A rescued the growth defi-ciency of the �elo2�elo3 mutant strain (Fig. 4A). Functionalcomplementation was further validated by incubation ofmicrosomal fractions prepared from the two respective com-plemented strains with radiolabeled malonyl-CoA and palmi-toyl-CoA (31). As expected, this resulted in chain elongation ofthe substrate (data not shown). Similarly, when expressed inyeast cells, the T. gondii candidates for dehydratase and enoylreductase were found to complement the growth of mutants intheir respective yeast homologs (S. cerevisiae Phs1p and Tsc13,see Fig. 4B and “Experimental Procedures” for detail). Overall,these experiments suggest that the T. gondii candidate genesindeed encode the enzymes of the fatty acid elongation system.We cannot make firm conclusions for TgELO-A and TgKCRthat failed to complement. This may be due to lack of expres-

TABLE 1Percent labeling of fatty acids derived from [U-13C]glucose-fedtachyzoites and HFFWild type RH parasites were used to infect HFF and labeled in situwith [13C]glu-cose. Uninfected HFF were labeled with lsqb]13C]glucose under identicalconditions. Percent labeling was determined by GC-MS analysis.

Cell type RH RH HFF

�/� ATc � � �C14:0 87.73 ( 3.86) 84.68 ( 0.50) NDC16:0 61.56 ( 0.58) 59.01 ( 1.09) 0.00 ( 0.00)C18:0 12.60 ( 0.55) 12.64 ( 0.73) 0.00 ( 0.00)C20:0 23.64 ( 12.81) 16.84 ( 6.41) 0.00 ( 0.00)C22:0 1.14 ( 2.95) 4.23 ( 1.05) 0.00 ( 0.00)C24:0 14.57 ( 4.41) 15.38 ( 1.53) 0.00 ( 0.00)C16:1 65.15 ( 1.62) 63.76 ( 0.47) 0.00 ( 0.00)C18:1 57.97 ( 3.88) 54.59 ( 0.15) 1.76 ( 1.77)C20:1 70.41 ( 0.82) 68.62 ( 0.71) 0.00 ( 0.00)C22:1 6.81 ( 8.79) 9.76 ( 3.44) 0.00 ( 0.00)C24:1 1.31 ( 1.87) 0.00 ( 2.58) 0.00 ( 0.00)C26:1 82.83 ( 4.14) 80.37 ( 2.75) ND



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sion or poor interaction of the heterologous enzymes with thenative yeast machinery.Genetic Ablation of Fatty Acid Elongases in T. gondii—To

further investigate the functional significance of the fatty acid

elongase system in Toxoplasma, we generated conditional par-asite mutants for the condensing enzymes of the pathway. Aparasite strain expressing epitope-tagged ELO-B under a tetra-cycline-regulated promoter from an ectopic locus was con-

FIGURE 2. Growth of �ACP/ACPi parasites in media supplemented with myristic and palmitic acid. Growth of �ACP/ACPi parasites stably expressing adTomatoRFP transgene was evaluated by measuring parasite fluorescence daily in a 96-well culture format (35). Parasites were grown in normal growth mediasupplemented with fatty acid-free BSA (A) or media supplemented with 1 �M of each myristic and palmitic acid coupled to BSA (3:1 molar ration) (B). Parasiteswere grown in the presence (black) or absence (white) of ATc. The average of fluorescence intensity (arbitrary units) for three independent replicates is shown,and error bars represent the standard deviation for each data point. Continuous culture in BSA-conjugated fatty acids (or higher serum supplementation) didnot lead to continuous growth of the ACP mutant in the presence of Atc. FA, fatty acids.

FIGURE 3. T. gondii fatty acid elongation pathway is localized to the parasite endoplasmic reticulum. Transgenic parasite lines were allowed to infectcoverslip cultures and were fixed and processed for immunofluorescence assay 24 h later. Transgenic proteins were marked with an HA (A–C) or a Myc (D–F)epitope tag detected with suitable antibodies (red channel). All strains carried a Der1GFP marker (green) previously shown to localize to the endoplasmicreticulum (17). DAPI and phase contrast images are shown for comparison. See supplemental Table S1 for reference to T. gondii gene models and “ExperimentalProcedures” for details on gene identification and cloning. Note that all six candidate fatty acid elongation enzymes are found to be associated with theendoplasmic reticulum. Tagged ELO-C produces a more restricted pattern at the apical side of the nuclear envelope.



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structed. In this background, we replaced the chromosomalcopy of this gene with a chloramphenicol resistance cassetteusing homologous recombination. Chloramphenicol-resistantparasites were screened by PCR to identify the clones withgenetic disruption for ELO-B. Using a similar strategy, we alsoengineered conditional mutants for ELO-C and ELO-A (see“Experimental Procedures” for details and note that some dele-tions were constructed using plasmids (9) and others with cos-mids (19)). Deletion of the respective locus was confirmed foreach mutant by Southern blot (Fig. 5, G–I). We evaluated theATc dependence of the expression of each of the genes byimmunofluorescence assay and Western blot assays. Forimmunofluorescence assay,mutant parasiteswere grown in theabsence and presence of ATc for 24 h prior to fixation andprocessing. ForWestern blots, parasites were grown in ATc for0–4 days. Both assays show rapid loss of the tagged proteinsupon ATc addition. The proteins are essentially undetectableafter 24 h of treatment (Fig. 5). To determine the effect of theloss of these proteins on parasite growth, we next performedplaque assays. Confluent host cell monolayers were infectedwith the mutant parasites in the absence or presence of ATc.Plaques in the host monolayer were visualized by staining after10 days. We detected no significant difference in plaque num-ber or size for each of the three elongase mutants (Fig. 5L). TheACPmutant was used as a positive control, and reduced plaqueformation was readily observed as described previously (9).Weconclude that individual loss of ELO-A, ELO-B, or ELO-C doesnot impede parasite growth in tissue culture.

T. gondii ELO Proteins Are Required for the Synthesis of VeryLong Chain Fatty Acids—Wedo not detect a growth phenotypein single elongase-deficient parasites. This could reflect thepossibility that fatty acids synthesized by the elongase systemare not essential, the ELO enzymes are functionally redundant(27), and/or the intracellular parasites can compensate for theloss of the pathways by salvaging necessary fatty acids from thehost. To determine the specific biochemical role of individualelongases, the fatty acid biosynthesis capabilities of eachmutant were initially analyzed using [14C]acetate labeling. Par-asites were grown in the absence or presence of ATc, liberatedfrom their host cells, and incubated in fatty acid-free mediumsupplemented with 10 �Ci of [14C]acetate. Lipids wereextracted after 4 h, and the corresponding fatty acid methylesters were analyzed by thin layer chromatography (Fig. 6). Theloss of ELO-B resulted in a moderate yet reproducible decreasein the labeling of all long chain fatty acids. Loss of ELO-C pro-duced the complete loss of labeling in a single very long chainfatty acid. In addition, we noted a marked increase in theabundance of a shorter fatty acid, suggesting that this may bea precursor that accumulates in the absence of the down-stream enzyme (Fig. 6C). Ablation of ELO-A did not producerecognizable changes in the labeling pattern. Although theseanalyses indicate clear changes in the biosynthetic capabili-ties of two of our mutants, they are limited in their sensitivityand resolution. In particular, they do not provide accuratemeasurement for the length and saturation of the affectedfatty acids.

FIGURE 4. T. gondii genes complement yeast fatty acid elongation mutants. Yeast mutants with deletions in specific genes encoding fatty acid elongationenzymes were transformed with plasmids carrying T. gondii candidate genes as detailed under “Experimental Procedures.” The resulting strains were used toconduct spotting assays on YPGalGlu media and demonstrate complementation (panels on the left). Control experiments using the same yeast cultures weredone on media that select against the yeast rescue plasmid and thus indicate that phenotypic complementation is due to presence of the T. gondii constructsonly (5�FOA-GalGlu), or media that indicate presence of the yeast rescue plasmids only (ScUra-GalGlu) (center and right panels, respectively). Yeast cultures wereprogressively diluted prior to plating from left to right as indicated. A, complementation analyses for the yeast elongase Sc�elo2�elo3 double mutant and theyeast (B) enoyl-CoA reductase (Sc�ECR) and dehydratase (Sc�DEH) mutants, respectively. Note that under restrictive conditions (5�FOA-GalGlu) growthdepends on the presence of the T. gondii genes. Only experiments that resulted in phenotypic complementation are shown here. See Table 2 for full detail onthe genotype of all yeast strains.



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ELO-B and ELO-C Produce Monounsaturated Long ChainFatty Acids in a Two-step Mechanism—To further investigatethe substrate specificity of the differentT. gondii fatty acid elon-gases, each of the ELO mutant parasites was metabolicallylabeled with [13C]glucose in absence or presence of ATc, andfatty acid mass isotopomers were analyzed by GC-MS. Thelabeling of saturated long chain fatty acids up to C16:0 wasunaffected by the individual loss of activity of ELO-A, ELO-B,and ELO-C (Fig. 7), consistent with their synthesis being regu-lated by FASII. Loss of ELO-A activity was associated with amarked reduction in labeling of saturated and unsaturated fattyacids with chain lengths of 18–24 carbons. Ablation of ELO-Aalso resulted in a reduction in the cellular levels of these fatty

acids. Significantly, both the abundance and level of labeling ofthe unsaturated fatty acid C16:1 increased in the absence ofELO-A. These results suggest that ELO-A is required for effi-cient elongation of C16:0 or C16:1 to longer species. In theabsence of ELO-A, de novo synthesized C16:0 is apparentlychanneled into increased synthesis of C16:1.In contrast, loss of expression of ELO-B or ELO-C resulted in

enhanced labeling of long chain fatty acids but reduced labelingof several very long chain fatty acids. Specifically, down-regula-tion of ELO-B activity severely ablated labeling of the monoun-saturated very long chain fatty acids C20:1, C22:1, C24:1, andC26:1 (Fig. 7A and Table 2). However, down-regulation ofELO-C activity led to a highly selective loss of labeling of C26:1

FIGURE 5. Conditional mutants lacking individual fatty acid elongases exhibit normal growth. We constructed conditional mutants for each of the threeT. gondii ELO genes. A–F, immunofluorescence assay detecting the expression of the HA epitope-tagged ectopic allele for the indicated ELO genes when grownin the absence (�ATc) or presence (�ATc) for 24 h. Merge images also show DAPI staining of DNA. G–I, Southern blot analysis probing genomic DNA from RHwild type, a strain carrying both the native and the conditional allele, and the mutant carrying only the conditional allele (from left to right for each of therespective mutants as indicated). Probes detect the respective coding region of each gene and are detailed in supplemental Table S2. In each blot the expectedsize of the restriction fragment for the native locus (e.g. ELO-B) and the conditional locus (e.g. ELO-Bi) is indicated with an arrowhead. Note loss of native loci. Jand K, Western blot analysis detecting ELO-Bi and ELO-Ci expression using an antibody against the HA tag. GRA8 is shown as a loading control. L, growth of theconditional ELO mutants was measured by plaque assay in absence and presence of ATc as indicated. �ACP/ACPi serves as positive control.

FIGURE 6. Radiolabeling analysis shows a reduction in long chain fatty acid synthesis in the T. gondii ELO-B and ELO-C mutant. T. gondii ELO mutants (A,�ELO-A; B, �ELO-B; and C, �ELO-C) were metabolically labeled with [14C]acetate; lipids were extracted, and fatty acid methyl esters were analyzed by reversephase thin layer chromatography, and representative autoradiographs for each mutant are shown. Each panel shows the parental strain carrying native andconditional locus (e.g. ELO-B/ELO-Bi) to the left and the mutant (�ELO-B/ELO-Bi) to the right grown in the absence (�) or presence (�) of ATc.



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and the accumulation of C22:1. These results suggest thatELO-B is largely responsible for the elongation of C18:1/C20:1to C22:1, whereas the primary role of ELO-C is to elongate fattyacids from C22:1 to C26:1. This model is also supported by ourmeasurements of overall fatty acid abundance (Fig. 7B). Loss ofELO-C results in a severe drop in the abundance of only C26:1,

in contrast, loss of ELO-B affected several shorter monounsat-urated fatty acids in addition toC26:1. This is reminiscent of thesituation in S. cerevisiae, where ScElo2p (equivalent toTgELO-B) is capable of elongating fatty acids up to 24 carbons,preferring a substrate with �22 carbon length. Likewise,ScElo3p and TgELO-C are alike in preferential production of

FIGURE 7. Loss of elongases results in selective reductions in the rate of synthesis and cellular levels of unsaturated fatty acids. Intracellular tachyzoitesof the three �ELO mutants were labeled with [U-13C]glucose in the presence (black) or absence (white) of ATc as detailed in Fig. 1. A, level of 13C incorporationinto the major fatty acids (FA) of the isolated tachyzoites. Error bars represent standard deviation where n � 3– 4. B, changes in the relative abundance ofindividual fatty acids in �ELO tachyzoites in the presence of ATc as measured by GC-MS (values are given relative to the abundance of this fatty acid species inthe no ATc control). Results are indicative of 3 individual replicates. Individual values for all experiments are listed in Table 4. Fatty acids for which significantchanges in labeling were observed in the presence and absence of ATc using the Wilcoxon Rank Sum Test (p values less than 0.05) are indicated with an asterisk.

TABLE 2Percent labeling of fatty acids derived from [U-13C]glucose-fed tachyzoitesMutant parasites were used to infect HFF and labeled in situwith [13C]glucose in the presence or absence of ATc. Percent labelingwas determined byGC-MS analysis. Fattyacids for which significant changes in labeling were observed in the presence and absence of ATc using the Wilcoxon rank sum test (p values less than 0.05) are indicatedby asterisk.

Cell type ACP ELOA ELOA ELOB ELOB ELOC ELOC

�/�ATc � � � � � � �C14:0 54.08* ( 0.99) 7.66* ( 1.78) ND ND ND ND ND NDC16:0 33.58* ( 0.68) 6.06* ( 0.22) 45.17 ( 0.64) 44.24 (0.19) 50.01 ( 0.45) 46.34 ( 1.47) 45.65 ( 1.28) 45.14 ( 0.35)C18:0 2.38* ( 0.51) 0.26* ( 0.28) 10.02* ( 0.50) 3.08* ( 0.03) 8.40 ( 0.40) 10.18 ( 0.54) 5.95 ( 0.53) 7.02 ( 0.18)C20:0 0.00 (0.00) 0.00 ( 0.00) 17.79* ( 3.14) 2.28* ( 1.60) 0.00 ( 0.00) 20.99 ( 8.83) 0.00 ( 3.65) 1.97 ( 7.81)C22:0 0.00 ( 0.00) 0.00 ( 0.39) 9.27 ( 1.61) ND 1.26 ( 2.64) 4.62 ( 3.55) 0.00 ( 2.47) 0.00 ( 0.00)C24:0 0.00 ( 4.82) 1.23 ( 0.08) 10.21* ( 1.47) 3.75*( 0.12) 4.18 ( 0.42) 8.87 ( 3.04) 3.12 ( 0.81) 3.17 ( 1.14)C16:1 17.76* ( 1.76) 10.01* ( 3.17) 35.63 ( 3.90) 47.04 ( 1.82) 41.27 ( 2.78) 25.38 ( 7.47) 23.94 ( 4.33) 36.37 ( 2.07)C18:1 16.37* ( 0.86) 4.96* ( 0.15) 31.95* ( 0.25) 5.42* ( 0.25) 20.15 ( 0.08) 27.88 ( 0.64) 28.86 ( 0.53) 28.83 ( 0.19)C20:1 24.05* ( 2.99) 11.21* ( 0.61) 45.56* ( 1.48) 21.54* ( 3.51) 40.60* ( 1.72) 8.46* ( 1.93) 43.64 ( 0.96) 45.09 ( 0.83)C22:1 0.00 ( 1.82) 1.05 ( 1.50) 0.51 ( 1.26) 0.55 ( 0.82) 20.12* ( 3.21) 0.00* ( 0.19) 4.91 ( 1.38) 24.13 (2.22)C24:1 0.31 ( 0.52) 0.00 ( 1.58) 8.36 ( 1.75) 5.29 ( 4.18) 9.42* ( 2.07) 2.63* ( 1.14) 8.10 ( 2.10) 8.39 ( 1.70)C26:1 36.45 ( 11.48) 27.56 ( 1.86) 59.32* ( 4.71) 47.30* ( 3.37) 70.80* ( 2.30) 0.00* ( 0.00) 59.41* ( 3.28) 0.00* ( 0.00)



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very long chain fatty acids. Note that both T. gondii enzymescomplement the yeast double mutant indicating that they canhave overlapping and slightly broader specificity.To distinguish between the potential activities of ELO-B, we

returned to the complemented yeast strains and analyzed theirfatty acid contents. The wild type yeast, mutant �elo2�elo3complemented S. cerevisiae ELO3, as well as the mutant com-plemented by expression of TgELO-B were grown on YPGal-Glu medium. Total lipids were extracted and analyzed byGC-MS (see under “Experimental Procedures”). As shown pre-viously, when compared with wild type, the �elo2�elo3/ScELO3mutant shows amuchdecreased level of fatty acidsC20and higher. The double deletion strain carrying the TgELO-Bconstruct, displayed some restoration of its longer chain fattyacids (C20–24) but not C26 or C26:1. Interestingly, thereseemed to be an overabundance of unsaturated fatty acids withno increase or only a modest increase in saturated fatty acids(see GC-MS tracks in Fig. 8, A and B). In particular, there was astriking accumulation of C22:1 compared with all other strains(see Tables 3 and 4 and Fig. 8A).

DISCUSSION

Intracellular pathogens need to either scavenge or synthesizelipids to maintain the integrity of their ownmembranes and/orthe membrane of specialized vacuoles within which they sur-vive and proliferate. Lipid and fatty acid salvage mechanisms

have been shown to be important for the intracellular growth ofseveral intracellular bacteria. Similarly, apicomplexan parasiteshave been shown to salvage lipids from the host endocytic path-way or cytoplasmic lipid bodies (12, 32–34, 36). Nonetheless,recent bioinformatic, genetic, and biochemical studies havesuggested that intracellular T. gondii tachyzoite stages expressmultiple pathways of fatty acid biosynthesis and may bedependent on de novo synthesis of fatty acids for the synthesisof bulk and specialized lipid classes (1). To dissect this complexnetwork of uptake and synthesis, we have combined reversegenetics with metabolomic analyses. We define the function ofindividual fatty acid synthesis pathways and have developed amodel that incorporates these findings (Fig. 9). The FASII path-way is localized to the apicoplast (Fig. 9, shown in green) and isthought to utilize acetyl-CoA derived from glycolytic triosephosphates that are imported into the apicoplast via a phos-phate translocator (19, 37). Our labeling studies provide strongsupport for this model. Specifically, although [14C]acetate wasnot effectively incorporated into FASII products, 13C derivedfrom [U-13C]glucose was strongly incorporated into this path-way. Ablation of FASII activity resulted in a global reduction infatty acid labeling. This was most apparent for the saturatedlong chain fatty acids C14:0 and C16:0, indicating that FASII islargely responsible for maintaining the cellular levels of thesefatty acids. The pattern of 13C labeling of these fatty acids inFASII-competent parasites was also consistent with de novosynthesis from short chain (�C4) fatty acid precursors. Synthe-sis of this precursor is most likely dependent on apicoplastFabD (38), but apicoplast ACP has also been demonstrated tobe capable of self-malonylation (39). Note that T. gondii alsoexpresses two differentially localized acetyl-CoA carboxylases,one in the apicoplast the other in the cytoplasm (40), that couldprovide malonyl-CoA for FASII and the elongation pathway,respectively. These findings demonstrate that the apicoplast,like the plant chloroplast, is themain cellular site of fatty acid de

FIGURE 8. GC-MS analysis of fatty acid methyl esters from S. cerevisiae complemented with T. gondii ELO-B. Fatty acids from yeast �elo2/�elo3 doubledeletion strain complemented by TgELO-B (see Fig. 5) were extracted and subjected to GC-MS analysis as described under “Experimental Procedures.” Anoverlay of the chromatogram of fatty acid methyl esters from this strain and the wild type is shown. A, enlargement of the region of the trace showing C22 andC24 methyl esters. B, region showing C26 methyl esters.

TABLE 3Yeast strains constructed for complementation analysis

Name Genotype of the yeast strain

Sc�ELO2�ELO3 � ScELO3 W1536 elo2�elo3�/YCp33 GAL1-ScELO3Sc�ELO2�ELO3 � TgELO-B W1536 �elo2�elo3/YCP111 GAL TgELO-BSc�ELO2�ELO3 � TgELO-C W1536 �elo2�elo3/YCP111 GAL TgELO-CSc�ECR � ScECR W1536 �tsc13/YCP33 GAL ScTsc13Sc�ECR � TgECR W1536 �tsc13/YCP111 GAL TgECRSc�DEH � ScDEH W1536 �phs1/YCP33 GAL ScPhs1Sc�DEH � TgDEH W1536 �phs1/YCP111 GAL TgDEH



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novo synthesis. There are a number of important functions forfatty acid synthesis directly in the plastid, including lipoylation,acylation of plastid proteins, and the synthesis of plastid lipids.Indeed, the loss of FASII in the apicoplast and the plant chlo-roplast perturbs organelle biogenesis and division leading toorganelle loss (9, 41).Does this suggest the significance for FASII to be entirely

local? Following this idea, the main function of FASII could betomaintain the apicoplast to support the organelle as the site ofisoprenoid precursor synthesis (42, 43). In plant chloroplasts,FASII-derived fatty acids are rapidly exported from the plastidinto the cytoplasm (44). Our experiments do not formally testthis and do not establish the localization of FASII-derived lip-ids. We note, however, that our measurements indicate thatmost (60–80%) of the parasiteC14:0 andC16:0 are 13C-labeled,a finding that is inconsistent with FASII only supplying a smallorganellar pool of lipids. Moreover, our data suggest that C16:0and C16:1 fatty acids synthesized by the apicoplast FASII com-plex are further elongated by the ER elongases, indicating that asignificant fraction of the apicoplast-synthesized fatty acids are

exported to the ER. We propose that the apicoplast is the mainsource of C14:0 and C16:0 for intracellular stages of T. gondiiand the liver stage of Plasmodium, but in the Plasmodiumerythrocyte and mosquito stages, this need can be satisfied bysalvage of fatty acids from the host. A greater dependence of T.gondii tachyzoites on de novo fatty acid synthesis is further sup-ported by our finding that loss of FASII was only partially res-cued by providing excess C14:0 and C16:0 in the culturemedium. Overall, these observations are consistent withimportant functions of FASII-derived products within andbeyond the apicoplast in T. gondii.In addition to the FASII complex, we provide direct evidence

for the activity of an elaborate fatty acid elongation pathwaylocalized to the parasite endoplasmic reticulum. This fatty acidelongation pathway consists of four enzymatic steps. Theextension of the carbon chain is initiated by a condensationreaction between acyl-CoA and malonyl-CoA catalyzed by afatty acid elongase. This is followed by reduction through keto-acyl-CoA reductase, dehydration to enoyl-CoA, and finalreduction to the elongated fatty acyl-CoA. We identified a full

TABLE 4Fatty acid profiles of yeast wild type and complement elo2/elo3 mutant strainValues indicated are % of relative response units of total fatty acid methyl esters. p values were calculated using a two-tailed Student’s t test, setting not detected fatty acidspecies to 0. The results of fatty acid extractions from two independent sets of cultures are shown. Fatty acyl species were identified by GC/MALDI mass spectroscopy asdescribed under “Experimental Procedures.” ND means not detected.

C14 C15 C16:1/16 C18:1/18 C20:1/20 C22:1 C22 C24:1 C24 C26 C26-OH

TgELO-B 2.5 1.2 4.4 1.9 26.1a 3.8 36.3 5.0 12.4b 3.2 14.8b 1.3 2.2 1 1.6b 0.2 ND/0.1 ND NDa

TgELO-C 2.1 0.6 3.6 2.3 47.2 3.9 45.0a 0.4 2.2 1.6 0.2 / ND ND ND ND ND NDa

ScELO3 2.5 0.1 2.4 1.5 40.6 0.8 51.9 1.8 ND ND ND ND ND 1.1 0.6 1 0.1WT 3.1 1.4 1.6 1.5 47.2 6.5 46.1 2.2 0.5 0.1 0.3 0.2 0.5 0.5 ND 0.1/ND 0.6 0,4 0.7/ND

a p � 0.05 is only in comparison with the S. cerevisiae elo2/elo3 double mutant complemented with S. cerevisiae ELO3.b p � 0.05 is in comparison with both S. cerevisiaeWT and elo2/elo3 double mutant complemented with S. cerevisiae ELO3.

FIGURE 9. Toxoplasma acquires fatty acids through a complex network of synthesis and uptake. A, T. gondii (pink) is an intracellular pathogen capable offatty acid and lipid salvage from the host cell (blue). This process can intersect host cell import as well as synthesis routes. B, in addition, the parasite harborsthree fatty acid synthesis pathways that are localized to different cellular compartments. C, apicoplast (green)-localized FASII pathway produces significantamounts of myristic and palmitic acid in addition to lipoic acid relying on cytoplasmic glycolysis for precursors. E, Toxoplasma also maintains an ER-associatedelongase system that synthesizes very long chain monounsaturated fatty acids, subsequently using the activity of ELO-B and ELO-C. D, T. gondii FASI remainslargely uncharacterized. Its stage-specific expression pattern and localization are not established. It is also unclear whether this megasynthase synthesizes fattyacids de novo like the FASI of humans or acts as an elongase for saturated fatty acids, as demonstrated for the FASI of C. parvum. Major products are highlightedin red. Des, desaturase; PEP, phosphoenolpyruvate; Mal, malonate; Ac, acetate; vlcFA, very long chain fatty acids.



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complement of six enzymes and support the sequence similar-ity-based assignments experimentally by yeast complementa-tion analysis for several candidates. Our [U-13C]glucose-labeling experiments in parasite mutants and heterologousexpression in yeast mutants suggest that each of the elongasesmay act sequentially as outlined schematically in Fig. 9E. TheT.gondii elongation systemgenerates very long chainmonounsat-urated fatty acids. ELO-B elongates C18:1 to C22:1 in two car-bon increments. Based on the yeast data, its activity fromC20:1to C22:1 appears to be particularly robust. C22:1 is elongated intwo final rounds by ELO-C to C26:1 (note that we do not fullyresolve whether the synthesis of C24 is an overlapping functionbetween the two elongases). The T. gondii ELO-B and ELO-Cpathway is the sole source of C26:1 for the parasite and does notappear to elongate saturated fatty acids. The significance of thisstriking preference for unsaturated fatty acids is unclear at thispoint. Unsaturated fatty acids have distinct physical propertiesresulting in differences in e.g.membrane fluidity, and thosemaybemore suited to the needs of the parasite. It is notable that thelevels of C26:1 in fibroblasts are very low (Table 1 and Fig. 10),possibly accounting for the inability to generate multiple ELOgenes.What is the source of the ELO-B substrates? Monounsatu-

rated fatty acids can be synthesized by the desaturation of acorresponding saturated fatty acid. A route used by manyorganisms is the conversion of C18:0 to C18:1 oleic acid bystearoyl-CoA desaturase (45). This enzyme is present inT. gon-dii (TGGT1_053030) and has been characterized in detail in P.falciparum (46). In Plasmodium, it was found to be localized tothe membrane of the endoplasmic reticulum and thus couldwork hand in hand with the ELO system. This system couldpotentially tap into the FASII-generated fatty acid pool throughthe activity of ELO-A, which appears to be required for elonga-

tion of C16:0 and C16:1 to C18:0 and C18:1, respectively. Spe-cifically, ablation of ELO-A resulted in a marked decrease insynthesis of C18:0, which was associated with the concomitantincrease in both the labeling and steady state concentration ofC16:0/C16:1 (Fig. 7). It is notable that only a minor pool ofC18:0 is labeled with [U-13C]glucose in wild type parasites(�10%) indicating that this fatty acid is primarily salvaged fromthehost. In contrast, C18:1 is labeled to a similar extent asC16:0and C16:1 (40%). These findings suggest that ELO-A primar-ily utilizes C16:0 synthesized in the apicoplast and C16:1 syn-thesized by the putative tachyzoite ER desaturase. The reducedcellular levels and rates of labeling of longer chain saturated/unsaturated fatty acids in ELO-A ablated parasites indicate thatthe C18:0 and C18:1 synthesized by ELO-A is selectively uti-lized by ELO-B and ELO-C, whereas the larger cellular pool ofsalvaged C18:0 appears to be poorly used. Collectively, theseobservations indicate that synthesis of very long chain fattyacids in intracellular tachyzoites involves extensive intraorgan-ellar transport as well as some degree of substrate channeling. Itshould be noted that theT. gondii genome encodes a FASI path-way (Fig. 9D), but its role in the T. gondii fatty acid metabolismis not yet established.Although we demonstrate clear biochemical effects of sup-

pressing individual ELO genes, we do not measure parasitegrowth impairment under tissue culture conditions. It is possi-ble that the parasite can satisfy its needs through uptake. Arecently described lipid uptake system in T. gondii may facili-tate this process (47). Alternatively, the lack of growth defectsmay be due to redundancy within the ELO system itself. Inyeast, the loss of a fatty acid elongase 2 or fatty acid elongase 3can be tolerated. However, loss of both elongases results in syn-thetic lethality. It has been suggested that the enzymes partiallycomplement each other and that the cells can tolerate the

FIGURE 10. Total fatty acid composition of uninfected HFF host cells and RH tachyzoites. The total lipid extract of human foreskin fibroblasts (HFF) (A) andisolated purified tachyzoites (labeled in their host cells as detailed in the “Experimental Procedures”) (B) was subjected to solvolysis in methanolic-HCl andderivatized in trimethylsilyl reagent, and fatty acid methyl esters were detected by GC-MS. The mol % of the major fatty acids are shown (mean S.D.). Thelower panels show same data set scaled to a maximum of 5% fractional abundance to display low abundance species.



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resulting slight differences in their fatty acid pools (27). Ouryeast complementation results indicate that the T. gondiienzymes may also have overlapping substrate specificity. Toestablish whether a similar redundancy exists in T. gondii, wesought to engineer a more complete ablation of the parasiteELO system. Our first approach was to attempt to generate aconditional double mutant in ELO-B and -C. However, ourattempts to generate such a mutant failed recurrently despitethe fact that gene targeting was efficient for individual loci inprevious experiments (data not shown). An alternativeapproach would be to delete a nonredundant downstreamenzyme of the pathway, thereby affecting the activity of all threeelongases. We attempted to generate conditional mutants fordehydratase and enoyl-CoA reductase. Again, we were unsuc-cessful in generating amutant (data not shown). These negativeresults do not formally establish the essential nature of the T.gondii ELO pathway, but they are consistent with such a role.Which aspect of parasite biology generates the heavy need

for elongated fatty acids served by these elaborate and poten-tially redundant synthesis and uptake pathways? Very longchain fatty acids are required for the synthesis of highly special-ized lipids such as waxes in plants, but they are also importantcomponents of phospholipids (27, 48). Loss of the elongationpathway affects the overall phospholipid pool in these organ-isms. T. gondii harbors a variety of phospholipids that not onlydiffer in their headgroup but also in the length of the acyl chains(49). The parasites can take up phospholipids but appear todepend on the synthesis of some species (47, 50, 51). Overall,our results demonstrate that despite its substantial complexitythe parasite fatty acid metabolism can be unraveled in molecu-lar detail. Stable isotope labeling followed by GC-MS can trackand quantify the synthesis of a large number of metabolites invivo and was particularly useful for resolving the activities ofdifferent fatty acid biosynthetic enzymes. Furthermore, the factthat 13C-labeled precursors can be added at normal physiolog-ical concentrationsmay increase the efficiency with which theycan access intracellular compartments such as the T. gondiiparasitophorous vacuole rather than being diverted into hostcell metabolism. Conditional gene ablation provides focus andspecificity to this broader metabolomic analysis. We providefunctional annotation for two important parasite pathwayshere and suggest that this strategy could be used to great effectto understand the metabolic interaction between host and par-asite in T. gondii and many other intracellular pathogens.

Acknowledgments—We thank Dr. Ulrich Bergmann (Department ofBiochemistry, University of Oulu) andDr. Päivi Joensuu (Departmentof Chemistry, University of Oulu) for assistance with the yeast fattyacid analyses and BenjaminWoodcroft (Department of Biochemistryand Molecular Biology, University of Melbourne) for assistance withthe statistical analyses and ApiLoc annotations.

REFERENCES1. Mazumdar, J., and Striepen, B. (2007) Make it or take it. Fatty acid metab-

olism of apicomplexan parasites. Eukaryot. Cell 6, 1727–17352. Valdivia, R. H. (2008) Chlamydia effector proteins and new insights into

chlamydial cellular microbiology. Curr. Opin. Microbiol. 11, 53–593. Bhatt, A., Molle, V., Besra, G. S., Jacobs, W. R., Jr., and Kremer, L. (2007)

TheMycobacterium tuberculosis FAS-II condensing enzymes. Their rolein mycolic acid biosynthesis, acid-fastness, pathogenesis, and in futuredrug development.Mol. Microbiol. 64, 1442–1454

4. Waller, R. F., Keeling, P. J., Donald, R. G., Striepen, B., Handman, E.,Lang-Unnasch, N., Cowman, A. F., Besra, G. S., Roos, D. S., and McFad-den, G. I. (1998) Nuclear encoded proteins target to the plastid in Toxo-plasma gondii and Plasmodium falciparum. Proc. Natl. Acad. Sci. U.S.A.95, 12352–12357

5. Hiltunen, J. K., Chen, Z., Haapalainen, A. M., Wierenga, R. K., and Kas-taniotis, A. J. (2010) Mitochondrial fatty acid synthesis. An adopted set ofenzymes making a pathway of major importance for the cellular metabo-lism. Prog. Lipid Res. 49, 27–45

6. Goodman, C. D., and McFadden, G. I. (2007) Fatty acid biosynthesis as adrug target in apicomplexan parasites. Curr. Drug Targets 8, 15–30

7. Surolia, N., and Surolia, A. (2001) Triclosan offers protection againstblood stages of malaria by inhibiting enoyl-ACP reductase of Plasmodiumfalciparum. Nat. Med. 7, 167–173

8. McLeod, R., Muench, S. P., Rafferty, J. B., Kyle, D. E., Mui, E. J., Kirisits,M. J., Mack, D. G., Roberts, C. W., Samuel, B. U., Lyons, R. E., Dorris, M.,Milhous, W. K., and Rice, D. W. (2001) Triclosan inhibits the growth ofPlasmodium falciparum and Toxoplasma gondii by inhibition of apicom-plexan Fab I. Int. J. Parasitol. 31, 109–113

9. Mazumdar, J. H., Wilson, E., Masek, K. A., Hunter, C., and Striepen, B.(2006) Apicoplast fatty acid synthesis is essential for organelle biogenesisand parasite survival in Toxoplasma gondii. Proc. Natl. Acad. Sci 103,13192–13197

10. Vaughan, A. M., O’Neill, M. T., Tarun, A. S., Camargo, N., Phuong, T. M.,Aly, A. S., Cowman, A. F., and Kappe, S. H. (2009) Type II fatty acidsynthesis is essential only formalaria parasite late liver stage development.Cell. Microbiol. 11, 506–520

11. Yu,M., Kumar, T. R., Nkrumah, L. J., Coppi, A., Retzlaff, S., Li, C. D., Kelly,B. J., Moura, P. A., Lakshmanan, V., Freundlich, J. S., Valderramos, J. C.,Vilcheze, C., Siedner,M., Tsai, J. H., Falkard, B., Sidhu, A. B., Purcell, L. A.,Gratraud, P., Kremer, L.,Waters, A. P., Schiehser, G., Jacobus, D. P., Janse,C. J., Ager, A., Jacobs, W. R., Jr., Sacchettini, J. C., Heussler, V., Sinnis, P.,and Fidock, D. A. (2008) The fatty acid biosynthesis enzyme FabI plays akey role in the development of liver-stage malarial parasites. Cell HostMicrobe 4, 567–578

12. Charron, A. J., and Sibley, L. D. (2002) Host cells. Mobilizable lipid re-sources for the intracellular parasite Toxoplasma gondii. J. Cell Sci. 115,3049–3059

13. Gerold, P., Dieckmann-Schuppert, A., and Schwarz, R. T. (1994) Glyco-sylphosphatidylinositols synthesized by asexual erythrocytic stages of themalarial parasite, Plasmodium falciparum. Candidates for plasmodial gly-cosylphosphatidylinositol membrane anchor precursors and pathogenic-ity factors. J. Biol. Chem. 269, 2597–2606

14. Zhu, G. (2004) Current progress in the fatty acid metabolism in Crypto-sporidium parvum. J. Eukaryot. Microbiol. 51, 381–388

15. Zhu, G., Li, Y., Cai, X., Millership, J. J., Marchewka, M. J., and Keithly, J. S.(2004) Expression and functional characterization of a giant Type I fattyacid synthase (CpFAS1) gene from Cryptosporidium parvum. Mol.Biochem. Parasitol. 134, 127–135

16. Zhu, G., Shi, X., and Cai, X. (2010) The reductase domain in a Type I fattyacid synthase from the apicomplexan Cryptosporidium parvum. Re-stricted substrate preference towards very long chain fatty acyl thioesters.BMC Biochem. 11, 46–46

17. Agrawal, S., van Dooren, G. G., Beatty, W. L., and Striepen, B. (2009)Genetic evidence that an endosymbiont-derived endoplasmic reticulum-associated protein degradation (ERAD) system functions in import of api-coplast proteins. J. Biol. Chem. 284, 33683–33691

18. Striepen, B., and Soldati, D. (2007) in Toxoplasma gondii, the Model Api-complexan- Perspective and Methods (Weiss, L. D., and Kim, K., eds) pp.391–415, Elsevier, London

19. Brooks, C. F., Johnsen, H., van Dooren, G. G., Muthalagi, M., Lin, S. S.,Bohne,W., Fischer, K., and Striepen, B. (2010) TheToxoplasma apicoplastphosphate translocator links cytosolic and apicoplast metabolism and isessential for parasite survival. Cell Host Microbe 7, 62–73

20. Meissner, M., Schlüter, D., and Soldati, D. (2002) Role of Toxoplasma



at Australian N

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AU

L), on Novem

ber 4, 2012w

ww

.jbc.orgD

ownloaded from

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gondii myosin A in powering parasite gliding and host cell invasion. Sci-ence 298, 837–840

21. Carey, K. L., Donahue, C. G., and Ward, G. E. (2000) Identification andmolecular characterization of GRA8, a novel, proline-rich, dense granuleprotein of Toxoplasma gondii. Mol. Biochem. Parasitol. 105, 25–37

22. Saunders, E. C., Ng,W.W., Chambers, J. M., Ng,M., Naderer, T., Krömer,J. O., Likic, V. A., and McConville, M. J. (2011) Isotopomer profiling ofLeishmaniamexicana promastigotes reveals important roles for succinatefermentation and aspartate uptake in tricarboxylic acid cycle (TCA)anaplerosis, glutamate synthesis, and growth. J. Biol. Chem. 286,27706–27717

23. Amberg, D. C., Burke, D., Strathern, J. N., and Cold Spring Harbor, L.(2005)Methods in Yeast Genetics, p. 207, Cold Spring Harbor LaboratoryPress, Cold Spring Harbor, NY

24. Kastaniotis, A. J., Autio, K. J., Sormunen, R. T., and Hiltunen, J. K. (2004)Htd2p/Yhr067p is a yeast 3-hydroxyacyl-ACP dehydratase essential formitochondrial function and morphology.Mol. Microbiol. 53, 1407–1421

25. Song,W.Q., Qin, Y.M., Saito,M., Shirai, T., Pujol, F.M., Kastaniotis, A. J.,Hiltunen, J. K., and Zhu, Y. X. (2009) Characterization of two cottoncDNAs encoding trans-2-enoyl-CoA reductase reveals a putative novelNADPH-binding motif. J. Exp. Bot. 60, 1839–1848

26. Qin, Y. M., Pujol, F. M., Hu, C. Y., Feng, J. X., Kastaniotis, A. J., Hiltunen,J. K., and Zhu, Y. X. (2007) Genetic and biochemical studies in yeast revealthat the cotton fiber-specific GhCER6 gene functions in fatty acid elonga-tion. J. Exp. Bot. 58, 473–481

27. Oh, C. S., Toke, D. A., Mandala, S., and Martin, C. E. (1997) ELO2 andELO3, homologues of the Saccharomyces cerevisiae ELO1 gene, functionin fatty acid elongation and are required for sphingolipid formation. J. Biol.Chem. 272, 17376–17384

28. Hager, K.M., Striepen, B., Tilney, L. G., and Roos, D. S. (1999) The nuclearenvelope serves as an intermediary between the ER and Golgi complex inthe intracellular parasite Toxoplasma gondii. J. Cell Sci. 112, 2631–2638

29. Cinti, D. L., Cook, L., Nagi, M. N., and Suneja, S. K. (1992) The fatty acidchain elongation system of mammalian endoplasmic reticulum. Prog.Lipid Res. 31, 1–51

30. Denic, V., and Weissman, J. S. (2007) A molecular caliper mechanism fordetermining very long-chain fatty acid length. Cell 130, 663–677

31. Qin, Y. M., Pujol, F. M., Shi, Y. H., Feng, J. X., Liu, Y. M., Kastaniotis, A. J.,Hiltunen, J. K., and Zhu, Y. X. (2005) Cloning and functional character-ization of two cDNAs encoding NADPH-dependent 3-ketoacyl-CoA re-ductased from developing cotton fibers. Cell Res. 15, 465–473

32. Coppens, I., Dunn, J. D., Romano, J. D., Pypaert,M., Zhang, H., Boothroyd,J. C., and Joiner, K. A. (2006) Toxoplasma gondii sequesters lysosomesfrom mammalian hosts in the vacuolar space. Cell 125, 261–274

33. Mikolajczak, S. A., Jacobs-Lorena, V., MacKellar, D. C., Camargo, N., andKappe, S. H. (2007) L-FABP is a critical host factor for successful malarialiver stage development. Int. J. Parasitol. 37, 483–489

34. Sharma, A., Yogavel, M., Akhouri, R. R., Gill, J., and Sharma, A. (2008)Crystal structure of soluble domain of malaria sporozoite protein UIS3 incomplex with lipid. J. Biol. Chem. 283, 24077–24088

35. Gubbels, M. J., Li, C., and Striepen, B. (2003) High-throughput growthassay forToxoplasma gondii using yellow fluorescent protein.Antimicrob.Agents Chemother. 47, 309–316

36. Coppens, I. (2006) Contribution of host lipids to Toxoplasma pathogene-sis. Cell. Microbiol. 8, 1–9

37. Lim, L., Linka, M., Mullin, K. A.,Weber, A. P., andMcFadden, G. I. (2010)

The carbon and energy sources of the nonphotosynthetic plastid in themalaria parasite. FEBS Lett. 584, 549–554

38. Prigge, S. T., He, X., Gerena, L., Waters, N. C., and Reynolds, K. A. (2003)The initiating steps of a type II fatty-acid synthase in Plasmodium falcip-arum are catalyzed by pfACP, pfMCAT, and pfKASIII. Biochemistry 42,1160–1169

39. Misra, A., Surolia, N., and Surolia, A. (2009) Catalysis and mechanism ofmalonyl transferase activity in type II fatty acid biosynthesis acyl carrierproteins.Mol. Biosyst. 5, 651–659

40. Jelenska, J., Crawford, M. J., Harb, O. S., Zuther, E., Haselkorn, R., Roos,D. S., and Gornicki, P. (2001) Subcellular localization of acetyl-CoA car-boxylase in the apicomplexan parasite Toxoplasma gondii. Proc. Natl.Acad. Sci. U.S.A. 98, 2723–2728

41. Wu, G. Z., and Xue, H. W. (2010) Arabidopsis �-ketoacyl-[acyl carrierprotein] synthase I is crucial for fatty acid synthesis and plays a role inchloroplast division and embryo development. Plant Cell 22, 3726–3744

42. Nair, S. C., Brooks, C. F., Goodman, C. D., Strurm, A., McFadden, G. I.,Sundriyal, S., Anglin, J. L., Song, Y., Moreno, S. N., and Striepen, B. (2011)Apicoplast isoprenoid precursor synthesis and the molecular basis of fos-midomycin resistance inToxoplasma gondii. J. Exp.Med. 208, 1547–1559

43. Yeh, E., and DeRisi, J. L. (2011) Chemical rescue of malaria parasites lack-ing an apicoplast defines organelle function in blood-stage Plasmodiumfalciparum. PLoS Biol. 9, e1001138

44. Koo, A. J., Ohlrogge, J. B., and Pollard, M. (2004) On the export of fattyacids from the chloroplast. J. Biol. Chem. 279, 16101–16110

45. Stukey, J. E., McDonough, V. M., andMartin, C. E. (1990) TheOLE1 geneof Saccharomyces cerevisiae encodes the �9 fatty acid desaturase and canbe functionally replaced by the rat stearoyl-CoA desaturase gene. J. Biol.Chem. 265, 20144–20149

46. Gratraud, P., Huws, E., Falkard, B., Adjalley, S., Fidock, D. A., Berry, L.,Jacobs, W. R., Jr., Baird, M. S., Vial, H., and Kremer, L. (2009) Oleic acidbiosynthesis in Plasmodium falciparum: characterization of the stearoyl-coA desaturase and investigation as a potential therapeutic target. PLoSONE 4, e6889

47. Lige, B., Jayabalasingham, B., Zhang, H., Pypaert, M., and Coppens, I.(2009) Role of an ancestral d-bifunctional protein containing two sterol-carrier protein-2 domains in lipid uptake and trafficking in Toxoplasma.Mol. Biol. Cell 20, 658–672

48. Bach, L., Michaelson, L. V., Haslam, R., Bellec, Y., Gissot, L., Marion, J., DaCosta, M., Boutin, J. P., Miquel, M., Tellier, F., Domergue, F., Markham,J. E., Beaudoin, F., Napier, J. A., and Faure, J. D. (2008) The very long-chainhydroxy fatty acyl-CoA dehydratase PASTICCINO2 is essential and lim-iting for plant development. Proc. Natl. Acad. Sci. U.S.A. 105,14727–14731

49. Welti, R., Mui, E., Sparks, A., Wernimont, S., Isaac, G., Kirisits, M., Roth,M., Roberts, C.W., Botté, C., Maréchal, E., andMcLeod, R. (2007) Lipido-mic analysis of Toxoplasma gondii reveals unusual polar lipids. Biochem-istry 46, 13882–13890

50. Coppens, I., and Vielemeyer, O. (2005) Insights into unique physiologicalfeatures of neutral lipids in Apicomplexa. From storage to potential me-diation in parasite metabolic activities. Int. J. Parasitol. 35, 597–615

51. Gupta, N., Zahn, M. M., Coppens, I., Joiner, K. A., and Voelker, D. R.(2005) Selective disruption of phosphatidylcholine metabolism of the in-tracellular parasite Toxoplasma gondii arrests its growth. J. Biol. Chem.280, 16345–16353



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