Báo cáo khoa học: Trypanosoma brucei: a model micro-organism to study eukaryotic phospholipid biosynthesis docx

REVIEW ARTICLE Trypanosoma brucei: a model micro-organism to study eukaryotic phospholipid biosynthesis Mauro Serricchio and Peter Bu ¨ tikofer Institute of Biochemistry and Molecular Medicine, University of Bern, Switzerland Introduction Trypanosoma brucei is a eukaryotic protozoan parasite causing African sleeping sickness in humans and nagana in domestic animals. During its complex life cycle, it migrates between the blood and tissue fluids of a mammalian host and several compartments of the insect vector, the tsetse fly. Trypanosoma brucei is not only a devastating pathogen, affecting social and eco- nomic development in sub-Saharan Africa, but has also become an interesting model organism to study general biological processes. RNA editing [1], glycosyl- phosphatidylinositol (GPI) anchoring [2], trans-splicing [3] and antigenic variation [4] represent biological phe- nomena that were initially discovered in trypanosomes and have later been observed in other eukaryotic organisms as well. The T. brucei genome with  9000 protein-coding genes has been sequenced [5] and the parasite is amenable to reverse genetic approaches, such as gene knockout by homologous recombination, Keywords biosynthesis; eukaryote; glycerophospholipid; phospholipid; RNAi; sphingophospholipid; trypanosome Correspondence P. Bu ¨ tikofer, Institute of Biochemistry & Molecular Medicine, University of Bern, Bu ¨ hlstrasse 28, 3012 Bern, Switzerland Fax: +41 31 631 3737 Tel: +41 31 631 4113 E-mail: peter.buetikofer@mci.unibe.ch Website: http://ntbiomol.unibe.ch/Buetikofer/ (Received 12 November 2010, revised 23 December 2010, accepted 7 January 2011) doi:10.1111/j.1742-4658.2011.08012.x Although the protozoan parasite, Trypanosoma brucei, can acquire lipids from its environment, recent reports have shown that it is also capable of de novo synthesis of all major phospholipids. Here we provide an over- view of the biosynthetic pathways involved in phospholipid formation in T. brucei and highlight differences to corresponding pathways in other eukaryotes, with the aim of promoting trypanosomes as an attractive model organism to study lipid biosynthesis. We show that de novo synthesis of phosphatidylethanolamine involving CDP-activated intermediates is essential in T. brucei and that a reduction in its cellular content affects mitochondrial morphology and ultrastructure. In addition, we highlight that reduced levels of phosphatidylcholine inhibit nuclear division, suggest- ing a role for phosphatidylcholine formation in the control of cell division. Furthermore, we discuss possible routes leading to phosphatidylserine and cardiolipin formation in T. brucei and review the biosynthesis of phosphati- dylinositol, which seems to take place in two separate compartments. Finally, we emphasize that T. brucei represents the only eukaryote so far that synthesizes all three sphingophospholipid classes, sphingomyelin, inosi- tolphosphorylceramide and ethanolaminephosphorylceramide, and that their production is developmentally regulated. Abbreviations CEPT, CDP-choline ⁄ ethanolamine:diacylglycerol phosphotransferase; CL, cardiolipin; CPT, CDP-choline:diacylglycerol phosphotransferase; CT, CTP:phosphocholine cytidylyltransferase; EPC, ethanolaminephosphorylceramide; EPT, CDP-ethanolamine:diacylglycerol phosphotransferase; ER, endoplasmic reticulum; ET, CTP:phosphoethanolamine cytidylyltransferase; GPI, glycosylphosphatidylinositol; IPC, inositolphosphorylceramide; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PGP, phosphatidylglycerophosphate; PI, phosphatidylinositol; PS, phosphatidylserine; RNAi, RNA interference; SM, sphingomyelin. FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 1035 or RNA interference (RNAi)-mediated downregulation of gene expression. In addition, factors required for adaptation and growth of different life cycle forms cannot only be investigated in cell culture, but also in suitable animal models (tsetse flies, rodents), allow- ing host–pathogen interactions to be studied (reviewed in [6]). Furthermore, in vitro differentiation of T. brucei from bloodstream- to procyclic (insect)-form parasites may reveal changes in gene expression and metabolism that are essential for the parasite life cycle (reviewed in [7]). Interestingly, T. brucei and other flagellates of the order Kinetoplastida contain single (e.g. mitochondria) and unique (e.g. glycosomes) organelles that undergo dramatic functional and morphological changes during differentiation, making T. brucei an interesting model organism to study organelle biogenesis and turnover (reviewed in [8,9]), and cell division (reviewed in [10]). It has long been known that T. brucei bloodstream forms acquire lipids from their mammalian hosts. For this reason, the study of lipid biosynthesis in trypano- somes has received little attention in the past. Only recently, a series of reports demonstrated that both bloodstream- and insect (procyclic)-form parasites are capable of de novo synthesis of lipids (recently reviewed in [11]). Identification of eukaryotic routes for lipid biosynthesis and of novel, parasite-typical pathways raised a new interest in T. brucei as a model organism to study eukaryotic lipid homeostasis. This review provides an overview of the biosynthetic path- ways for phospholipid synthesis in T. brucei and high- lights differences and unique features that may make trypanosomes an attractive model micro-organism to study lipid turnover and lipid–protein interactions in eukaryotes. Biosynthesis of phosphatidylcholine (PC) PC represents the most abundant glycerophospholipid class in most eukaryotes (reviewed in [12,13]). In all mammalian cells capable of de novo synthesis of phos- pholipids, PC is generated by the CDP-choline path- way, often referred to as the CDP-choline branch of the Kennedy pathway [14]. It involves the sequential action of three enzymes to generate PC from its pre- cursors, choline and diradylglycerol, via the high- energy intermediate CDP-choline. Although in most mammalian cells this pathway is responsible for the production of almost the entire pool of PC (reviewed in [15]), human liver cells synthesize approximately one-third of their PC via sequential methylation of phosphatidylethanolamine (PE) [16], a reaction cata- lysed by PE N-methyltransferase [17]. Both PC biosynthetic pathways are involved in the regulation of lipoprotein metabolism in mice [18,19]. However, they generate distinct pools of PC consisting of different molecular species [20]. A similar observation has also been reported in the yeast, Saccharomyces cerevisiae [21]. Very recently, the lack of PE N-methyltransferase was shown to protect mice against diet-induced obesity and insulin resistance [22], suggesting that this path- way may be linked to the regulation of body energy metabolism. All three enzymes involved in PC formation via the CDP-choline branch of the Kennedy pathway have been identified and characterized in mammalian cells. The cytosolic enzyme choline kinase catalyses the first step in the reaction sequence, phosphorylating choline in an ATP-dependent reaction to phosphocholine. Choline kinases are ubiquitously distributed among eukaryotes [23] and, in general, use both choline and ethanolamine as substrates (reviewed in [24]). In mam- malian cells, choline kinase exists as three different isoforms encoded by two separate genes. Recent studies suggest an important role for choline kinase in cancer cell proliferation (reviewed in [25]). The second enzyme in the CDP-choline pathway, CTP:phospho- choline cytidylyltransferase (CT), uses phosphocholine and CTP as substrates to form CDP-choline, thereby releasing pyrophosphate. In mammalian cells, several isoforms of the enzyme have been described (reviewed in [15,26]), consisting of up to four distinct conserved domains [23,27]. Upon stimulation by lipids, CT is converted from a soluble to a membrane-bound form (reviewed in [28]). In many cells, CT has been localized to the nucleus, but cytosolic forms of the enzyme have also been reported [15]. The reaction catalysed by CT is considered the rate-limiting step in PC synthesis. In the final step of the CDP-choline pathway, a choline phosphotransferase activity transfers phosphocholine from CDP-choline to diradylglycerol to yield PC, releasing CMP as by-product. Two different enzymes catalysing this reaction were identified and character- ized in mammalian cells, a CDP-choline ⁄ ethanol- amine:diacylglycerol phosphotransferase (CEPT) that uses both CDP-choline and CDP-ethanolamine as sub- strates [29] and a CDP-choline:diacylglycerol phospho- transferase (CPT) that uses CDP-choline only as the substrate [30]. Both CEPT and CPT are predicted to be integral membrane proteins and have been reported to localize to the endoplasmic reticulum (ER) ⁄ nuclear membrane and Golgi, respectively [31]. CEPT and CPT activities have also been identified in S. cerevisiae [32,33]. In T. brucei, candidate genes encoding all enzymes of the CDP-choline pathway have been identified Phospholipid biosynthesis in T. brucei M. Serricchio and P. Bu ¨ tikofer 1036 FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS (reviewed in [11]). Choline kinase, which in contrast to mammalian cells is encoded by a single gene in T. brucei [23], has been characterized experimentally in bloodstream-form trypanosomes and displays dual specificity for choline and ethanolamine, with choline being the preferred substrate [34], thus reflecting the situation in most mammalian cells [26]. The second enzyme of the CDP-choline pathway, CT, has not been studied experimentally in T. brucei. A recent report suggests that part of the substrate for CT, phospho- choline, may derive from sphingomyelin (SM) degrada- tion by neutral sphingomyelinase [35]. Based on the importance of the first two enzymes of the CDP-cho- line branch of the Kennedy pathway in mammalian cells and yeast, they are probably essential in T. brucei, but experimental evidence is lacking. Interestingly, all kinetoplastid CTs are unusual fusion proteins in hav- ing a cytidylyltransferase domain fused to a CDP-alco- hol phosphatidyltransferase domain that is normally found in CEPT and CDP-ethanolamine:diacylglycerol phosphotransferase (EPT) [23]. The function of this additional domain is unknown. The third enzyme, CEPT, has been characterized in T. brucei procyclic forms and is involved in the synthesis of both PC and PE. Ablation of CEPT activity using RNAi caused a reduction in PC and PE levels and a growth arrest of parasites in culture [36] (Fig. 1). Subsequent flow cytometry and cytology studies demonstrated that knocking-down CEPT expression inhibits nuclear divi- sion [37], suggesting, for the first time in a eukaryotic organism, a role for CEPT in the control of cell divi- sion. Although at present there is no information available on the localization of CEPT in trypano- somes, its involvement in nuclear division suggests that it may associate with the nuclear envelope membrane in T. brucei parasites, which would be consistent with its localization in mammalian cells [31]. In contrast to mammalian cells and yeast, the T. brucei genome lacks a candidate gene for PE N-methyltransferase. Accordingly, experiments in both bloodstream [38] and procyclic forms [36] have shown that methylation of PE to PC does not occur in T. brucei. Biosynthesis of PE and phosphatidylserine (PS) PE generally represents the second major glycero- phospholipid class in eukaryotes, whereas PS occurs in small amounts only (reviewed in [13,39]). Apart from being a major structural component of eukaryotic and prokaryotic membranes, PE has been shown to affect protein folding [40] and promote membrane fusion and fission events [41]. In addition, PE can serve as a mem- brane anchor for proteins [42], and represents the donor of the ethanolamine moiety for GPI anchor bio- synthesis [43] and the ethanolamine phosphoglycerol modification of eukaryotic elongation factor 1A [44]. It is worth mentioning that the dependence of GPI and ethanolamine phosphoglycerol synthesis on PE as the ethanolamine donor was demonstrated using T. brucei as the model organism, although these pro- tein modifications had been reported for the first time in other eukaryotic organisms. AB C Fig. 1. PE and PC formation in T. brucei. (A) In T. brucei, EPT catalyses the final reaction in alk-1-enyl-acyl PE formation by the Kennedy pathway, whereas the dual-specificity enzyme, CEPT, generates diacyl PE and PC. (B) RNAi-mediated ablation of EPT results in a reduction in alk-1-enyl-acyl PE species and an accumulation of diacyl PE and PC. (C) RNAi-mediated knockdown of CEPT results in a reduction in PC and diacyl PE species and a small increase in alk-1-enyl-acyl PE. Changes in the PE and PC contents in (B) and (C) relative to control cells (A) are reflected by the sizes of the circles and the numbers. The morphological and biochemical changes caused by RNAi are indicated at the bottom. M. Serricchio and P. Bu ¨ tikofer Phospholipid biosynthesis in T. brucei FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 1037 The biosynthesis and turnover of the two amino- phospholipids PS and PE is metabolically closely inter- related. In mammalian cells, PS is synthesized via head group exchange with an existing phospholipid, i.e. by replacing the choline group of PC or the ethanolamine group of PE with the amino acid, l-serine. The reac- tions are catalysed by two distinct activities, PS syn- thase-1 and PS synthase-2, showing different substrate specificities for PC and PE, respectively [45,46]. Both enzymes are localized to mitochondria-associated membranes, i.e. special subdomains of the ER that transiently come in contact with mitochondrial outer membranes [47]. However, the different tissue distribu- tion of the two enzymes suggests that they may have different functions (reviewed in [39]). Knockout mice for PS synthase-1 or PS synthase-2 are viable and exhi- bit minor phenotypes only [48–50], indicating that the two enzymes may have complementary functions in the maintenance of PS homeostasis. In contrast, S. cerevisiae generates PS from CDP- diacylglycerol and l-serine by the action of PS synthe- tase [51], a membrane protein localizing to a special subfraction of microsomes [52]. A similar reaction involving a membrane-associated enzyme also occurs in Gram-positive bacteria [53]. However, in Gram-neg- ative bacteria, PS is synthesized by a cytosolic enzyme, which only associates with membranes upon interac- tion with lipid substrates [54], suggesting that Gram- positive and Gram-negative enzymes evolved from different origins (reviewed in [55]). Whereas PS synthe- tase from the Gram-positive bacterium Bacillus subtilis shows 35% overall amino acid sequence homology to S. cerevisiae PS synthetase, with particularly high homology in the conserved CDP-alcohol phosphatidyl- transferase domain, it shows little homology to PS syn- thetase from Escherichia coli [56]. Interestingly, conserved amino acid motifs in E. coli PS synthetase indicate that it belongs to a large superfamily of pro- teins that includes PS synthetases of other Gram-nega- tive bacteria, bacterial cardiolipin (CL) synthases, phospholipases D, nucleases and pox envelope proteins [57]. PS is not only a membrane component and mediates important cellular functions (reviewed in [58]), but also serves as the substrate for PE formation. In most bac- teria, conversion of PS to PE, a reaction catalysed by PS decarboxylase, represents the only pathway for PE synthesis (reviewed in [59]). Similarly, in yeast and many mammalian cells, decarboxylation of PS is a major pathway for PE formation (reviewed in [39,60,61]). Eukaryotic PS decarboxylases belong to two distinct classes of enzyme that localize to different intracellular compartments. Type I PS decarboxylases are found in mitochondria, whereas type II enzymes localize to the endomembrane system. Typically, PS decarboxylases are membrane proteins consisting of two nonidentical subunits that are generated from sin- gle proenzymes. The contribution of PS decarboxyl- ation to cellular PE formation varies between different cell types or organisms. A PS decarboxylase knockout in mice results in mitochondrial defects and lethality between days 8 and 10 of embryonic development [62]. In eukaryotes, PE can also be synthesized via the CDP-ethanolamine branch of the Kennedy pathway [14], by head group exchange with PS, or by acylation of lyso-PE (reviewed in [58]). The contributions of the latter two pathways to de novo synthesis of PE are unclear. The first reaction of the CDP-ethanolamine branch of the Kennedy pathway is catalysed by etha- nolamine kinase, resulting in the formation of phosphoethanolamine, which in turn is activated using CTP by CTP:phosphoethanolamine cytidylyltransfer- ase (ET) to form CDP-ethanolamine. Alternatively, phosphoethanolamine may also be produced via degra- dation of sphingosine-1-phosphate by sphingosine-1- phosphate lyase [63]. The contribution of this reaction to de novo PE formation in mammalian cells has not been firmly established. The final step in PE formation by the Kennedy pathway is catalysed by the dual-spec- ificity enzyme CEPT, transferring the ethanolamine moiety to diradylglycerol. Interestingly, it has long been thought that CEPT provides all of the ethanol- amine phosphotransferase activity for PE formation. However, recently, a human cDNA encoding a CDP- ethanolamine-specific phosphotransferase (EPT) was isolated and its transcripts were found ubiquitously expressed in multiple tissues [64]. The contribution of the PS decarboxylation reaction and the CDP-ethanolamine branch of the Kennedy pathway to PE formation in mammalian cells has been experimentally addressed using pathway-specific stable isotope labelling experiments, revealing a preferential use of the CDP-ethanolamine pathway over PS decar- boxylation in a ratio of approximately 2 : 1 [65]. In addition, the two pathways were found to generate distinct PE molecular species, with the PS decarboxyl- ation route having a preference for long-chain, polyun- saturated molecular species. Deletion of the ET gene in mice causes embryonic lethality, indicating that PE levels cannot be maintained by PS decarboxylation [66]. In T. brucei, de novo synthesis of PE occurs exclu- sively via the CDP-ethanolamine branch of the Ken- nedy pathway [36]. All enzymes have been identified and experimentally validated [36,38,44]. Disruption of the pathway by downregulation of any of the three Phospholipid biosynthesis in T. brucei M. Serricchio and P. Bu ¨ tikofer 1038 FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS enzymes using RNAi results in growth arrest of the parasites. To our knowledge, T. brucei represented the first eukaryotic organism in which the PE branch of the Kennedy pathway was shown to be essential for cell growth. Only very recently, the essential nat- ure of the Kennedy pathway was also demonstrated in Plasmodium berghei blood stage parasites [67]. Analysis of the phospholipid composition of T. brucei parasites after RNAi against ethanolamine kinase or ET showed alterations not only in PE but also in PC and PS levels [36,37]. In addition, inhibition of PE synthesis also blocked de novo synthesis of GPI anchors and prevented ethanolamine phosphoglycerol addition to eukaryotic elongation factor 1A [44], demonstrating the above-mentioned precursor–prod- uct relationship between PE and ethanolamine-con- taining protein modifications. Furthermore, ablation of ET activity resulted in disruption of mitochondrial morphology and ultrastructure [37], demonstrating for the first time a direct effect of reduced PE levels on mitochondrial integrity. Interestingly, a similar observation has recently been made in mitochondria of mammalian cells. Preliminary work showed that a reduction in mitochondrial PE levels, after depletion of PS decarboxylase capacity, caused alterations in mitochondrial morphology and motility (J. E. Vance, personal communication), suggesting that the effects seen in T. brucei may represent a widespread phe- nomenon. Remarkably, T. brucei PE consists of high levels of ether-type molecular species [36,68,69]. RNAi against EPT and CEPT demonstrated that bulk alk-1-enyl-acyl PE is synthesized by EPT, whereas diacyl-type PE is primarily produced by CEPT [36] (Fig. 1). It is tempt- ing to speculate that the two enzymes may be involved in generating two spatially and functionally distinct pools of PE in T. brucei. The contribution of phosphoethanolamine generated via sphingosine-1-phosphate degradation to PE forma- tion in T. brucei has not been determined. However, in Leishmania parasites, this pathway was shown to be essential if exogenous ethanolamine as the substrate for the Kennedy pathway was absent from the culture medium [70]. The pathway for PS formation in T. brucei has not been firmly established. At present, it is unclear if PS is synthesized from CDP-diacylglycerol and l-serine by PS synthetase [38], or by head group exchange with PE involving PS synthase-2 [37]. Preliminary findings in our laboratory using RNAi against a candidate gene encoding PS synthase-2 indicate that PS formation is essential for parasite viability (J. Jelk & P. Bu ¨ tikofer, unpublished data). Biosynthesis of phosphatidylglycerol (PG) and CL PG and CL represent minor glycerophospholipid classes in eukaryotes. CL is predominantly found in the inner mitochondrial membrane [52] or at contact sites of inner and outer mitochondrial membranes [71]. CL is rather unique in that it has a dimeric structure, consist- ing of two phosphatidyl moieties attached to glycerol and a small negatively charged head group, providing distinct physicochemical properties to CL and CL- containing membranes (reviewed in [72]). Among the many roles of CL, it has been shown to be required for proper function of key mitochondrial enzymes and proteins involved in ATP production via oxidative phosphorylation, as well as for mitochondrial transport systems (reviewed in [73–75]). In addition, CL organizes into membrane domains and participates in the formation and maintenance of dynamic protein–lipid and protein–protein interactions (reviewed in [76]). Remarkably, a reduction in CL levels and changes in the fatty acyl composition of CL have been linked to human diseases, such as Barth syndrome, an X-linked recessive human disorder caused by a defect in the enzyme tafazzin, which is involved in CL acyl chain remodelling in mammalian cells (reviewed in [77]). In contrast to their low abundance in eukaryotic cells, PG and CL represent the major anionic glycero- phospholipid classes in most Gram-positive and Gram- negative bacteria, accounting for  20 and 5% of total phospholipids, respectively [78]. In both prokaryotes and eukaryotes, CL and its biosynthetic precursor, PG, are synthesized from phosphatidic acid (reviewed in [79]). Phosphatidic acid is first activated with CTP to CDP-diacylglycerol by the enzyme CDP-diacylglyc- erol synthase. Following condensation with glycerol-3- phosphate to phosphatidylglycerophosphate (PGP) by PGP synthase, the terminal phosphate group is hydro- lysed to form PG. Interestingly, although bacterial enzymes catalysing PGP dephosphorylation were reported almost 30 years ago [80], the first eukaryotic PGP phosphatase has only recently been identified in S. cerevisiae [81]. The final biosynthetic step in CL formation, catalysed by CL synthase, differs between prokaryotes and eukaryotes. In prokaryotes, PG and a phosphatidyl moiety from another PG are condensed to CL, whereas in eukaryotes, PG and CDP-diacyl- glycerol are fused to CL (reviewed in [79]). PGP synthase and CL synthase each belong to two distinct protein families. The CDP-alcohol phosphat- idyltransferase family includes phosphatidyl- and phos- photransferases acting on CDP-alcohols, whereas the phospholipase D fami ly contains phosphatidyltransferases M. Serricchio and P. Bu ¨ tikofer Phospholipid biosynthesis in T. brucei FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 1039 having active sites related to those found in phospholi- pase D [57]. Bacteria commonly use CDP-alcohol phosphatidyltransferases for the PGP synthase reac- tion, whereas mammals and yeast have phospholipase D-like PGP synthases [23]. Conversely, almost all prokaryotes use phospholipase D-like enzymes for the CL synthase reaction, whereas eukaryotic CL synthas- es belong to the CDP-alcohol phosphatidyltransferase family (reviewed in [23,79]). Experimental evidence for the presence of CL and PG in both T. brucei bloodstream and procyclic forms has been reported [68,69]. However, the pathway for CL synthesis has not been elucidated. Recently, candi- date genes encoding enzymes for all steps in CL syn- thesis have been identified using bioinformatic tools [11]. Preliminary results indicate that PG and CL synthesis in T. brucei is essential for parasite growth (M. Serricchio & P. Bu ¨ tikofer, unpublished data). Biosynthesis of phosphatidylinositol (PI) and GPI PI is a glycerophospholipid class containing an inositol head group derived from the polyol, myo-inositol. PI or derivatives thereof are found in all eukaryotes, including fungi and protozoa, but also in archaea and some pathogenic bacteria [82,83]. In eukaryotes, PI constitutes 3–20% of cellular phospholipids [13]. Apart from being a structural membrane component, PI and its phosphorylated forms also serve as precursors for cell signalling molecules [84] and the biosynthesis of GPI anchors (reviewed in [85,86]). Intracellular myo-inositol can be generated de novo in a two-step reaction process involving inositol-3-phos- phate synthase, generating inositol-3-phosphate from glucose-6-phosphate, and inositol monophosphatase, catalysing dephosphorylation of inositol-3-phosphate to inositol (reviewed in [84]). Alternatively, myo-inositol can be taken up from the environment by inositol trans- porters (reviewed in [87]). Subsequently, myo-inositol is transferred to CDP-diacylglycerol by the action of PI synthase, an enzyme that is conserved in all eukaryotes [88]. In T. brucei, the pathway for myo -inositol synthesis and PI formation has been established. It has been proposed that T. brucei bloodstream forms contain two pools of PI synthase [89,90]. One pool localizes to the ER and uses myo-inositol generated de novo from glucose-6-phosphate whereas the other pool associates with the Golgi and uses myo-inositol taken up from the environment (Fig. 2). In T. brucei bloodstream forms, de novo formation of myo-inositol is essential [90]. In addition, recent results indicate that in both procyclic- and bloodstream-form trypanosomes, uptake of myo-inositol via a specific transporter is nec- essary for normal growth (A. Gonzalez Salgado & P. Bu ¨ tikofer, unpublished data). Interestingly, the PI pool formed from endogenously produced myo-inositol is primarily used for GPI synthesis, whereas exogenous myo-inositol is used for bulk PI formation [89,90]. This two-pool model is consistent with a previous report showing the presence of a subset of distinct PI molecu- lar species that is used for GPI anchor biosynthesis [91]. However, it does not explain why cytosolic myo- inositol produced from glucose-6-phosphate does not (freely) exchange with myo-inositol taken up from the medium, unless de novo-synthesized myo-inositol or its precursor, myo-inositol-3-phosphate, are sequestered from exogenous myo-inositol, as has been suggested [90]. PI synthase was found to be essential in both bloodstream- [89] and procyclic-form trypanosomes (M. Serricchio & P. Bu ¨ tikofer, unpublished data). Although several candidate genes encoding PI kinases have been identified in T. brucei [11], the reactions leading to the production of phosphorylated PIs have not been examined experimentally. In contrast, the involvement of PI as a precursor for GPI anchor synthe- sis has been extensively studied in both bloodstream- and procyclic-form T. brucei (reviewed in [86]). In fact, it was in T. brucei where, for the first time, the entire GPI biosynthetic pathway leading to the formation of the GPI core precursor, ethanolamine-phosphate- Manal-2Manal-6Manal-GlcN-PI [92,93], and the first Fig. 2. Biosynthesis of inositol-containing lipids in T. brucei.In T. brucei bloodstream forms, two pools of PI synthase have been reported, one localizing to the ER and one to the Golgi [89]. The corresponding model proposes that the ER enzyme preferentially uses inositol formed de novo from glucose-6-phosphate to generate PI for GPI anchor synthesis, whereas the Golgi enzyme primarily uses inositol taken up from the environment via a putative myo-inositol transporter (MIT) for PI and IPC synthesis. It is not clear how the two pools of myo-inositol in the cytosol are seques- tered, or exchange with each other. Phospholipid biosynthesis in T. brucei M. Serricchio and P. Bu ¨ tikofer 1040 FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS complete chemical structure of a GPI anchor [2], were established. In addition, T. brucei was the first organism in which remodelling of the acyl chain composition of PI was demonstrated. During GPI anchor synthesis and after GPI attachment to protein in bloodstream-form parasites, the PI acyl chains are replaced by myristate [94,95]. More recent data demonstrate that GPI lipid remodelling also occurs in procyclic-form T. congolense and, possibly, T. brucei [96]. Following the discovery in T. brucei, remodelling of GPIs was also reported in many other organisms (reviewed in [97,98]), indicating that this process probably represents a general event during GPI anchoring of proteins. Biosynthesis of sphingophospholipids The sphingophospholipids, consisting of SM, ethanol- aminephosphorylceramide (EPC) and inositolphos- phorylceramide (IPC), represent key structural components of virtually all eukaryotic membranes. In addition, they represent reservoirs for important sig- nalling molecules, such as sphingosine, sphingosine-1- phosphate and ceramide (reviewed in [99,100]). In most mammalian cells, SM is by far the most abundant sphingophospholipid class (reviewed in [13]), whereas EPC, which represents the major sphingophospholipid in Drosophila melanogaster [101], only occurs in trace amounts [102]. IPCs, or derivatives thereof, have not been detected in mammalian cells, instead they repre- sent prominent sphingophospholipid classes in fungi, plants and protozoa [103–107]. The biosynthesis of sphingo(phospho)lipids starts with the condensation of l-serine with palmitoyl-CoA to form 3-ketosphinganine, a reaction catalysed by serine palmitoyltransferase. After reduction of the product, dihydrosphinganine is N-acylated to dihydro- ceramide by a family of ceramide synthases, with its members showing distinct substrate specificities for fatty acyl-CoAs [108,109]. Dihydroceramide is subse- quently desaturated to ceramide, the central metabolite in sphingolipid metabolism and branch point for the synthesis of SM, EPC and IPC. The formation of SM is catalysed by SM synthase and involves transfer of phosphocholine from PC to ceramide to generate SM and diradylglycerol. In mam- malian cells, two SM synthases have been identified, one located in the lumen of the Golgi and the other in the plasma membrane [110]. In addition, mammalian cells express an SM synthase-related protein, SMSr, that was recently shown to have EPC synthase activity and is localized in the ER [111,112], confirming earlier reports on the presence of such an activity in rat liver and brain microsomes [102,113]. The distinct localization of the SM synthases presumably reflects their roles in de novo SM synthesis (Golgi enzyme) and regeneration of SM from ceramide (plasma membrane enzyme). IPC synthase is an essential enzyme in fungi [114,115] and localizes to the Golgi [116]. Recently, its function and localization in yeast was shown to be dependent on the expression of an additional protein, Kei1, suggesting that IPC synthase may consist of multiple subunits [117]. In T. brucei, candidate genes for all enzymes involved in ceramide synthesis have been identified [11]. However, with the exception of serine palmitoyl- transferase [118,119], individual enzymes have not been studied experimentally. In contrast, the subsequent steps in SM, IPC and EPC formation in T. brucei have recently been characterized in detail [107,120]. The reactions are catalysed by a family of sphingolipid syn- thases, TbSLS1-4, showing distinct substrate specifici- ties. Using a cell-free synthesis system for the expression of polytopic membrane proteins [121], TbSLS1 was identified as IPC synthase, TbSLS2 as EPC synthase, whereas TbSLS3 and TbSLS4 show dual specificities for PC and PE as head group donors to produce SM and EPC, respectively [120]. Interest- ingly, the production of the different sphingophospho- lipid classes is developmentally regulated, with IPC being produced preferentially in T. brucei procyclic forms and EPC in bloodstream forms, whereas SM is generated in both life cycle forms [107,120] (Fig. 3). The localization of T. brucei sphingolipid synthases has not been reported. Whether sphingophospholipids are involved in protein trafficking to the cell surface in T. brucei bloodstream forms is unclear [35,119]. To our knowledge, T. brucei represents the only organism so far that synthesizes all three sphingophospholipid classes, SM, IPC and EPC, and thus represents an ideal model organism to study their biosynthesis, regulation and functional roles. Fig. 3. Sphingophospholipid formation in T. brucei.InT. brucei, all three classes of sphingophospholipids, EPC, IPC and SM, are gen- erated. A family of sphingolipid synthases (TbSLS1–4) is responsi- ble for the stage-specific production of the different classes in bloodstream- (BSF) and procyclic-form (PCF) parasites. The horizon- tal line indicates that the lipid class is present in trace amounts only. M. Serricchio and P. Bu ¨ tikofer Phospholipid biosynthesis in T. brucei FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 1041 Acknowledgements Work in PB’s laboratory is supported by Swiss National Science Foundation grant 31003A_130815 and Sinergia grant CRSII3_127300. We thank J.E. Vance for communicating unpublished findings, and I. Roditi and J.D. Bangs for comments on parts of the manuscript. PB thanks S. Harley and M. Bu ¨ tikofer for stimulation and input. MS thanks K. Durrer for support and advice. References 1 Blum B, Bakalara N & Simpson L (1990) A model for RNA editing in kinetoplastid mitochondria: ‘‘guide’’ RNA molecules transcribed from maxicircle DNA provide the edited information. Cell 60, 189– 198. 2 Ferguson MA, Homans SW, Dwek RA & Rademacher TW (1988) The glycosylphosphatidylinositol membrane anchor of Trypanosoma brucei variant surface glycoprotein. Biochem Soc Trans 16, 265–268. 3 Sutton RE & Boothroyd JC (1986) Evidence for trans splicing in trypanosomes. Cell 47, 527–535. 4 Cross GA (1977) Antigenic variation in trypanosomes. Am J Trop Med Hyg 26, 240–244. 5 Berriman M, Ghedin E, Hertz-Fowler C, Blandin G, Renauld H, Bartholomeu DC, Lennard NJ, Caler E, Hamlin NE, Haas B et al. (2005) The genome of the African trypanosome Trypanosoma brucei. Science 309, 416–422. 6 Roditi I & Lehane MJ (2008) Interactions between trypanosomes and tsetse flies. Curr Opin Microbiol 11, 345–351. 7 Fenn K & Matthews KR (2007) The cell biology of Trypanosoma brucei differentiation. Curr Opin Micro- biol 10, 539–546. 8 Schneider A (2001) Unique aspects of mitochondrial biogenesis in trypanosomatids. Int J Parasitol 31, 1403–1415. 9 He CY (2007) Golgi biogenesis in simple eukaryotes. Cell Microbiol 9 , 566–572. 10 Vaughan S & Gull K (2008) The structural mechanics of cell division in Trypanosoma brucei. Biochem Soc Trans 36, 421–424. 11 Smith TK & Bu ¨ tikofer P (2010) Lipid metabolism in Trypanosoma brucei. Mol Biochem Parasitol 172, 66–79. 12 Li Z & Vance DE (2008) Phosphatidylcholine and cho- line homeostasis. J Lipid Res 49, 1187–1194. 13 van Meer G, Voelker DR & Feigenson GW (2008) Membrane lipids: where they are and how they behave. Nat Rev Mol Cell Biol 9, 112–124. 14 Kennedy EP & Weiss SB (1956) The function of cyti- dine coenzymes in the biosynthesis of phospholipides. J Biol Chem 222, 193–214. 15 Vance JE & Vance DE (2004) Phospholipid biosynthe- sis in mammalian cells. Biochem Cell Biol 82, 113–128. 16 Sundler R & Akesson B (1975) Regulation of phospho- lipid biosynthesis in isolated rat hepatocytes. Effect of different substrates. J Biol Chem 250, 3359–3367. 17 Bremer J & Greenberg DM (1960) Biosynthesis of choline in vitro. Biochim Biophys Acta 37, 173–175. 18 Noga AA, Zhao Y & Vance DE (2002) An unexpected requirement for phosphatidylethanolamine N-methyl- transferase in the secretion of very low density lipopro- teins. J Biol Chem 277, 42358–42365. 19 Robichaud JC, Francis GA & Vance DE (2008) A role for hepatic scavenger receptor class B, type I in decreasing high density lipoprotein levels in mice that lack phosphatidylethanolamine N-methyltransferase. J Biol Chem 283, 35496–35506. 20 DeLong CJ, Shen YJ, Thomas MJ & Cui Z (1999) Molecular distinction of phosphatidylcholine synthesis between the CDP-choline pathway and phosphatidyl- ethanolamine methylation pathway. J Biol Chem 274, 29683–29688. 21 Boumann HA, Damen MJ, Versluis C, Heck AJ, de Kruijff B & de Kroon AI (2003) The two biosynthetic routes leading to phosphatidylcholine in yeast produce different sets of molecular species. Evidence for lipid remodeling. Biochemistry 42, 3054–3059. 22 Jacobs RL, Zhao Y, Koonen DP, Sletten T, Su B, Lingrell S, Cao G, Peake DA, Kuo MS, Proctor SD et al. (2010) Impaired de novo choline synthesis explains why phosphatidylethanolamine N-methyltrans- ferase-deficient mice are protected from diet-induced obesity. J Biol Chem 285, 22403–22413. 23 Lykidis A (2007) Comparative genomics and evolution of eukaryotic phospholipid biosynthesis. Prog Lipid Res 46, 171–199. 24 Aoyama C, Liao H & Ishidate K (2004) Structure and function of choline kinase isoforms in mammalian cells. Prog Lipid Res 43, 266–281. 25 Wu G & Vance DE (2010) Choline kinase and its func- tion. Biochem Cell Biol 88, 559–564. 26 Gibellini F & Smith TK (2010) The Kennedy pathway – de novo synthesis of phosphatidylethanolamine and phosphatidylcholine. IUBMB Life 62 , 414–428. 27 Lykidis A, Baburina I & Jackowski S (1999) Distribu- tion of CTP:phosphocholine cytidylyltransferase (CCT) isoforms. Identification of a new CCTbeta splice vari- ant. J Biol Chem 274, 26992–27001. 28 Cornell RB & Northwood IC (2000) Regulation of CTP:phosphocholine cytidylyltransferase by amphitro- pism and relocalization. Trends Biochem Sci 25, 441– 447. 29 Henneberry AL & McMaster CR (1999) Cloning and expression of a human choline ⁄ ethanolaminephospho- transferase: synthesis of phosphatidylcholine and phosphatidylethanolamine. Biochem J 339, 291–298. Phospholipid biosynthesis in T. brucei M. Serricchio and P. Bu ¨ tikofer 1042 FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 30 Henneberry AL, Wistow G & McMaster CR (2000) Cloning, genomic organization, and characterization of a human cholinephosphotransferase. J Biol Chem 275, 29808–29815. 31 Henneberry AL, Wright MM & McMaster CR (2002) The major sites of cellular phospholipid synthesis and molecular determinants of fatty acid and lipid head group specificity. Mol Biol Cell 13, 3148–3161. 32 Henneberry AL, Lagace TA, Ridgway ND & McMas- ter CR (2001) Phosphatidylcholine synthesis influences the diacylglycerol homeostasis required for SEC14p- dependent Golgi function and cell growth. Mol Biol Cell 12, 511–520. 33 Boumann HA, de Kruijff B, Heck AJ & de Kroon AI (2004) The selective utilization of substrates in vivo by the phosphatidylethanolamine and phosphatidylcho- line biosynthetic enzymes Ept1p and Cpt1p in yeast. FEBS Lett 569, 173–177. 34 Gibellini F, Hunter WN & Smith TK (2008) Biochemi- cal characterization of the initial steps of the Kennedy pathway in Trypanosoma brucei: the ethanolamine and choline kinases. Biochem J 415, 135–144. 35 Young SA & Smith TK (2010) The essential neutral sphingomyelinase is involved in the trafficking of the variant surface glycoprotein in the bloodstream form of Trypanosoma brucei. Mol Microbiol 76, 1461–1482. 36 Signorell A, Rauch M, Jelk J, Ferguson MA & Bu ¨ tikofer P (2008) Phosphatidylethanolamine in Trypanosoma brucei is organized in two separate pools and is synthesized exclusively by the Kennedy pathway. J Biol Chem 283, 23636–23644. 37 Signorell A, Gluenz E, Rettig J, Schneider A, Shaw MK, Gull K & Bu ¨ tikofer P (2009) Perturbation of phosphatidylethanolamine synthesis affects mitochon- drial morphology and cell-cycle progression in procy- clic-form Trypanosoma brucei. Mol Microbiol 72 , 1068– 1079. 38 Gibellini F, Hunter WN & Smith TK (2009) The etha- nolamine branch of the Kennedy pathway is essential in the bloodstream form of Trypanosoma brucei. Mol Microbiol 73, 826–843. 39 Vance JE (2008) Phosphatidylserine and phosphatidyl- ethanolamine in mammalian cells: two metabolically related aminophospholipids. J Lipid Res 49, 1377– 1387. 40 Dowhan W & Bogdanov M (2009) Lipid-dependent membrane protein topogenesis. Annu Rev Biochem 78, 515–540. 41 Emoto K & Umeda M (2000) An essential role for a membrane lipid in cytokinesis. Regulation of contrac- tile ring disassembly by redistribution of phosphatidyl- ethanolamine. J Cell Biol 149, 1215–1224. 42 Ichimura Y, Kirisako T, Takao T, Satomi Y, Shimonishi Y, Ishihara N, Mizushima N, Tanida I, Kominami E, Ohsumi M et al. (2000) A ubiquitin-like system mediates protein lipidation. Nature 408, 488– 492. 43 Menon AK, Eppinger M, Mayor S & Schwarz RT (1993) Phosphatidylethanolamine is the donor of the terminal phosphoethanolamine group in trypanosome glycosylphosphatidylinositols. EMBO J 12 , 1907–1914. 44 Signorell A, Jelk J, Rauch M & Bu ¨ tikofer P (2008) Phosphatidylethanolamine is the precursor of the etha- nolamine phosphoglycerol moiety bound to eukaryotic elongation factor 1A. J Biol Chem 283, 20320–20329. 45 Kuge O, Nishijima M & Akamatsu Y (1985) Isolation of a somatic-cell mutant defective in phosphatidylserine biosynthesis. Proc Natl Acad Sci USA 82, 1926–1930. 46 Voelker DR & Frazier JL (1986) Isolation and charac- terization of a Chinese hamster ovary cell line requiring ethanolamine or phosphatidylserine for growth and exhibiting defective phosphatidylserine synthase activ- ity. J Biol Chem 261, 1002–1008. 47 Stone SJ & Vance JE (2000) Phosphatidylserine syn- thase-1 and -2 are localized to mitochondria-associated membranes. J Biol Chem 275, 34534–34540. 48 Bergo MO, Gavino BJ, Steenbergen R, Sturbois B, Parlow AF, Sanan DA, Skarnes WC, Vance JE & Young SG (2002) Defining the importance of phos- phatidylserine synthase 2 in mice. J Biol Chem 277, 47701–47708. 49 Steenbergen R, Nanowski TS, Nelson R, Young SG & Vance JE (2006) Phospholipid homeostasis in phospha- tidylserine synthase-2-deficient mice. Biochim Biophys Acta 1761, 313–323. 50 Arikketh D, Nelson R & Vance JE (2008) Defining the importance of phosphatidylserine synthase-1 (PSS1): unexpected viability of PSS1-deficient mice. J Biol Chem 283, 12888–12897. 51 Letts VA, Klig LS, Bae-Lee M, Carman GM & Henry SA (1983) Isolation of the yeast structural gene for the membrane-associated enzyme phosphatidylserine syn- thase. Proc Natl Acad Sci USA 80, 7279–7283. 52 Zinser E, Sperka-Gottlieb CD, Fasch EV, Kohlwein SD, Paltauf F & Daum G (1991) Phospholipid synthe- sis and lipid composition of subcellular membranes in the unicellular eukaryote Saccharomyces cerevisiae. J Bacteriol 173, 2026–2034. 53 Kanfer JN & Kennedy EP (1962) Synthesis of phos- phatidylserine by Escherichia coli. J Biol Chem 237, PC270–PC271. 54 Louie K, Chen YC & Dowhan W (1986) Substrate- induced membrane association of phosphatidylserine synthase from Escherichia coli. J Bacteriol 165, 805–812. 55 Matsumoto K (1997) Phosphatidylserine synthase from bacteria. Biochim Biophys Acta 1348, 214–227. 56 Okada M, Matsuzaki H, Shibuya I & Matsumoto K (1994) Cloning, sequencing, and expression in Escheri- chia coli of the Bacillus subtilis gene for phosphatidyl- serine synthase. J Bacteriol 176, 7456–7461. M. Serricchio and P. Bu ¨ tikofer Phospholipid biosynthesis in T. brucei FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS 1043 57 Koonin EV (1996) A duplicated catalytic motif in a new superfamily of phosphohydrolases and phospholipid synthases that includes poxvirus envelope proteins. Trends Biochem Sci 21, 242–243. 58 Vance JE & Steenbergen R (2005) Metabolism and functions of phosphatidylserine. Prog Lipid Res 44, 207–234. 59 Raetz CR & Dowhan W (1990) Biosynthesis and func- tion of phospholipids in Escherichia coli. J Biol Chem 265, 1235–1238. 60 Choi JY, Wu WI & Voelker DR (2005) Phosphatidyl- serine decarboxylases as genetic and biochemical tools for studying phospholipid traffic. Anal Biochem 347, 165–175. 61 Schuiki I & Daum G (2009) Phosphatidylserine decar- boxylases, key enzymes of lipid metabolism. IUBMB Life 61, 151–162. 62 Steenbergen R, Nanowski TS, Beigneux A, Kulinski A, Young SG & Vance JE (2005) Disruption of the phos- phatidylserine decarboxylase gene in mice causes embryonic lethality and mitochondrial defects. J Biol Chem 280, 40032–40040. 63 Zhou J & Saba JD (1998) Identification of the first mammalian sphingosine phosphate lyase gene and its functional expression in yeast. Biochem Biophys Res Commun 242, 502–507. 64 Horibata Y & Hirabayashi Y (2007) Identification and characterization of human ethanolaminephosphotrans- ferase1. J Lipid Res 48, 503–508. 65 Bleijerveld OB, Brouwers JF, Vaandrager AB, Helms JB & Houweling M (2007) The CDP-ethanolamine pathway and phosphatidylserine decarboxylation generate different phosphatidylethanolamine molecular species. J Biol Chem 282, 28362–28372. 66 Fullerton MD, Hakimuddin F & Bakovic M (2007) Developmental and metabolic effects of disruption of the mouse CTP:phosphoethanolamine cytidylyltransfer- ase gene (Pcyt2). Mol Cell Biol 27, 3327–3336. 67 Dechamps S, Wengelnik K, Berry-Sterkers L, Cerdan R, Vial HJ & Gannoun-Zaki L (2010) The Kennedy phospholipid biosynthesis pathways are refractory to genetic disruption in Plasmodium berghei and therefore appear essential in blood stages. Mol Biochem Parasitol 173, 69–80. 68 Patnaik PK, Field MC, Menon AK, Cross GA, Yee MC & Bu ¨ tikofer P (1993) Molecular species analysis of phospholipids from Trypanosoma brucei bloodstream and procyclic forms. Mol Biochem Parasitol 58, 97–105. 69 Richmond GS, Gibellini F, Young SA, Major L, Den- ton H, Lilley A & Smith TK (2010) Lipidomic analysis of bloodstream and procyclic form Trypanosoma bru- cei. Parasitology 137, 1357–1392. 70 Zhang K, Pompey JM, Hsu FF, Key P, Bandhuvula P, Saba JD, Turk J & Beverley SM (2007) Redirection of sphingolipid metabolism toward de novo synthesis of ethanolamine in Leishmania. EMBO J 26, 1094–1104. 71 Ardail D, Privat JP, Egret-Charlier M, Levrat C, Lerme F & Louisot P (1990) Mitochondrial contact sites. Lipid composition and dynamics. J Biol Chem 265, 18797–18802. 72 Lewis RN & McElhaney RN (2009) The physicochemi- cal properties of cardiolipin bilayers and cardiolipin- containing lipid membranes. Biochim Biophys Acta 1788, 2069–2079. 73 Klingenberg M (2009) Cardiolipin and mitochondrial carriers. Biochim Biophys Acta 1788, 2048–2058. 74 Schlame M & Ren M (2009) The role of cardiolipin in the structural organization of mitochondrial mem- branes. Biochim Biophys Acta 1788, 2080–2083. 75 Bogdanov M, Mileykovskaya E & Dowhan W (2008) Lipids in the assembly of membrane proteins and orga- nization of protein supercomplexes: implications for lipid-linked disorders. Subcell Biochem 49, 197–239. 76 Mileykovskaya E & Dowhan W (2009) Cardiolipin membrane domains in prokaryotes and eukaryotes. Biochim Biophys Acta 1788, 2084–2091. 77 Houtkooper RH & Vaz FM (2008) Cardiolipin, the heart of mitochondrial metabolism. Cell Mol Life Sci 65, 2493–2506. 78 Cronan JE (2003) Bacterial membrane lipids: where do we stand? Annu Rev Microbiol 57, 203–224. 79 Schlame M (2008) Cardiolipin synthesis for the assem- bly of bacterial and mitochondrial membranes. J Lipid Res 49, 1607–1620. 80 Icho T & Raetz CR (1983) Multiple genes for membrane-bound phosphatases in Escherichia coli and their action on phospholipid precursors. J Bacteriol 153, 722–730. 81 Osman C, Haag M, Wieland FT, Brugger B & Langer T (2010) A mitochondrial phosphatase required for cardiolipin biosynthesis: the PGP phosphatase Gep4. EMBO J 29, 1976–1987. 82 Jackson M, Crick DC & Brennan PJ (2000) Phosphati- dylinositol is an essential phospholipid of mycobacte- ria. J Biol Chem 275, 30092–30099. 83 Koga Y & Morii H (2007) Biosynthesis of ether-type polar lipids in archaea and evolutionary considerations. Microbiol Mol Biol Rev 71, 97–120. 84 Michell RH (2008) Inositol derivatives: evolution and functions. Nat Rev Mol Cell Biol 9, 151–161. 85 Orlean P & Menon AK (2007) Thematic review series: lipid posttranslational modifications. GPI anchoring of protein in yeast and mammalian cells, or: how we learned to stop worrying and love glycophospholipids. J Lipid Res 48, 993–1011. 86 Hong Y & Kinoshita T (2009) Trypanosome glycosyl- phosphatidylinositol biosynthesis. Korean J Parasitol 47, 197–204. Phospholipid biosynthesis in T. brucei M. Serricchio and P. Bu ¨ tikofer 1044 FEBS Journal 278 (2011) 1035–1046 ª 2011 The Authors Journal compilation ª 2011 FEBS [...]... Allegood JC, Rawat SS, Edwards MB, Wang X, Merrill AH Jr, Acharya U & Acharya JK (2007) Ceramide transfer protein function is essential for normal oxidative stress response and lifespan Proc Natl Acad Sci USA 104, 11364–11369 Malgat M, Maurice A & Baraud J (1986) Sphingomyelin and ceramide-phosphoethanolamine synthesis by microsomes and plasma membranes from rat liver and brain J Lipid Res 27, 251–260 Smith... Kuroda M, Hashida-Okado T, Yasumoto R, Gomi K, Kato I & Takesako K (1999) An aureobasidin A resistance gene isolated from Aspergillus is a homolog of yeast AUR1, a gene responsible for inositol phosphorylceramide (IPC) synthase activity Mol Gen Genet 261, 290–296 Levine TP, Wiggins CA & Munro S (2000) Inositol phosphorylceramide synthase is located in the Golgi apparatus of Saccharomyces cerevisiae Mol... 117 Sato K, Noda Y & Yoda K (2009) Kei1: a novel subunit of inositolphosphorylceramide synthase, essential for its enzyme activity and Golgi localization Mol Biol Cell 20, 4444–4457 118 Fridberg A, Olson CL, Nakayasu ES, Tyler KM, Almeida IC & Engman DM (2008) Sphingolipid synthesis is necessary for kinetoplast segregation and cytokinesis in Trypanosoma brucei J Cell Sci 121, 522– 535 119 Sutterwala SS,... (1995) Structural analysis of inositol phospholipids from Trypanosoma cruzi epimastigote forms Biochem J 310, 255–261 Markham JE, Li J, Cahoon EB & Jaworski JG (2006) Separation and identification of major plant sphingolipid classes from leaves J Biol Chem 281, 22684– 22694 Sutterwala SS, Hsu FF, Sevova ES, Schwartz KJ, Zhang K, Key P, Turk J, Beverley SM & Bangs JD (2008) Developmentally regulated sphingolipid... Sanyal S, Menon AK & Bangs JD (2007) De novo sphingolipid synthesis is essential for viability, but not for transport of glycosylphosphatidylinositol-anchored proteins, in African trypanosomes Eukaryot Cell 6, 454–464 120 Sevova ES, Goren MA, Schwartz KJ, Hsu FF, Turk J, Fox BG & Bangs JD (2010) Cell-free synthesis and functional characterization of sphingolipid synthases from parasitic trypanosomatid... (1974) Inositol phosphorylceramide, a novel substance and the chief member of a major group of yeast sphingolipids containing a single inositol phosphate J Biol Chem 249, 3395–3405 Kaneshiro ES, Jayasimhulu K & Lester RL (1986) Characterization of inositol lipids from Leishmania donovani promastigotes: identification of an inositol sphingophospholipid J Lipid Res 27, 1294–1303 Bertello LE, Goncalvez MF,... 1970–1977 93 Masterson WJ, Doering TL, Hart GW & Englund PT (1989) A novel pathway for glycan assembly: biosynthesis of the glycosyl-phosphatidylinositol anchor of the trypanosome variant surface glycoprotein Cell 56, 793–800 94 Masterson WJ (1990) Biosynthesis of the glycosyl phosphatidylinositol anchor of Trypanosoma brucei variant surface glycoprotein Biochem Soc Trans 18, 722–724 95 Buxbaum LU, Milne... (1987) Sidedness of ceramide-phosphoethanolamine synthesis on rat liver and brain microsomal membranes J Lipid Res 28, 138–143 Nagiec MM, Nagiec EE, Baltisberger JA, Wells GB, Lester RL & Dickson RC (1997) Sphingolipid synthesis as a target for antifungal drugs Complementation of the inositol phosphorylceramide synthase defect in a mutant strain of Saccharomyces cerevisiae by the AUR1 gene J Biol Chem... Buxbaum LU, Milne KG, Werbovetz KA & Englund PT (1996) Myristate exchange on the Trypanosoma brucei variant surface glycoprotein Proc Natl Acad Sci USA 93, 1178–1183 96 Butikofer P, Greganova E, Liu YC, Edwards IJ, ¨ Lehane MJ & Acosta-Serrano A (2010) Lipid remodelling of glycosylphosphatidylinositol (GPI) glycoconjugates in procyclic-form trypanosomes: biosynthesis and processing of GPIs revisited... Conzelmann A (2007) Multiple functions of inositolphosphorylceramides in the formation and intracellular transport of glycosylphosphatidylinositolanchored proteins in yeast Biochem Soc Symp 74, 199–209 98 Fujita M & Kinoshita T (2010) Structural remodeling of GPI anchors during biosynthesis and after attachment to proteins FEBS Lett 584, 1670–1677 99 Merrill AH Jr, Sullards MC, Wang E, Voss KA & Riley . Switzerland Introduction Trypanosoma brucei is a eukaryotic protozoan parasite causing African sleeping sickness in humans and nagana in domestic animals. During its complex life cycle, it migrates. M, Hashida-Okado T, Yasumoto R, Gomi K, Kato I & Takesako K (1999) An aureobasidin A resis- tance gene isolated from Aspergillus is a homolog of yeast AUR1, a gene responsible for inositol. palmitoyl-CoA to form 3-ketosphinganine, a reaction catalysed by serine palmitoyltransferase. After reduction of the product, dihydrosphinganine is N-acylated to dihydro- ceramide by a family