Biogenesis and cellular dynamics of aminoglycerophospholipids

Biogenesis and Cellular Dynamics of Aminoglycerophospholipids Ruth Birner and GLinther Daum Institut ffir Biochemie, Technische Universit~itGraz, Petersgasse 12/2, A-8010 Graz, Austria

Aminoglycerophospholipids phosphatidylserine (PtdSer), phosphatidylethanolamine (PtdEtn), and phosphatidylcholine (PtdCho) comprise about 80% of total cellular phospholipids in most cell types. While the major function of PtdCho in eukaryotes and PtdEtn in prokaryotes is that of bulk membrane lipids, PtdSer is a minor component and appears to play a more specialized role in the plasma membrane of eukaryotes, e.g., in cell recognition processes. All three aminoglycerophospholipid classes are essential in mammals, whereas prokaryotes and lower eukaryotes such as yeast appear to be more flexible regarding their aminoglycerophospholipid requirement. Since different subcellular compartments of eukaryotes, namely the endoplasmic reticulum and mitochondria, contribute to the biosynthetic sequence of aminoglycerophospholipid formation, intracellular transport, sorting, and specific function of these lipids in different organelles are of special interest. KEY WORDS: Aminoglycerophospholipid, Phosphatidylserine, Phosphatidylethanolamine, Phosphatidylcholine, Lipid biosynthesis, Lipid Transport, Function. ®2003, ElsevierScience (USA).

I. Introduction Glycerophospholipids play multiple roles in cells. First, they are the major components of membrane bilayers providing the permeability barrier between the interior of the cell and its environment and between the lumen of organelles and the cytosol. Membranes, however, are not merely static; they are highly dynamic structures involved in vesicle transport, cell division, or spore formation by

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fusion/fission events. Second, glycerophospholipids not only provide the hydrophobic matrix for membrane proteins, but also specifically interact with enzymes by affecting their assembly and activity. Third, some glycerophospholipids are second messengers involved in recognition processes and signal transduction. Finally, glycerophospholipids are precursors of several macromolecules, such as bacterial lipopolysaccharide (LPS) or eukaryotic glycosylphosphatidylinositol anchor (GPI). The variety of functions of different subcellular membranes may be one reason for phospholipid diversity. While the number of nucleotides or amino acids forming DNA or proteins, respectively, has been highly conserved during the evolution, the number of different lipid molecules has increased during the development of species. Thus, lipid diversity may play a major role in the formation of the highly specified membranes characteristic of highly developed organisms. The major lipids of bacterial membranes are certain glycerophospholipid species, whereas eukaryotic membranes contain in addition sterols, sphlngolipids, and glycolipids. Glycerophospholipids contain a glycerol backbone, a polar head group, which is characteristic for each glycerophospholipid class, and a hydrophobic tail consisting of acyl chains that vary in length and degree of saturation (Fig. 1). The most important glycerophospholipid classes are as follows: phosphatidic acid (PA), phosphatidylserine (PtdSer), phosphatidylethanolamine (PtdEtn), phosphatidylcholine (PtdCho), phosphatidylinositol (PtdIns), pbosphatidylglycerol (PtdGro), and cardiolipin (CL). Lyso-glycerophospholipids contain only one acyl chain either in snl or in sn2 position. Because PtdSer, PtdEtn, and PtdCho share an amino group as a common structural feature in their head group, they are called aminoglycerophospholipids. While PA, PtdSer, PtdIns, PtdGro, and CL are anionic phospholipids, PtdCho and PtdEtn are zwitterionic phospholipids. The head groups of PtdSer and PtdEtn are rather small compared to that of PtdCho. This is an important property because the size of the head group determines the shape of individual phospholipid species and thus affects the structure and dynamics of membranes. Different molecular properties of tipids result in lipid polymorphism (Dowhan, 1997a; de Kruij ff, 1997), bilayer and nonbilayer structures (e.g., inverted hexagonal phase) being most prominent representatives. PtdCho is a typical bilayer lipid because of its cylindrical shape, whereas the conical PtdEtn forms nonbilayer structures, as do PA and CL in the presence of divalent cations. A bilayer-to-nonbilayer phase transition allows the membrane to undergo local rearrangement which is needed for fusion, cell division, or budding processes. The lipid composition of cells varies greatly with growth conditions, with growth phase, and among organelles, implying the existence of specific lipid requirements and sorting mechanisms. For example, the aminoglycerophospholipids PtdCho and PtdEtn are considered major bulk membrane components, whereas CL and PtdGro are restricted to mitochondrial membranes and are important for respiration. As another example, the minor aminoglycerophospholipid PtdSer, together with the majority of sterols, sphingolipids, and GPI-anchored proteins, is found

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in the plasma membrane and appears to be specifically sorted along the secretory pathway. Asymmetric lipid composition of the two leaflets of a membrane bilayer is the rule, not the exception. The best-studied example in that respect is the plasma membrane, with PtdCho being present in both leaflets and PtdSer and PtdEtn exclusively localized to the inner (cytosolic) side. As a consequence of the above-mentioned features, aminoglycerophospholipids are of great importance for cellular structure, function, and maintenance of organelles.

II. Biogenesis and Function of Aminoglycerophospholipids A. Biosynthesis of Phosphatidylserine, Phosphatidylethanolamine, and Phosphatidylcholine Aminoglycerophospholipids PtdSer, PtdEtn, and PtdCho are metabolically related, although different pathways lead to their formation in different species (Fig. 2) (Kent, 1995; Daum et al., 1998; Vance, 1998).

1. Synthesis of Phosphatidylserine In prokaryotes and lower eukaryotes, PtdSer is synthesized from cytidinediphosphate diacylglycerol (CDP-DAG) and serine (Ser) by PtdSer synthase. The precursor for this reaction, CDP-DAG, is synthesized from PA and cytidinetriphosphate (CTP) by CDP-DAG synthase (CDS), which has been identified in E. coli, S. cerevisiae, Drosophila, plants, mammalian cells (Dowhan, 1997b; Heacock and Agranoff, 1997), and Plasmodium falciparum (Martin et aL, 2000). While the yeast CDS1 gene is essential, suggesting that only one CDS exists or Cdslp forms an essential pool of CDP-DAG, two mammalian isoforms of CDS, namely a microsomal CDS1 and a mitochondrial CDS2, have been cloned (Lykidis et al., 1997; Halford et al., 1998; Volta et al., 1999). PtdSer synthase (PSS) has been characterized from several microorganisms, including E. coli (DeChavigny et aL, 1991), Bacillus subtilis (Okada et al., 1994), Helicobacter pylori (Ge and Taylor, 1997), and yeast (Yamashita and Nikawa, 1997). While the E. coli PSS, PssAp, is associated with ribosomes and binds to the membrane through electrostatic force (Louie et al., 1986), the PSS proteins from other species appear to be true membrane components. As an example, the yeast Pssl/Cholp is associated with the endoplasmic reticulum (ER) and related membranes (Kuchler et al., 1986; Gaigg et al., 1995). In mammalian cells, PtdSer synthesis localized to the ER and related membranes occurs in a different reaction. Mammalian PtdSer is synthesized in a Ca 2+-dependent base exchange catalyzed by two PtdSer synthases: PSS 1 primarily uses PtdCho as a substrate for exchange of the head group (Kuge and Nishijima,

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1997), whereas PSS2 converts PtdEtn to PtdSer in vivo (Saito et al., 1998). Both enzymes were cloned from Chinese hamster ovary (CHO) cells (Kuge et al., 1991, 1997) and later from mouse (Stone et al., 1998; Stone and Vance, 1999). Both PSS 1 and PSS2, which are highly hydrophobic, contain multiple potential transmembrane domains and typical ER targeting sequences. Murine PSS 1 mRNA is most abundant in kidney, brain, and liver, while PSS2 mRNA is expressed at the highest rate in testis (Sturbois-Balcerzak et al., 2001). In CHO cells and in resting keratinocytes, PtdCho is the major precursor for PtdSer synthesis (Arthur and Lu, 1993; Stone and Vance, 1999). In plants, synthesis of PtdSer occurs by both PtdSer synthase and serine exchange (Delhaize et al., 1999; Manoharan et al., 2000). The plant serine exchange enzyme accepts PtdEtn and PtdCho as substrates. Neither enzyme, however, has yet been identified at the molecular level. Both enzymatic activities, the PtdSer synthase and the serine exchange enzyme, are thought to be localized to the ER, but an additional serine exchange activity has also been localized to the plant plasma membrane (Vincent et aL, 1999).

2. Synthesis of Phosphatidylethanolamine by Decarboxylation of Phosphatidylserine Biosynthesis of PtdEtn in prokaryotes and eukaryotes is accomplished by decarboxylation of PtdSer (Voelker, 1997). In E. coli and most other bacteria this is the only pathway of PtdEtn synthesis, whereas in eukaryotes PtdEtn is also formed through the CDP-Etn branch of the Kennedy pathway (see Section II.A.4). Bacterial PtdSer decarboxylase is an integral membrane protein, which was purified from E. coli (Dowhan et al., 1974) and cloned from B. subtilis (Matsumoto et al., 1998). In yeast, two different PtdSer decarboxylases were cloned: PtdSer decarboxylase 1, Psdlp (Trotter et al., 1993), which is localized to the inner mitochondrial membrane (Zinser et al., 1991), and Psd2p (Trotter et al., 1995), which is present in a Golgi/vacuolar compartment (Trotter and Voelker, 1995). Mutants defective in one of the PtdSer decarboxylases, Psdlp or Psd2p, grow like wildtype on glucose medium, but p s d l A p s d 2 A double mutants are auxotrophic for ethanolamine (Etn) or choline (Cho), depending on the carbon source (Trotter and Voelker, 1995; Birner et al., 2001; Storey e¢ al., 2001a). In mammalian cells, only one PtdSer decarboxylase has been identified so far. Like the yeast Psdlp, the rat liver enzyme was localized to the external side of the inner mitochondrial membrane (Zborowski et al., 1983). In most cell types, PtdSer decarboxylation is the major route of PtdEtn synthesis (Voelker, 1984; Vance, 1988).

3. Synthesis of Phosphatidylcholine by Methylation of Phosphatidylethanolamine In liver and in yeast, PtdCho is synthesized by methylation of PtdEtn. Yeast cells contain two PtdEtn methyltransferases (PEMT) (Kanipes and Henry, 1997):

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Pemlp/Cho2p methylates PtdEtn to yield phosphatidylmonomethylethanolamine, and Pem2p/Opi2p preferentially catalyzes the two subsequent methylation steps. Since Pem2p can replace Pernlp to some extent, and mono- and dimethylated PtdEtn appear to replace PtdCho sufficiently in S. cerevisiae, p e m l and pem2 single mutants are not auxotrophic for Cho, whereas growth of a p e m l pem2 double mutant requires exogenous Cho (Summers et al., 1988; Kodaki and Yamashita, 1989). In the yeast Schizosaccharomyces pombe, the two methyltransferases do not complement each other (Kanipes and Henry, 1997). In liver, all three methylation steps are catalyzed by a single PtdEtn methyltransferase, which was purified and cloned in mouse (Cui et al., 1993). Three human splice variants of PEMT cDNA were cloned, and the respective proteins were found at highest levels in liver, heart, and testis in a tissue-specific expression pattern (Walkey et al., 1999; Shields et aL, 2001). The majority of routine PEMT activity was found in mitochondria-associated membranes (MAM), a specific fraction of the ER (see Section III.B. 1), although a phospholipid methyltransferase activity has also been measured on the outer side of the plasma membrane of isolated rat hepatocytes (Bontemps and van den Berghe, 1998). A plant PEMT, which catalyzes all three methylation steps from PtdEtn to PtdCho, was recently cloned from spinach (Nuccio et aL, 2000). A screen for the Arabidopsis PEMT by complementation of the yeastpeml mutant failed, but led to the isolation of an S-adenosylmethionine:phosphoethanolamineN-methyltransferase which converts phosphoethanolamine to phosphocholine. The latter component can be used in the CDP-Cho pathway for PtdCho synthesis (Bolognese and McGraw, 2000), which appears to be the major route of PtdCho formation in tobacco. Three parallel interconnected pathways of PtdCho formation involving methylation of Etn, Etn-P, or PtdEtn were detected in flowering plants (McNeil et aL, 2000). Bacterial genes for PEMTs were cloned from Rhodobacter (Arondel et aL, 1993) and Acetobacter (Hanada et al., 2001). A novel Cho-dependent pathway of PtdCho biosynthesis was discovered in Sinorhizobium meliloti (de Rudder et aL, 1999; Sohlenkamp et al., 2000; Lopez-Lara and Geiger, 2001). In this pathway, a PtdCho synthase condenses Cho directly with CDP-DAG to form PtdCho in one step.

4. The Kennedy Pathway of Phosphatidylethanolamine and Phosphatidylcholine Formation In mammalian ceils, PtdCho is predominantly synthesized via the CDP-Cho branch of the Kennedy pathway (Kent, 1995). The CDP-Etn branch of the Kennedy pathway can significantly contribute to PtdEtn biosynthesis when mammalian cells are supplied with Etn (Tijburg et al., 1989; Arthur and Lu, 1993). Both branches of the Kennedy pathway are also active in yeast, although with minor efficiency (Daum et al., 1998). Nevertheless, yeast cells unable to synthesize PtdSer, or lacking PtdSer decarboxylase or PtdEtn methyltransferase activities, can be rescued by exogenous Etn/Cho.

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The first step of exogenous Etn/Cho utilization is their transport into the cell. In yeast, both Etn and Cho are imported by the same carrier, Ctrlp (Nikawa et al., 1986, 1990). A suppressor for the yeast ctrl mutation has been cloned from Torpedo electric lobe. Homologs of this choline transporter-like gene, CTL1, were also identified in mammals and C. elegans (O'Regan et al., 2000). Intracellular Etn and Cho are phosphorylated to Etn-P and Cho-P, respectively, by cytosolic Etn and Cho kinases. Two enzymes with overlapping substrate specificities, Ekilp (Kim et al., 1999) and Ckilp (Yamashita and Hosaka, 1997), were cloned in yeast. Purification of the yeast Ckilp revealed that native Ckilp exists in oligomeric structures of dimers, tetramers, and octamers (Kim et al., 1998). Several CKI isoforms with dual substrate specificity for Etn and Cho are present in mammalian cells (Uchida and Yamashita, 1992; Uchida, 1994; Aoyama et al., 1998, 2000). A number of Etn-specific kinases (EKIs) were purified from rat liver (Weinhold and Rethy, 1974; Uchida, 1997) and human liver (Draus et al., 1990) and cloned from Drosophila (Pavlidis et al., 1994) and human (Lykidis et al., 2001a). In the next step, Etn-P and Cho-P are converted to CDP-Etn and CDP-Cho, catalyzed by CDP-phosphoethanolamine cytidylyltransferase (ECT) and CDPphosphocholine cytidylyltransferase (CCT) (Bladergroen and van Golde, 1997; Kent, 1997). Only one ECT exists in eukaryotes, which has been purified from rat liver (Vermeulen et al., 1994) and cloned in human (Nakashima et al., 1997) and yeast (Min-Seok et al., 1996). The plant ECT activity was localized to the outer mitochondrial membrane and ER (Wang and Moore, 1991), whereas mammalian ECT is present in cisternae of the rough ER and the cytosol (van Hellemond et al., 1994). Three CCT isoenzymes--CCTc~, CCTfll, and CCT/32--were identified in human cells (Kalmar et al., 1994; Lykidis et al., 1998, 1999), but only one enzyme of this kind, CCT1/PCT1, was cloned in yeast (Tsukagoshi et al., 1991). Human CCTc¢ is expressed ubiquitously, while CCTfl is expressed primarily in placenta and testis (Lykidis et al., 1998). All three human CCT isoforms associate to some extent with the ER (Lykidis et al., 1999). The majority of CCTo~, however, is localized to the nucleus in most mammalian cells (Lykidis et al., 1998; DeLong et al., 2000), and CCTc~ of pulmonary tissues localizes to the cytosol (Ridsdale et al., 2001). Additional cDNAs for CCTs have been cloned from several mammalian cell types (Kent, 1997), plants (Jones et al., 1998; Choi et al., 2001; Kent, 1997), P l a s m o d i u m f a l c i p a r u m (Kent, 1997), Caenorhabditis elegans (Friesen et al., 2001a), and Streptococcus pneumoniae (Campbell and Kent, 2001). In the last step of the Kennedy pathway, CDP-Etn and CDP-Cho react with DAG to form PtdEtn and PtdCho (McMaster and Bell, 1997a,b). Two CDPcholine: 1,2-diacylglycerol cholinephosphotransferase activities were described in yeast. Cptlp accepts only CDP-choline as a substrate, while Eptlp can utilize CDPCho and CDP-Etn with similar efficiency (McGee et al., 1994; McMaster and Bell, 1994; Williams and McMaster, 1998). Cpt 1p- and Ept lp-derived cholinephosphotransferase activities can significantly overlap in vivo with Eptlp, contributing

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60% to net PtdCho synthesis (Henneberry et al., 2001). In mammalian cells, cholinephosphotransferase and ethanolaminephosphotransferase revealed similar substrate specificities. The human CPT1 gene product complements for the loss of PtdCho, but not of PtdEtn synthesis in a c p t l A e p t l A yeast mutant (Henneberry et al., 2000), whereas the human CEPT1 gene product rescues both branches of the yeast pathway (Henneberry and McMaster, 1999). 5. Endogenous Sources of Free Ethanolamine and Choline Alternatively to exogenous Etn/Cho, free Etn/Cho produced by lipolytic cellular activities is recycled to PtdEtn/PtdCho formation via the Kennedy pathway. Catabolic enzymes responsible for degradation of aminoglycerophospholipids comprise the phospholipases D, producing free Etn/Cho from PtdEm/PtdCho, and the phospholipases C, yielding Etn-P/Cho-P. Moreover, catabolic enzymes degrading sphingomyelin or phosphorylated sphingoid bases produce Cho-P or Etn-P, respectively. In fibroblasts, phosphocholine formed through lysosomal degradation of PtdCho and sphingomyelin is exported from the lysosome and incorporated into PtdCho without prior hydrolysis to choline (Jansen et al., 2001). In the yeast, as in mammalian cells, the CDP-Etn branch of the Kennedy pathway is linked to sphingolipid catabolism through the reaction of dihydrosphingosinephosphate lyase, Dpllp, which releases long-chain aldehydes and Etn-P (Saba et al., 1997; Mandala et al., 1998). Overexpression of Dpllp rescues the Etn auxotrophy of yeast p s s l A / c h o l A and p s d l A psd2A mutants, since the produced Etn-P is reutilized for PtdEtn biosynthesis by the CDP-Etn branch of the Kennedy pathway (Birner et al., 2001). B. Cell Biological Function of Phosphatidylserine 1. Phenotypes of Cells with Defects in Phosphatidylserine Synthesis PtdSer is essential for viability of mammalian cells (Kuge et al., 1986a; Voelker and Frazier, 1986). Exogenous serine is required for synthesis of PtdSer and sphingolipids in cultured hippocampal neurons (Mitoma et al., 1998). In contrast to mammalian cells, PtdSer-deficient yeast cells (pssl/chol) are viable, but depend on supplementation with Etn or Cho for alternative synthesis of PtdEtn and PtdCho through the Kennedy pathway (Atkinson et al., 1980a,b). pssl/chol mutants have a growth defect on nonfermentable carbon sources and accumulate respiratory-defcient (petite) cells. Yeast diploids homozygous for the pssl/chol mutation are defective in sporulation. Loss of PtdSer synthesis results in fragmentation of vacuoles, increased susceptibility to Ca 2+, Zn 2+, Mn 2+, L-lysine, and L-arginine (Hamamatsu et aL, 1994), and a defect in tryptophan transport (Nakamura et al., 2000). Escherichia coli pssA mutants are unable to synthesize

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PtdSer and PtdEtn, but are viable when supplemented with tryptophan and divalent cations (DeChavigny et al., 1991).

2. Species and Localization of Phosphatidylserine In rat liver microsomes, the pattern of molecular species of PtdSer is markedly different from that of the precursor PtdEtn and PtdCho inasmuch as stearic acid/ arachidonic acid (18:0/20:4) and stearic acid/docosahexaenoic acid (18:0/22:6) containing PtdEtn and PtdCho are preferentially used for PtdSer synthesis via base exchange (Ellingson and Seenalah, 1994). Docosahexaenoic acid deficiency results in reduced PtdSer levels (Garcia et al., 1998), while treatment of cells with docosahexaenoic acid promotes PtdSer enrichment in neuronal membranes (Kim et al., 2000). Although both PtdSer and PtdIns are derived from CDP-DAG in yeast, these two phospholipids vary greatly in their degree of acyl chain saturation (Wagner and Paltauf, 1994; Schneiter et al., 1999). This observation may be explained by different pools of CDP-DAG or different substrate specificities of PtdSer synthase and PtdIns synthase. PtdSer is the major phospholipid of the yeast plasma membrane (Zinser et al., 1991; Zinser and Daum, 1995; Tuller et al., 1999). In this compartment, the majority of molecular species of PtdSer contain one saturated acyl chain at the expense of diunsaturated species (Schneiter et al., 1999). A similar enrichment of saturated PtdSer species was reported for the plasma membrane of mammalian cells (van Meer, 1998). In plants, vesicles budding from the ER are enriched in PtdSer synthesized by base exchange, especially in species containing very long-chain fatty acids (VLCFA) (Sturbois-Balcerzak et al., 1999). 3. Asymmetric Distribution of Phosphatidylserine in Membranes PtdSer is abundant in the cytoplasmic leaflet but more or less absent from the outer leaflet of the plasma membrane of eukaryotes (Devaux, 1991). Enzymes facilitating aminoglycerophospholipid transport across the plasma membrane, such as aminophospholipid translocase, scramblase, and floppases, were identified (for details see Section III.C). Inhibition of aminophospholipid translocase (which facilitates the transport of PtdSer and PtdEtn from the outer to the inner leaflet) or activation of scramblase (which catalyzes random transport of phospholipids across the bilayer) results in loss of plasma membrane asymmetry. Plasma membrane asymmetry depends on many different parameters: it is decreased with age in human lymphocytes and platelets corresponding to aging changes in superficial membrane fluidity (Noble et aL, 1999; Pereira et aL, 1999), regulated by store-operated Ca 2+ entry (Kunzelmann-Marche et aL, 2001) and lipid raft integrity (Kunzelmann-Marche et al., 2002), and affected by apoptogenic

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agents (Schlegel and Williamson, 2001). Loss of lipid asymmetry creates a procoagulant surface on platelets (Comfurius et al., 1994). During formation of a platelet plug at the damaged vessel wall, factors IXa and VIIIa form the "tenase" complex, leading to activation of factor X on the surface of activated platelets. Subsequently, factors Xa and Va form the "prothrombinase" complex, which catalyzes the formation of thrombin from prothrombin and leads to fibrin formation. An enhancement of negatively charged PtdSer in the outer membrane leaflet resulting from a breakdown of the phospholipid asymmetry is essential for the formation of the procoagulant surface (Solum, 1999). PtdSer in membranes was found to bind and regulate factor Xa, an activator of prothrombin (Wu et al., 2002; Banerjee et aL, 2002; Srivastava et al., 2002), to activate factor Va (Srivastava et al., 2001) by conforrnational changes (Zhai et al., 2002), and to interact specifically with prothrombin (Falls et aL, 2001). Exposure of PtdSer on the outer leaflet of the plasma membrane is an early sign of apoptosis (Schlegel and Williamson, 2001). Externalized PtdSer serves as a trigger for macrophage recognition and phagocytosis of apoptotic cells (Fadok et al., 1992, 2001a). Recent data suggest that both stimulation of phospholipid scramblase and inhibition of aminophospholipid translocase are relevant for PtdSer translocation to the surface during apoptosis. Caspase 3 activation stimulates phospholipid scramblase activity via protein kinase C8 (Frasch et aL, 2000) and inhibits aminophospholipid translocase activity, leading to PtdSer externalization and phagocytosis of oxidatively stressed erythrocytes (Mandal et aL, 2002). Moreover, selective oxidation of PtdSer by cytochrome c, which is set free from mitochondria into the cytosol during apoptosis, may cause inhibition of the aminophospholipid translocase (Tyurina et al., 2000). Phagocytosis of apoptotic cells is mediated by a cell-surface PtdSer receptor (PtdSerR) on macrophages (Fadok et al., 2001b; Somersan and Bhardwaj, 2001). Recently, it was shown that swelling of cells in oncosis, the early phase of primary necrosis, is also associated with exposure of PtdSer (Lecoeur et aL, 2001). Thus, PtdSer, which is externalized on both apoptotic and necrotic cells, may not be specific for the recognition of either one (Cocco and Ucker, 2001). PtdSer exposure on the erythrocyte surface endows the cell with the propensity of adhering to vascular endothelium. Red cell-endothelial adhesion might have implications in the pathology of sickle cell disease, falciparum malaria, and diabetes, which are associated with loss of erythrocyte membrane asymmetry (Closse et al., 1999; Manodori et al., 2000; Setty et al., 2002). Moreover, the presence of anionic phospholipids, such as PtdSer, on the exterior face of the plasma membrane increases uptake of the terminal complement proteins C5b-7 and subsequent hemolysis, which may also be involved in the pathology of sickle cell disease (Liu et al., 1999). Infection of host cells by Chlamydia induces rapid, transient PtdSer externalization, which may he an important factor in the pathogenesis of chlamydial infections (Goth and Stephens, 2001).

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Maintenance of asymmetric distribution of phospholipids is important not only for membrane mechanical stability of red blood cells (Manno et al., 2002), but also for membrane budding and endocytosis (Farge et al., 1999). Stimulation of the aminophospholipid translocase provokes endocytic-like vesicles in erythrocytes and stimulates endocytosis in K562 cells (Devaux, 2000). Scrambling may induce echinocytosis and stimulate microvesiculation of erythrocytes (Kamp et al., 2001). Exocytic release of neurotransmitters from cholinergic nerve terminals isolated from electric ray is associated with rapid transbilayer phospholipid redistribution (Lee et al., 2000). 4. Interaction of Phosphatidylserine with the Cytoskeleton Interaction with negatively charged phospholipids such as PtdSer can serve to enrich cytoskeletal proteins at the membrane in a Ca2+-dependent process (Axelrod and Wang, 1994). The negatively charged head group of PtdSer controls Ca 2+ binding by lowering the electric surface potential and elevating cation concentration at the membrane/water interface (Huster et al., 2000). PtdSer binds to annexins in a Ca2+-dependent manner (Reutelingsperger, 2001) through a highly conserved PtdSer binding site (Montaville et al., 2002). Annexin V appears to form a two-dimensional network adsorbed to a lipid bilayer, strongly reducing lateral diffusion of PtdSer (Saurel et al., 1998). Actin, but also tropomyosin, myosin and the Ca2+/calmodulin-binding protein caldesmon, binds PtdSer of the ER membrane (Makowski et al., 1997). The PtdSer-caldesmon interaction reduces the polar head group mobility of phospholipids and inhibits PtdEtn biosynthesis via a phospholipid base-exchange reaction with PtdSer as substrate. 5. Involvement of Phosphatidylserine in Signaling and Enzyme Activation PtdSer is also involved in signaling because it is an essential cofactor for the activation of protein kinase C (PKC) (Bell and Bums, 1991; Nishizuka, 1992) through Ca2+-dependent binding to certain domains of the protein (Verdaguer et al., 1999). Similarly, activation of phospholipase C (PLC) appears to involve a ternary complex of PtdSer, Ca 2+, and the C2 domain ofPLC (Lomasney et al., 1999). Moreover, PtdSer has been shown to regulate the activities of many other enzymes, such as DAG kinase (Sakane et al., 1991), dynamin (Tuma et al., 1993), B-Raf protein kinase (Ghosh et al., 1994), Na+/K+-ATPase (Stekhoven et al., 1994), acetylcholine receptor (Sunshine and McNamee, 1992), and glutamate receptor (Gagne et al., 1996). The Semliki Forest virus mRNA capping enzyme requires association with anionic membrane phospholipids, especially PtdSer, which may explain why the replication complexes of all positive-strand RNA viruses are associated with membranes (Ahola et al., 1999).

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C. Cell Biological Function of Phosphatidylethanalamine

1. Phenotypes of Cells with Defects in Phosphatidylethanolamine Biosynthesis PtdEtn is a major membrane lipid component of prokaryotes and eukaryotes. With its small head group and its conelike shape, PtdEtn is a non-bilayer-forming lipid. Escherichia coli pssA or p s d mutants are conditional-lethal and rely on supplementation with Mg 2+, Ca 2+, or Sr2+ (DeChavigny et al., 1991). The deficiency of PtdEtn is compensated by elevated levels of PtdGro and CL in these mutants, suggesting that PtdEtn is not essential for viability of E. coli. The presence of cations guarantees that the bilayer-to-nonbilayer transition temperature remains conserved, which is essential for function (e.g., fusion, division, and budding reactions) and provides the lateral packing pressure needed for proper protein folding (Rietveld et al., 1993; Morein et al., 1996). In contrast to E. coli, pss and p s d mutants of Bacillus subtilis grow without supplementation of divalent cations (Matsumoto, 1997). In these mutant strains, increased levels of glucosyldiacylglycerol appear to compensate for the lack of PtdEtn. In contrast, PtdEtn synthesis in yeast is essential (Birner et al., 2001; Robl et al., 2001 ; Storey et al., 2001a).

2. Phosphatidylethanolamine Affects Enzymes in Prokaryotes and Eukaryotes Escherichia colipssA orpsd mutants exhibit defects in secondary transport systems

for sugars (lactose permease and melibiose transporter) and amino acids (proline, tryptophan, and lysine) (DeChavigny et al., 1991). In yeast, amino acid transport depends on PtdEtn (Rob1 et al., 2001). PtdEtn was shown to act as a chaperone for E. coli lactose permease (LacY) (Bogdanov and Dowhan, 1998; Bogdanov et al., 1999). The N-terminal half of LacY in PtdEtn-lacking E. coli adopts an inverted topology (Bogdanov et al., 2002). Normal conformation and topology of at least one LacY subdomain accompanied by restoration of active transport is triggered by post-assembly synthesis of PtdEtn. Similarly, lyso-PtdEtn of E. coli, whose amount was found to increase upon heat shock, displays chaperone-like properties on hydrophilic proteins, such as citrate synthase and ~-glycosidase (Kern et al., 2001). The nonbilayer property of the membrane is also essential for the function of the SecYEG translocase of E. coli and B. subtilis (Rietveld et al., 1995; van der Does et al., 2000). PtdEtn mediates insertion of the catalytic domain of leader peptidase in membranes (van Klompenburg et al., 1998; van den Brink-van der Laan et al., 2001) and increases the association rate of SecA, a soluble component of the translocase, with membranes (Ahn et al., 2001). A similar mechanism for binding of FtsY, the E. coli homolog of the eukaryotic signal recognition particle receptor ot-subunit, to the cytoplasmic membrane via interactions with PtdEtn

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has been proposed (Millman et al., 2001). Depletion of PtdEtn affects secretion and transcription of the E. coli alkaline phosphatase (Mikhaleva et al., 200l). In mammals, PtdEtn modulates the activity of Ca 2+-ATPase in the sarcoplasmic reticulum (Hunter et al., 1999).

3. Phosphatidylethanolamine Affects Motility of Prokaryotes Escherichia colipssA or psd mutants are impaired in motility and chemotaxis (Shi et al., 1993). Pseudomonas aeruginosa exhibits directed twitching mobility, a type IV pilus-dependent form of motility, when attracted by PtdEtn (Kearns et al., 2001). PtdEtn also acts as a chemattractant for the soil bacterium Myxococcus xanthus, but this effect depends on an extracellular matrix-associated zinc metalloprotease (Kearns and Shimkets, 2001; Kearns et al., 2002). The bundle-forming pilus of enteropathogenic E. coli has a binding specificity for PtdEtu, suggesting

that PtdEtn may act as a bundle-forming pilus receptor for bacterial autoaggregation and may promote localized adherence to host cells, both of which contribute to bacterial pathogenesis (Khursigara et al., 2001). Enterohemorrhagic E. coli induce apoptosis, which augments bacterial binding and PtdEtn exposure on the plasma membrane outer leaflet (Barnett Foster et al., 2000).

4. Phosphatidylethanolamine Affects Cytokinesis PtdEtn has been implicated in cytokinesis in E. coli (Mileykovskaya et al., 1998). Lack of PtdEtu may affect the correct interaction of FtsZ, a protein essential for an early step in cytokinesis, with membrane nucleation sites, thereby altering tile FtsZ ring structure so as to prevent or delay its constriction. In mammalian cells, PtdEtn is also required for completion of cytokinesis (Emoto et al., 1996; Emoto and Umeda, 2000, 2001). Immobilization of cell surface PtdEtu by a PtdEtn-binding peptide inhibits disassembly of the contractile ring components, including myosin II and radixin, resulting in formation of a long cytoplasmic bridge between daughter cells. Thus, transbilayer redistribution of PtdEtu to the surface of the cleavage furrow is essential for contractile ring disassembly at the final stage of cytokinesis, and appears to play a critical role in mediating the coordinated movements between the contractile ring and the plasma membrane required for the proper progression of cytokinesis. 5. Phosphatidylethanolamine Is a Precursor for Complex Membrane Components PtdEtn acts as a donor of Etu-P or Etn-pyrophosphate for the biosynthesis of bacterial macromolecules, such as lipopolysaccharide (LPS) (Kanipes et al., 2001) and periplasmic membrane-derived oligosaccharide (MDO) (Huijbregts et al., 2000). In eukaryotes, PtdEtn is the precursor for the Etn-P bridge linking the

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C-terminal amino acid of glucosylphosphatidylinositol(GPI)-anchored proteins to GPI (Menon and Stevens, 1992), and also for Etn-P side chain addition to the mannose residues of the GPI anchor (Imhof et aL, 2000). Most of the yeast Etn-P transferases are essential, as is GPI-anchor synthesis, for cell wall maintenance (Hong etal., 1999), explaining the essential minimal requirement for PtdEtn (Birner etal., 2001). In mammalian cells, biosynthesis of the major lipofuscin fluorophore, pyridinium bisretinoid, involves the formation of the precursor PtdEtn-bisretinoid in the photoreceptor outer segment membrane (Liu et al., 2000). In plants, PtdEtn is the precursor for N-acyl PtdEtn, a minor phospholipid involved in signal transduction and membrane protection during stress response (Chapman, 2000). The plant eukaryotic elongation factor lot (eEF-1A) is covalently attached to PtdEtn (Ransom etaL, 1998), although this posttranslational modification does not affect function of eEF-1A. PtdEtn is also required for autophagy in yeast (Ichimura et al., 2000). Covalent linkage of Apg8/Aut7p to PtdEtn by an ubiquitin-like system is essential for membrane dynamics during autophagy and localization of autophagy factors to the pre-autophagosomal structure in the vicinity of the vacuole (Suzuki et aL, 2001). 6. Phosphatidylethanolamine Is an Essential Mitochondrial Component The requirement for PtdEtn in yeast is more stringent on nonfermentable carbon sources, i.e., when mitochondria are fully developed, than on fermentable carbon sources (Birner et al., 2001; Storey et al., 2001 a). Under nonfermenting conditions, synthesis of PtdSer in the ER, transport of PtdSer from the ER to mitochondria and subsequent decarboxylation to PtdEtn by Psdlp, or synthesis of PtdEtn through the Kennedy pathway becomes essential. A p s d l A mutant contains only a low level of mitochondrial PtdEtn and has an increased tendency to form respirationdeficient cells (petites) on glucose (Birner et aL, 2001) owing to instability of the mitochondrial genome (Birner et aL, unpublished). Thus, Psdlp is the major supplier of mitochondrial PtdEtn, confirming early results from our laboratory (Kuchler et al., 1986). Similar results were obtained for mammalian cells, from which the majority of mitochondrial PtdEtn is derived from decarboxylation of PtdSer and not from the CDP-Etn pathway (Voelker, 1984; Kuge et aL, 1986b; Shiao et al., 1995). 7. Significance of the CDP-Ethanolamine Pathway The CDP-Etn pathway is required for synthesis of Etn plasmalogen species (alkenyl/acyl species) in neuronal and cardiac cells. In these cells, Etn plasmalogens account for half of total phosphoglycerides (Yorek et al., 1985). In heart tissue, the decarboxylation pathway is less active than in liver or kidney (Arthur and Page, 1991). In rat hepatocytes, the two pathways of PtdEtn synthesis seem to

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have specific functions. PtdEtn and PtdCho derived from PtdSer are mainly used for the synthesis of secreted lipoproteins, whereas CDP-Etn derived PtdEtn and PtdCho is used for the formation of intracellular membranes (Vance and Vance, 1986; Vance, 1988).

D. Fundamental Role of Phosphatidylcholine as Bulk Lipid of Membranes 1. Phosphatidyleholine in Prokaryotes PtdCho is not only a major component of eukaryotic membranes, but is also present in some bacteria such as Acetobacter, Rhodobacter, Zymomonas, Rhizobium, and Spirochetes without being essential. As recent examples, PtdCho-deficient Acetobacter aceti was shown to be viable (Hanada et al., 2001), and Sinorhizobium meliloti double mutants bearing defects in PtdEtn methyltransferase and PtdCho synthase showed reduced growth but survived (de Rudder et al., 2000). The PtdCho synthase appears to be common to a number of bacteria displaying symbiotic or pathogenic associations with a eukaryotic host that provides choline for this pathway.

2. Essentiality of Phosphatidylcholine in Eukaryotes PtdCho is an essential structural component of eukaryotic membranes, comprising about 50% of the total amount of glycerophospholipids (Howe and McMaster, 2001). PtdCho is the major bilayer-forming lipid because of its large head group and cylindrical shape. Moreover, PtdCho is the precursor of mammalian sphingomyelin and a number of lipid-signaling molecules. Choline is an essential nutrient for mammalian cells. Its absence leads to apoptosis in PC12 cells, which is associated with a decrease of PtdCho and sphingomyelin, accumulation of ceramide and DAG, and activation of a caspase (Yen et al., 1999). Defects in the CDP-Cho pathway also cause apoptosis in mammalian cells (Cui et al., 1996; Anthony et al., 1999). PtdCho synthesis is important during mammalian embryonic development (Fisher et al., 2001, 2002), and its absence induces apoptosis in primary cultures of fetal neurons (Yen et aL, 200l). A decrease in PtdCho, which causes accumulation of triacylglycerol (TAG) in lipid droplets, also leads to apoptosis in Jnrkat T-cells (A1-Saffar et al., 2002). In contrast to mammalian cells, the Kennedy pathway is not essential in yeast (McGee et aL, 1994), although it is active whenever choline is present exogenously or as degradation product of PtdCho (Cleves et al., 1991). Yeast cells with defects in the PtdEtn methylation pathway are strictly auxotrophic for choline, demonstrating that PtdCho is essential as a membrane component (Summers et al., 1988; Kodaki and Yamashita, 1989; Kanipes and Henry, 1997).

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In rat liver, 70% of newly synthesized PtdCho is derived from the CDP-Cho pathway and 30% from the PEMT route (Reo et al., 2002). Mice homozygous for a disruption of the gene encoding PEMT display a normal phenotype as long as supplemented with choline (Walkey et al., 1997, 1998; Waite et al., 2002). Thus, the PEMT pathway appears to be maintained during evolution to provide PtdCho when dietary choline is insufficient. However, CDP-Cho and PtdEtn methylation pathways were shown to have opposite effects on proliferative characteristics of the liver. While increased expression of the CDP-Cho pathway favors the proliferation of hepatocytes, overexpression of PEMT strongly inhibits the growth of hepatorna cell lines (Vance et al., 1997) and induces apoptosis (Tessitore et al., 1999). Moreover, PtdCho synthesized by PEMT does not substitute for PtdCho synthesized through the CDP-Cho pathway. A block in the CDP-Cho pathway causes apoptosis in mammalian cells (Esko et al., 1981; Cui et al., 1996; Anthony et al., 1999) which cannot be rescued by expression of rat liver PEMT (Houweling et al., 1995). This failure appears to be due to inadequate long-term PtdCho synthesis, which causes TAG and lyso-PtdCho accumulation (Waite and Vance, 2000). Differences in the two PtdCho-synthesizing pathways in liver cells do not result from different subcellular locations of enzymes involved, but rather from the synthesis of distinct PtdCho pools (DeLong et al., 1999). While PtdCho derived from the CDP-Cho pathway mainly contains medium-chain and saturated fatty acid, PtdCho from the PEMT pathway comprises significantly more long-chain and polyunsaturated species, contains a higher percentage of arachidonate, and is in general more diverse. Kinetic analysis of PtdEtu and PtdCho biosynthesis in mammalian cells suggested that the CDP-Etu and CDP-Cho pathways and the PEMT pathway are channeled processes (Bladergroen et al., 1998; Reo et al., 2002). In this model, enzymes involved in a metabolic pathway are not randomly distributed throughout the cell or even through a compartment, but rather reaction intermediates pass from one enzyme of a biosynthetic sequence to the next without equilibration of pools. In fact, the intermediate metabolites directly derived from exogenous Cho and Etn do not completely mix with the intracellular pools, but are preferentially used for phospholipid synthesis (Reo et al., 2002). The cytoskeleton may play a role in the formation of functional complexes of enzymes and substrates (Bladergroen et al., 1998).

3. Interaction of Phosphatidylcholine with Enzymes and Traffic Vesicles Examples of specific interaction of PtdCho with proteins are rare. The most prominent example is (R)-3-hydroxybutyrate dehydrogenase, a mitochondrial enzyme that selectively binds PtdCho at its C terminus (Loeb-Hennard and McIntyre, 2000). PtdCho also interacts with glycerol-3-phosphate dehydrogenase (Gut2p) of yeast mitochondria, although the significance of this interaction is not clear (Janssen et al., 2002).

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Interaction of PtdCho metabolism with secretion of proteins was shown in yeast (Kent and Carman, 1999). PtdCho synthesis was reported to be a negative regulator of yeast Golgi secretory function through the Ptdlns/PtdCho transfer protein Secl4p (Xie et al., 2001). Ptdlns/PtdCho transfer proteins are abundant cytosolic proteins that were originally identified by their ability to act in vitro as specific transporters of Ptdlns and PtdCho between membranes. Recent data, however, suggest that Secl4p acts as a molecular switch depending on PtdCho and Ptdlns binding. In analogy to the mammalian Ptdlns/PtdCho transfer proteins, the yeast Sec14p inhibits consumption of DAG and production of PtdCho by the CDPcholine pathway in its PtdCho-bound form and stimulates Ptdlns-4-P synthesis in its Ptdins-bound form (Li et al., 2000). In contrast to PtdCho, DAG and/or its metabolic precursor phosphatidic acid (PA), produced by PtdCho turnover through phospholipases, and Ptdlns-4-P and its metabolically related product Ptdlns-4,5-P2 may stimulate vesicle biogenesis (see Section II.D.4). 4. Phosphatidylcholine Is a Precursor of Signaling Molecules

In addition to its structural role as a bulk phospholipid, PtdCho plays a significant role as a precursor for signaling molecules such as PA, DAG, lyso-PtdCho, and arachidonate (Cockcroft, 2001; Steed and Chow, 2001). In mammalian cells~ PtdCho-specific PLDs are coupled to immune receptors (Melendez and Allen, 2002). PA produced by turnover of PtdCho functions as a growth factor, stimulates PLC and protein/lipid kinase activity, mobilizes Ca 2+ flux, activates NADPH oxidase, and induces hormone release, platelet aggregation, gene transcription, and changes in cytoskeletal dynamics (Cockcroft, 2001; Steed and Chow, 2001). PA production by PLD is stimulated by monomeric GTPases of the Rho family, modulating actin assembly (Extort, 1997; Mackay and Hall, 1998). Moreover, PA produced by PLD-mediated turnover of PtdCho promotes vesicular budding from the Golgi (Siddhanta and Shields, 1998) and stimulates secretion (Cohen and Brown, 2001). Mammalian PLD1 has been localized in part to the Golgi, which is consistent with its involvement in vesicular trafficking (Freyberg et al., 2001). Inhibition of PA synthesis alters Golgi structure and inhibits secretion in endocrine cells, probably because PA is required for stimulation of Ptdlns-4phosphate-5-kinase PIP5K (Jones et aL, 2000; Siddhanta et al., 2000). On the other hand, mammalian PLDs require Ptdins-4,5-P2 for enzymatic activity. A pleckstrin homology domain required for Ptdlns-4,5-P2-dependent PLD localization and activation has been proposed (Hodgkin et aL, 2000). Thus, a positive stimulatory loop is created that would generate both PA and Ptdlns-4,5-P2 at specific sites within the cell. Therefore synthesis of PA and Ptdlns-4,5-P2 could be spatially and temporally regulated to modulate secretion and acfin assembly. Lyso-PtdCho produced by PLA2 appears to be involved in a variety of signaling pathways affecting cell proliferation, adhesion and differentiation of lymphoid cells, regulation of myocardial sensitivity to cholinergic stimulation, and

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contractility of arterial smooth muscle cells (Prokazova et al., 1998). Moreover, lyso-PtdCho induces both apoptotic and nonapoptotic death (Hsieh et al., 2000), inhibits insulin-induced Akt activation through protein kinase C-or in vascular smooth muscle cells (Motley et al., 2002), and induces Ca2+ mobilization and membrane current via a Rho-activation-dependent PLC pathway in endothelial cells (Yokoyama et aL, 2002). Membrane breakdown by PLA2 is involved in acute and chronic neurodegeneration, such as stroke and senile dementia (Klein, 2000), but also in schizophrenia (Schmitt et al., 2001). In yeast, deacylation of PtdCho to glycerophosphocholine is accelerated by the stress of elevated temperature (Dowd et aL, 2001).

E. Regulation of Aminoglycerophospholipid Biosynthesis

1. Regulation of Phosphatidylserine Synthesis In E. coli, reconstituted PtdSer synthase is activated by anionic phospholipids, such as PtdGro, CL, and PA (Rilfors et al., 1999). In mammalian cells, regulation

of PtdSer synthesis is more complex owing to the unique functions of the two PSS enzymes. Murine PSS1 and PSS2 have different substrate specificities and are differentially expressed in different tissues (Stone and Vance, 1999). In contrast to PSS 1, PSS2 is inhibited by PtdSer. Overexpression ofPSS 1 but not of PSS2 inhibits PtdEtn biosynthesis via the CDP-Etn pathway. In CHO cells, however, both PSS 1 and PSS2 are inhibited by PtdSer (Kuge et al., 1998, 1999). PtdSer homeostasis in glioma C6 cells is modulated by stimulation of PtdSer synthesis and inhibition of PtdSer decarboxylase activity by sphingosine, sphingosylphosphorylcholine, and sphingosine-l-phosphate (Wojcik and Baranska, 1999). In Jurkat T-cells, PtdSer synthesis is inhibited by hydrogen peroxide, but stimulated by antioxidants, suggesting that PtdSer synthesis is regulated by the redox status of the cells (Pelassy et al., 2001). In addition to transcriptional control (see Section II.E.4), the branch points of the phospholipid biosynthetic pathway are subject to biochemical regulation in yeast (Carman and Henry, 1999). The PtdSer to Ptdlns ratio is subjected to biochemical regulation of PtdSer synthase, Psslp. Activity of Psslp is positively affected by PA and diacylglycerol pyrophosphate, and reduced by phosphorylation, inositol, CL, DAG, sphingoid bases, and CTP.

2. Regulation of Phosphatidylethanolamine Synthesis It prokaryotes, a balanced phospholipid composition appears to be regulated by independent feedback regulation rather than by coordinate regulation of PtdSer synthase and PtdGroP synthase. In E. coli, activation ofPtdSer synthase by PtdGro, CL, and PA may be part of a regulatory mechanism that keeps a balance between

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PtdEtn and the total amount of PtdGro and CL (Rilfors et al., 1999). PtdSer decarboxylase is not rate-limiting and is regulated only by supply of its substrate PtdSer in E. coli. In mammalian cells, regulation of the two PtdEtn biosynthetic pathways, PtdSer decarboxylation and the CDP-Etn pathway, is coordinated in response to ethanolamine in the cell culture media to prevent excess production of PtdEtn (Tijburg et al., 1989; Arthur and Lu, 1993).

3. Regulation of Phosphatidylcholine Synthesis The C C T gene product, CDP-phosphocholine cytidylyltransferase, catalyzes the rate-limiting step in PtdCho biosynthesis of higher eukaryotes (Vance etal., 1980). Regulation of CCT has been studied extensively because PtdCho is the major precursor for PtdSer and PtdEtn in mammalian cells (see Section II.A.1) and its biosynthesis therefore affects total membrane homeostasis. Mammalian PtdCho synthesis is primarily regulated by reversible membrane association of CCTa, although both product and substrate of CCTa are water-soluble (Comell and Northwood, 2000). Stimulation of cellular PtdCho synthesis correlates with translocation of inactive CCTa from the intranuclear space to an active membrane-bound form at the nuclear envelope/ER, or from the cytosol to the plasma membrane and the ER (Ridsdale et al., 2001). Two domains of CCTa are involved in its regulation by lipids: the three amphipathic a-helical repeats that interact with both neutral and anionic lipid mixtures, and the C-terminal 57 amino acid residues that interact exclusively with anionic lipids (Lykidis et al., 2001b). Interaction of lipids with these domains removes an inhibitory constraint on CCT activity. Thus, the CCTa catalytic fragment has higher activity than the full-length protein in the absence of lipids (Ffiesen et al., 1999). CCT lipid association and activation appear to depend on the physical properties of the membrane, such as the stored curvature elastic stress caused by cone-shaped lipids like PtdEtn which accumulates during PtdCho starvation (Attard et al., 2000; Davies et al., 2001). Thus, a physical feedback signal may also play a role in the control of membrane lipid synthesis. In the lung, tissue injury during inflammation, which involves the release of tumor necrosis factor (TNF) and generation of ceramide, is accompanied by inhibition of surfactant PtdCho metabolism (Mallampalli et aL, 2000). This inhibition occurs either by modulating CCT protein stability via the ubiquitin/proteasome pathway or by generation of lyso-PtdCho as a consequence of the activation of cytosolic PLA2 (Awasthi et al., 2001). PtdCho homeostasis is achieved by a batance between the opposing activities of CCT and PLA2 (Lykidis and Jackowski, 2001). PLA2 activity is modulated in response to changes of PtdCho levels in CHO cells (Barbour et aL, 1999). In plants, PEMT overexpression and supplementation with Etn leads to an increase of free Cho without affecting the PtdCho content, suggesting that PtdCho degradation must be very efficient in these cells (McNeil et al., 2001).

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Cell-permeable ceramide may inhibit PtdEtn and PtdCho synthesis in rat fibroblasts by inhibiting CPT and EPT, which catalyze the final steps in the CDP-Cho and CDP-Etn branches of the Kennedy pathway (Bladergroen et al., 1999). In neuronal cells, an increased level of free DAG enhances the activity of CPT, causing a signifcant increase in the rate of PtdCho biosynthesis (Araki and Wurtman, 1997). In mammalian cells, PtdCho synthesis is also regulated in a cell-cycledependent manner, with PtdCho production peaking in G0/G1 state, declining during transition to G1/S, and remaining low during S and G2/M states (Tseu et al., 2002). These changes are not due to alterations in CCTa expression and degradation, but are a consequence of changes in CCTc~ activity through phosphorylation and membrane association. Increased transcription of the CCTa gene occurs during S-phase in preparation for mitosis (Golfman et al., 2001). Transcription of the murine CTTo~ is modulated by transcriptional enhancer factor-4 (TEF4), which can act as a suppressor via its direct binding to an upstream enhancer element or as an activator via its interactions with the basal transcriptional machinery (Sugimoto et al., 2001). Transcription of CCT is stimulated by lipid deprivation (Ryan et al., 2000). Expression of the murine CCT gene is inhibited by sterols via a sterol response element (SRE) and activated by mature (cleaved) SRE-binding proteins (SREBPs) (Lagace et al., 2000; Kast et al., 2001; Mallampalli et al., 2002). SREBPs are transcription factors that belong to the basic helix-loop-helix- leucine zipper family and regulate, besides CCT, enzymes of cholesterol, fatty acid, and TAG synthesis (Brown and Goldstein, 1997). The regulation of CCT by SREBP correlates with fatty acid synthesis and may occur through posttranscriptional activation of nuclear CCTol by fatty acids or a fatty acid-derived signal (Lagace et al., 2000). SREBPs are synthesized as precursors bound to the nuclear envelope and ER, and released from the membrane into the nucleus upon activation by cleavage. Interestingly, in flies, cleavage and subsequent release of SREBP from membranes depend on PtdEtn, the major phospholipid in Drosophila, exerting a feedback control on the synthesis of fatty acids and phospholipids (Dobrosotskaya et al., 2002). The finding that SREBP processing is controlled by different lipids in mammals and flies (sterols and PtdEtn, respectively) suggests that SREBP has an essential function in monitoring cell membrane composition and adjusting lipid synthesis accordingly. In plants, CCT expression increases after a shift to low temperatures (Choi et aL, 2001). In yeast, CCT activity becomes rate-limiting for PtdCho biosynthesis through the Kennedy pathway under conditions of choline limitation (McMaster and Bell, 1994). Yeast CCT is activated by anionic phospholipids. Kinetic characterization of purified yeast CCT demonstrated an increase in the kcat value, but not of Km values for the substrates CTP and phosphocholine upon lipid binding (Friesen et al., 2001b). In yeast, the contribution of the CDP-Cho as compared to the CDP-DAG pathway for PtdCho biosynthesis is regulated mainly by the availability of CTP for Cctlp, the rate-limiting enzyme of the Kennedy pathway. CTP synthase activity requires ATP/UTP-dependent tetramerization of the enzyme (Pappas et al., 1998),

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is inhibited by its product CTP, and is activated by phosphorylation (Ostrander et al., 1998). Overexpression of CTP synthase results in increased efficiency of the CDP-Cho at the expense of the CDP-DAG pathway (McDonough et al., 1995). The regulation includes supply of CTP for the phosphocholine cytidylyltransferase reaction in the CDP-Cho pathway and inhibition of PtdSer synthase in the CDPDAG pathway. In addition, Ckilp is positively regulated by ATP, but is inhibited by ADP and phosphorylation. Thus, the Kennedy pathway in yeast is sensitive to the energy state of the cell.

4. The Yeast INO-Regulation of Aminoglycerophospholipid Synthesis Transcriptional regulation of aminoglycerophospholipid biosynthesis has been extensively studied in the yeast S. cerevisiae (Carman and Henry, 1999). Genes encoding aminoglycerophospholipid biosynthetic enzymes, as well as many other phospholipid biosynthetic enzymes, are under coordinate transcriptional control of the inositol/choline response element (ICRE)/inositol-sensitive upstream activation sequence (UAS1/o) (5'-CATGTGAAAT-3/) in their promoters. These coordinately regulated enzymes have reduced activity levels in response to inositol, and even more dramatically in response to inositol plus choline, ethanolamine, or sefine during the exponential growth phase. Activity levels are respressed in the stationary growth phase independently of the presence of inositol and under nitrogen starvation (Griac and Henry, 1999). Among the enzymes affected are inositol-l-phosphate synthase (INO1), CDP-DAG synthase (CDS), PtdSer synthase (PSS1/CHO1), PtdSer decarboxylase 1 (PSD1), PtdEtn methyltransferases (PEM1 and PEM2), and choline kinase (CKII) (Carman and Henry, 1999). The yeast protein Opilp has been found to repress transcription of ICREregulated genes. A mutation in the OPI1 gene leads to overproduction of inositol (opi) (Carman and Henry, 1999). Tiffs opi-phenotype is also caused by mutations in CDS1, PSS1, PSD1, PEM1, PEM2, and URA7 (a CTP synthase mutant). Thus, ongoing PtdCho biosynthesis is required for global transcriptional regulation of phospholipid synthesis in response to inositol or nitrogen starvation. Increased turnover of PtdCho by PLD and decreased expression of CDP-DAG synthase lead to rapid derepression ofphospholipid biosynthetic genes, suggesting that accumulation of PA might be the signal for derepression of phospholipid biosynthetic genes. Under starvation, a low PA level might be the signal for the repression of phospholipid biosynthetic genes in the stationary phase regardless of inositol/chotine availability (Carman and Henry, 1999). Recently, links between the transcriptional regulation of phospholipid synthesis and the glucose and unfolded-protein response (UPR) pathways were established in yeast. Accumulation of unfolded protein in the ER initiates the UPR, which induces the transcriptional upregnlation of multiple ER-resident proteins involved in protein folding (Ma and Hendershot, 2001; Patil and Walter, 2001).

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Transcription of glucose-repressed genes under control of glucose depletion is facilitated via the glucose response signal transduction pathway (Carlson, 1999). Mutations in key proteins relaying either of the two signals to the nucleus were found to affect IN01 transcription, resulting either in inositol auxotrophy or in overproduction of inositol (opi) in the mutant cells (Cox et aL, 1997; Ouyang et aL, 1999; Shirra and Arndt, 1999). Thus, regulation of IN01 by UPR or glucose response pathways indirectly affects the transcriptional regulation of most glycerophospholipid biosynthetic genes by changing the cellular concentration of free inositoL

III. Lipid Dynamics and Membrane Assembly of Aminoglycerophospholipids A. Sorting of Aminoglycerophospholipid Species in Eukaryotic Cells Lipid analysis of subcellular fractions of rat liver, plant cells and yeast (Tables I-III) demonstrated that different organelles have different lipid compositions. The most striking differences are the high levels of PtdSer, sterols, and sphingolipids at the

TABLE I Lipicl Composition of Subcellular Fractions of Mammalian Cells (Rat Liver) Mitochondria a

Endoplasmic reticulurnb

Lysosomesc

Golgi b

Plasma membraneb

Phospholipid (mg/mg protein)

0.175

0.374

0.156

0.825

0.672

Sterol (rag/rag protein)

0.003

0.014

0.038

0.038

0.128

% of total phospholipids PtdCho

44

60

48

51

40

PtdEtn

34

23

17

21

24

Ptdlns

5

10

6

12

8

PtdSer

1

2

3

6

9

CL

14

1

1

1

1

PA

<1

1

1

<1

1

1

3

24

8

17

Sphingomyelin

Data are from: aZambrano et aL (1975); De Kroon et al. (1997). bZambrano et aL (1975). c Colbeau et al. (1971); Wherret and Huterer (1972).

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TABLE II Lipid Composition of Subcellular Fractions of Plant Cellsa Mitochondria (cauliflower)

Microsomes (castor bean)

Chloroplasts (cauliflower)

Peroxisomes (castor bean)

Tonoplast (mung bean)

Plasma membrane (mung bean)

Phospholipid (rag/rag protein)

0.356

0.930

0.046

0.488

1.545

0.983

Sterol (rag/rag protein)

0.016

0.008

--

0.002

0.420

0.460

% of total phospholipids PtdCho

44

47

55

54

47

47

PtdEtn

34

31

6

29

32

35 5

Ptdlns

7

14

14

10

11

PtdSer

0

2

0

2

5

8

PtdGro

4

4

23

3

5

5

11

2

2

2

0

0

CL

a Data are from Daum and Vance (1997) and references therein.

expense of PtdCho in the plasma membrane, and the high levels of CL and PtdEtn in mitochondria. It is obvious that cells must have developed elaborate mechanisms to maintain unique lipid compositions in the various organellar membranes. The characteristic lipid composition of organelles must be the result of a coordinate regulation and compartmentalization of lipid biosynthesis, degradation, and TABLE III Lipid Composition of Subcellular Fractions of Yeast (S. cerevisiae) a

Mitochondria

Endoplasmic reticulum

Vacuoles

Phospholipid (mg/mg protein)

0.09

0.22

0.51

0.23

Sphingolipids (mg/mg protein)

0.02

0.14

0.20

0.27

Sterol (mg/mg protein)

0.01

0.05

0.05

0.40

Plasma membrane

% of total phospholipids PtdCho

40

50

51

18

PtdEtn

27

24

21

22

Ptdlns

15

12

20

19

PtdSer

3

9

4

36

CL

13

1

2

<1

PA

2

4

2

4

aData are from Zinser and Daum (1995).

CELLBIOLOGYOFAMINOGLYCEROPHOSPHOLIPIDS

297

sorting. Some phospholipids are restricted to their organelle of synthesis, e.g., CL and PtdGro in mitochondria (see Tables I-III). Other glycerophospholipids, such as PtdCho, PtdEtn, Ptdlns, and PtdSer, are synthesized in the ER, Golgi, and mitochondria and are required in all organelles (Voelker, 1991; van Meer, 1993; Trotter and Voelker, 1994). Transport of lipids can be facilitated by vesicle flux (Puoti et al., 1991; Zinser et al., 1993), soluble carrier proteins (lipid transfer proteins) (Wirtz, 1991; Li et aL, 2000), or contact between donor and acceptor membranes (Daum and Vance, 1997; Voelker, 2000).

B. Compartmentalization of Aminoglycerophospholipid Biosynthetic Steps Aminoglycerophospholipid biosynthesis is a prominent example for compartmentalization of different biosynthetic steps of a pathway. Synthesis of PtdEtn and PtdCho requires crosstalk of mitochondria with the ER in mammalian and yeast ceils. Moreover, the Golgi/vacuole is involved in PtdEtn synthesis in yeast (Fig. 3) (Voelker, 2000). 1. The Route of Aminoglycerophospholipids from the ER to the Mitochondria

The substrate for the major pathway of mitochondrial PtdEtn biosynthesis in both mammals and yeast, PtdSer, is synthesized in the ER and transported to mitochondria for decarboxylation. The molecular mechanism of this interorganellar PtdSer transport is poorly understood. In isolated rat liver mitochondria, decarboxylation of PtdSer depends on substrate concentration, but not on cytosolic factors (Voelker, 1989). Similar studies in our laboratory using isolated yeast mitochondria (Simbeni et al., 1990, 1991, 1993) or permeabilized yeast cells (Acbleitner et al., 1995) revealed that import of PtdSer into mitochondria does not require ATP, an electrochemical gradient across the inner mitochondrial membrane, or cytosolic factors. Interestingly, mitochondria-associated membranes (MAM), ER-like membranes that are attached to mitochondria (Vance, 1990; Gaigg et al., 1995) and enriched in several lipid-biosynthetic activities, among them PtdSer synthase, appear to be involved in the supply of PtdSer to mitochondria. Although the function of MAM has not been firmly established, it is likely that import of PtdSer into mitochondria occurs through a membrane collision-based mechanism. In mammalian cells, MAM appear to be associated with contact sites between inner and outer mitochondrial membranes (Ardail et al., 1991, 1993). When PtdSer translocation to mitochondria or PtdSer decarboxylation is blocked, newly synthesized PtdSer accumulates in MAM but not in bulk microsomes, suggesting that PtdSer traverses the MAM on its route from the ER to mitochondria (Vance and Shiao, 1996). Respiratory activity of mitochondria may modulate the association of MAM and mitochondria in mammalian cells (Monni et al., 2000).

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Proteins of the outer mitochondrial membrane appear to be required for efficient PtdSer import in a reconstituted system of isolated membranes from CHO cells (Shiao etaL, 1998) or yeast (Achleitner etaL, 1999). A fusogenic protein associated with mitochondrial membranes was implicated in import of PtdSer into rat brain mitochondria (Camici and Corazzi, 1997). In permeabilized CHO cells, a cytosolic factor from bovine brain, the EF-hand-type calcium-binding protein S 100B, was shown to enhance transport-dependent PtdSer decarboxylation in the presence of Ca 2+ (Kuge et al., 2001). In the course of these studies, a mammalian mutant defective in intramitochondrial PtdSer transport was isolated (Emoto et al., 1999), but the gene affected has not yet been identified. In mammalian cells, the preference of PSD for certain PtdSer species in vitro does not reflect the mitochondrial PtdEtn profile, suggesting that remodeling or transport selectivity may be important (Kevala and Kim, 2001). Indeed, hydrophilic PtdSer species are translocated to mitochondria more efficiently than hydrophobic species. Thus, efflux of PtdSer from the ER membrane may be the rate-limiting step of PtdSer translocation to mitochondria (Heikinheirno and Somerharju, 1998, 2002). Recent work in our laboratory with yeast confirmed this view (Biirgermeister et al., unpublished). PtdSer synthesis was shown to be the driving force for mitochondrial PtdEtn formation in yeast rather than its transport in mitochondria (Birner et aL, unpublished). Experimental evidence for transport of PtdEtn and PtdCho from the ER to mitochondria is limited. Such experiments are difficult to perform because the import of PtdEtn and PtdCho, in contrast to PtdSer translocation to mitochondria, is not linked to metabolic conversion. In rat brain, extramitochondrially synthesized PtdEtn is mainly assembled into the outer mitochondrial membrane, while PtdEtn produced by mitochondrial decarboxylation is mainly confined to the inner mitochondrial membrane (Camici and Corazzi, 1995). In vitro translocation of PtdEtn from isolated ER to mitochondria is enhanced by Ca 2+ or addition of a nonspecific lipid-transfer protein. Alternatively, PtdEtn synthesized in MAM can be translocater to mitochondria in the reconstituted system without the aid of a transfer protein. A high rate of PtdCho transport from the ER to mitochondria in vivo was demonstrated in pnlse-chase experiments with yeast (Daum et aL, 1986). This transport does not distinguish between PtdCho synthesized by the CDP-Cho pathway or the PtdEtn methylation pathway (Janssen et al., 1999). Transmembrane movement of PtdCho across the outer mitochondrial membrane of yeast and liver is rapid, bidirectional, and independent of ATP and membrane potential (Lampl et al., 1994; Dolls et al., 1996). 2. The Route of Phosphatidylserine from the ER to the Golgi/Vacuole in Yeast Genetic screens to isolate yeast strains defective in interorganellar PtdSer transport revealed mutants with a defect in PtdS er transport from the ER to the Golgi/vacuole,

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which harbors Psd2p (Fig. 3) (Trotter et al., 1998). Psd2p contains a C2 homology domain that is not required for catalysis in vitro but is essential for in vivo function, suggesting that this domain plays a direct role in membrane docking and/or PtdSer transport to the enzyme (Kitamura et al., 2002). Identification of the SEC14 homolog PDR17 (van den Hazel et aL, 1999; Wu et al., 2000) and the essential PtdIns-4-kinase STT4 (Trotter et al., 1998) as components involved in this translocation process suggested vesicular flux as a possible mechanism. In contrast, the more recent observation that PtdSer transport from the ER to Golgi/vacuoles does not require ATP indicates that this route may also be nonvesicular (Wu and Voelker, 2001). During these screens, a mutation in the morphogenesis checkpointdependent M C D 4 gene, encoding a protein implicated in GPI-anchor biosynthesis, was identified (Storey et al., 2001b). The function of Mcd4p appears to be transfer of phosphoethanolamine from PtdEtn to the first mannose of the GPI anchor (Gaynor et al., 1999; Hong et al., 1999). Since the mutant M C D 4 allele did not cause a defect in GPI-anchor protein synthesis and did not accelerate PtdEtn degradation, transport-dependent metabolism of PtdSer may be affected (Storey et al., 2001b). 3. Vesicular Transport of Aminoglycerophospholipids to the Plasma Membrane Vesicular transport governing protein trafficking to the plasma membrane also appears to account for phospholipid transport to secretory organelles, namely the Golgi, the vacuole, and the plasma membrane, as demonstrated by lipid analysis of isolated yeast compartments (Zinser et al., 1991; Schneiter et al., 1999). The accumulation of PtdSer at the expense of PtdCho in the plasma membrane may be the result of specific sorting of PtdSer, together with sterols, sphingolipids, and GPI-anchored or acylated proteins, in lipid rafts. As another example, vesicles budding from plant ER are enriched in PtdSer, especially in species containing very long-chain fatty acids (Sturbois-Balcerzak et al., 1999; Vincent et al., 2001). Recent studies on intracellular transport of pulmonary surfactant PtdCho in alveolar type II cells, however, showed that transport of PtdCho along the secretory pathway does not require a functional Golgi apparatus (Osanai et al., 2001). In yeast, a subfraction of the ER with a high capacity to synthesize lipids was found to associate with the plasma membrane and may be involved in nonvesicular transport of lipids to the cell periphery (Pichler et al., 2001). Nonvesicular transport of fluorescent PtdCho analogs significantly contributes to the canalicular enrichment of PtdCho in hepatocytic cells (Wustner et al., 2001). Earlier studies with fluorescent analogs of phospholipids showed that fluorescent PtdSer analog is transported to the Golgi and to mitochondria by nonvesicular mechanisms in fibroblasts (Kobayashi and Arakawa, 1991). A fluorescent PtdCho analog is sorted by vesicular transport to the yeast vacuole for degradation, whereas a fluorescent PtdEtn analog is internalized to the nuclear envelope/ER and mitochondria, most likely independently from vesicular transport (Grant et al., 2001).

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C. Maintenance of Plasma Membrane Asymmetry

The asymmetric phospholipid composition of the two leaflets of a membrane bilayer appears to be maintained by the membrane-associated cytoskeleton and the action of phospholipid-transporting proteins (Gupta, 1990). Transport of phospholipids across a phospholipid bilayer may be facilitated by flip and flop. Flip is defined as the transport of a phospholipid molecule from the outer to the inner leaflet, and flop is the transport in the opposite direction. In E. coli, all lipid-synthesizing enzymes are localized to the cytosolic side of the inner membrane. Newly synthesized PtdEtn is translocated rapidly and independently from energy across the inner membrane (Huijbregts et al., 1998). Membrane-spanning peptides have been found to induce phospholipid flop across the inner membrane of E. coli (Kol et al., 2001). In yeast, the distribution of PtdEtn in the plasma membrane is regulated by an ATP-dependent, protein-mediated process (Balasubramanian and Gupta, 1996; Kean etal., 1997; Grant etal., 2001), which is significantly influenced by disrupting the actin cytoskeleton (Dixit and Gupta, 1998). Earlier studies with blood cells suggested that PtdEtn and PtdSer in the inner leaflet of the membrane may be stabilized by their interactions with cytoskeletal proteins (Williamson et aL, 1987; Gupta, 1990; Schroit and Zwaal, 1991; Williamson and Schlegel, 1994). Phospholipid translocation across the plasma membrane of mammalian cells has been reviewed by Bevers et aL (1999). The aminophospholipid translocase (flippase) is one of the major factors governing the asymmetric distribution of PtdSer and PtdEtn in the plasma membrane by flipping PtdSer and PtdEtn from the outer to the inner leaflet (Daleke and Lyles, 2000). A putative aminophospholipid translocase has been identified in bovine cells (ATPaselI) (Tang et aL, 1996; Ding et aL, 2000). This polypeptide belongs to a new subfamily of P-type ATPases which includes the human FIC-1 gene, the yeast DRS2 gene (Siegmund et al., 1998), and theArabidopsisALA1 gene (Gomes et al., 2000). However, the yeast drs2A mutant is not defective in transbilayer movement of fluorescent phospholipid analogs in the plasma membrane (Siegmund et aL, 1998; Marx et al., 1999), but is synthetically lethal with clathrin heavy-chain temperature-sensitive alleles. The drs2A mutant exhibits late Golgi defects, which may result from loss of clathrin function at this compartment, suggesting a possible link between membrane asymmetry and clathrin function at the Golgi complex (Chen et al., 1999). Deletion of the yeast DRS2 gene leads to a cold-sensitive growth phenotype and a defect in internalization of PtdSer; both defects are complemented by heterologous expression of ALA1 (Gomes et al., 2000). Downregulation of ALA1 in Arabidopsis results in cold-sensitive plants that are much smaller than the wild-type, suggesting a link between transmembrane bilayer asymmetry and cold adaptation. Selective degradation and synthesis of aminoglycerophospholipids in the two leaflets of the plasma membrane by deacylation/acylation reactions, called membrane polishing, is also involved in their transbilayer distribution (Rice, 1998).

302

BIRNERAND DAUM

Secreted PLA2 isoenzymes with specificity for aminophospholipids may preferentially remove PtdSer and PtdEtn from the exoplasmic leaflet of the plasma membrane. In bovine erythrocytes, the asymmetry of PtdEtn in the plasma membrane appears to be maintained by deacylation of PtdEtn at the membrane surface and selective reacylation of lyso-PtdEtn in the inner leaflet (Florin-Christensen et aL, 2001). Putative floppases (outwardly directed aminophospholipid translocases) were identified as ATP-binding cassette (ABC) transporters (Borst et al., 2000; Raggers et aL, 2000). ABC transporters are active transporters that bind their substrates and translocate it through the membrane in an ATP-dependent manner against a concentration gradient. The typical ABC transporter is a large protein with 12 transmembrane segments, two nucleotide binding sites, and modified with carbohydrate side chains. P-Glycoproteins encoded by the multidrug-resistance M D R 3 gene in humans and the M D R 2 gene in mice are primarily PtdCho transporters. An m d r 2 A knockout mouse is completely unable to secrete PtdCho into bile (Frijters et al., 1997). While the human MDR3 protein has specific PtdCho translocase activity, other ABC transporters, such as MDR1 P-glycoprotein and the multidrug-resistance (-associated) transporter MRP1, have a broad specificity for lipid analogs (Romsicki and Sharom, 2001) which may, however, be recognized rather as drugs than lipids. MDR3 also has significant drug transport activity (Smith et al., 2000). MDR1 appears to be involved in secretion of the platelet-activating factor, which is the natural short-chain phospholipid 1-O-alkyl2-acetyl-sn-glycero-3-phosphocholine (Raggers et al., 2001). ABCR (also called Rim protein) is an ABC transporter found exclusively in vertebrate photoreceptor outer segments. This polypeptide, whose defect causes Stargardt's macular dystrophy, may transport a complex of retinaldehyde and PtdEtn to the retina of the eye (Weng et al., 1999; Sun and Nathans, 2001). The ATP-binding cassette transporter 1 (ABC1) mediates release of membrane phospholipids and cholesterol to apo-AI of the cholesterol-shuttling high-density lipoprotein (HDL) particle and promotes Ca2+-induced exposure of PtdSer at the membrane (Hamon et al., 2000). Gain-of-function mutations in the transcription factors PDR1 and PDR3 (pleiotropic drug resistance), which are known to regulate transcription of ATPbinding cassette transporters in yeast, result in a dramatic upregulation of flop but also in downregulation of flip of fluorescent phospholipid analogs across the plasma membrane (Hanson and Nichols, 2001). Recently, three ABC transporters of Candida albicans, Cdrlp, Cdr2p, and Cdr3p, were shown to translocate phospholipid analogs (Smriti et aL, 2002). Cell activation, apoptosis, and injury are accompanied by random redistribution (called scrambling) of all major lipid classes between the leaflets of the plasma membrane, resulting in surface exposure of PtdSer and PtdEtn (see Section lI.B.3.). A protein called phospholipid scramblase (PLSCR1) exhibiting Ca 2+- or apoptosis-activated phospholipid scrambling activity has been cloned in human and mouse (Zhou et al., 1997). Recently, three human homologs of

CELLBIOLOGYOFAMINOGLYCEROPHOSPHOLIPIDS

303

PLSCR1 were identified (Wiedmer et al., 2000; Sims and Wiedmer, 2001). Activation of the same scramblase by Ca 2+ or apoptosis occurs through different signal transduction pathways in lymphocytes (Williamson et al., 2001). The PLSCR1 expression level regulates movement of PtdSer to the cell surface (Zhao et aL, 1998), and PLSCR1 upregulation in tumor cells exposed to interferon (Zhou et al., 2000) may contribute to the tumor-suppressive action of this cytokine (Silverman et al., 2002). The number of blood cells was normal in adult p l s c r l knockout mice, but depressed in fetus and newborn animals (Zhou et al., 2002). Hematopoietic precursor cells showed impaired differentiation to mature granulocytes. These results suggest that PLSCR 1 functionally contributes to cytokine-regulated cell proliferation and differentiation. It has been suggested that specific aminophospholipid flippases may also exist in ER membranes (Menon et aL, 2000; Nicolson and Mayinger, 2000). Membrane asymmetry, with PtdSer predominantly present in the inner and PtdEtn in the outer leaflet but with PtdCho equally distributed between both monolayers, has been demonstrated for the sarcoplasmic reticulum (Bick et al., 1998).

IV. Concluding Remarks During the last twenty years, significant progress has been made in the identification, characterization, and localization of enzymes of the aminoglycerophospholipid biosynthetic pathways in different cell types. Many genes encoding the respective proteins were identified. However, important questions concerning the overall organization of organellar membranes, the homeostasis of lipids, and the regulatory network of membrane component biosynthesis remain to be answered and are highly _n........mg fie~u~for future studies. Different subcellular membranes have specific lipid compositions. It is still a matter of dispute how the characteristic distribution of phospholipids is maintained. Highly specific sorting mechanisms and tightly regulated local phospholipid synthesis and turnover must have evolved. Data available so far suggest a major contribution of vesicular transport to phospholipid transport from the major site of phospholipid biosynthesis, the ER, to organelles of the secretory pathway including the plasma membrane. However, alternative mechanisms of transport, such as monomer transfer with or without the aid of transfer proteins or membrane contact between donor and acceptor organelles, appear to exist. Such mechanisms were suggested for organelles outside the secretory pathway, especially for mitochondria. The route of mitochondrial decarboxylation of PtdSer synthesized in the ER is a prominent example for the interplay of different cellular compartments for biosynthesis of aminoglycerophospholipids. Ongoing efforts to identify factors involved in aminoglycerophospholipid transport will help to clarify the mechanism(s) involved.

304

BIRNERAND DAUM

How is the asymmetry of cellular membranes maintained? Obviously, transporters, but also the interaction of the cytoskeleton with anionic phospholipid species such as PtdSer, play important roles in maintaining the asymmetry of the plasma membrane in mammalian cells. Whether this is also true for intracellular membranes remains to be established. What is the function of different phospholipid classes/species in different organellar membranes? How do they influence the properties of the membrane? Certain phospholipid species may specifically interact with proteins involved in all kinds of cellular processes, thus affecting protein conformation and enzymatic activity. PtdSer and PtdEtn exposed on the mammalian cell surface are important factors for intracellular communication and cytokinesis. PtdEtn synthesized in mitochondria appears to play a major role at the site of its synthesis. In all cases, species analysis will shed more light on specific properties of these lipids. The use of modern methods of mass spectrometry will provide the analytical basis to establish a cellular "lipidome," which will help us understand the molecular background of membrane properties. Finally, regulation of membrane homeostasis in different cell types needs more elucidation. Obviously, phospholipid synthesis and turnover are tightly and coordinately regulated. While sterols and fatty acids have been identified as membrane sensors in mammalian cells, PtdEtn serves as a membrane sensor in flies. The membrane sensor in yeast has not yet been identified. Recently, links between phospholipid regulation and the glucose response and unfolded protein response in yeast were demonstrated, suggesting that an overall regulatory network exists to coordinate protein and lipid synthesis in response to nutrients and stress. However, the molecular basis of regulation of phospholipid biosynthesis in the cellular regulatory network remains to be investigated.

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Bick, R. J., Buja, L. M., Van Winkle, W. B., and Taffet, G. E. (1998). Membrane asymmetry in isolated canine cardiac sarcoplasmic reticnlum: Comparison with skeletal muscle sarcoplasmic reticulum. J. Membr. BioL 164, 169-175. Birner, R., Biirgermeister, M., Schneiter, R., and Danm, G. (2001). Roles of phosphatidylethanolamine and of its several biosynthetic pathways in Saccharomyces cerevisiae. Mol. Biol. Cell 12, 997-1007. Bladergroen, B. A., and van Golde, L. M. (1997). CTP:phosphoethanolamine cytidylyltransferase. Biochim. Biophys. Acta 1348, 91-99. Bladergroen, B. A., Geelen, M. J., Reddy, A. C., Declercq, P. E., and Van Golde, L. M. (1998). Channelling of intermediates in the biosynthesis of phosphatidylcholine and phosphatidylethanolamine in mammalian cells. Biochem. J. 334, 511-517. Bladergroen, B. A., Bussiere, M., Klein, W., Geelen, M. J., Van Golde, L. M., and Houweling, M. (1999). Inhibition of phosphatidylcholine and phosphatidylethanolamine biosynthesis in rat-2 fibroblasts by cell-permeable ceramides. Eur. J. Biochem. 264, 152-160. Bogdanov, M., and Dowhan, W. (1998). Phospholipid-assisted protein folding: Phosphatidylethanolamine is required at a late step of the conformational maturation of the polytopic membrane protein lactose permease. EMBO J. 17, 5255-5264. Bogdanov, M., Umeda, M., and Dowhan, W. (1999). Phospholipid-assisted refolding of an integral membrane protein. Minimum structural features for phosphatidylethanolamineto act as a molecular chaperone. J. Biol. Chem. 274, 12339-12345. Bogdanov, M., Heacock, P. N., and Dowhan, W. (2002). A polytopic membrane protein displays a reversible topology dependent on membrane lipid composition. EMBO J. 21, 2107-2116. Bolognese, C. P., and McGraw, P. (2000). The isolation and characterization in yeast of a gene for Arabidopsis S-adenosylmethionine:phosphoethanolamineN-methyltransferase. Plant Physiol. 124, 1800-1813. Bontemps, E, and van den Berghe, G. (1998). Novel evidence for an ecto-phospholipid methyltransferase in isolated rat hepatocytes. Biochem. J. 330, 1-4. Brown, M. S., and Goldstein, J. L. (1997). The SREBP pathway: Regulation of cholesterol metabolism by proteolysis of a membrane-bound transcription factor. Cell 89, 331-340. Borst, P., Evers, R., Kool, M., and Wijnholds, J. (2000). A family of drug transporters: The multidrug resistance-associated proteins. J. Natl. Cancer Inst. 92, 1295-1302. Camici, O., and Corazzi, L. (1995). Import of phosphatidylethanolamine for the assembly of rat brain mitochondrial membranes. J. Membr. Biol. 148, 169-176. Camici. 0.. and Corazzi, L. (1997). Pbosphatidylserine translocation into brain mitochondria: Involvement of a fusogenic protein associated with mitochondrial membranes. Mol. CeIL Biochem. 175, 71-80. Campbell, H. A., and Kent, C. (2001). The CTP:phosphocholine cytidylyltransferase encoded by the licC gene of Streptococcus pneumoniae: Cloning, expression, purification, and characterization. Biochim. Biophys. Acta 1534, 85-95. Carlson, M. (1999). Glucose repression in yeast. Curr. Opin. Microbiol. 2, 202-207. Carman, G. M., and Henry, S. A. (1999). Phospholipid biosynthesis in the yeast Saccharomyces cerevisiae and interrelationship with other metabolic processes. Prog. Lipid Res. 38, 361-399. Chapman, K. D. (2000). Emerging physiological roles for N-acylphosphatidylethanolaminemetabolism in plants: Signal transduction and membrane protection. Chem. Phys. Lipids 108, 221-229. Chen, C. Y., Ingram, M. E, Rosal, P. H., and Graham, T. R. (1999). Role for Drs2p, a P-type ATPase and potential aminophospholipid translocase, in yeast late Golgi function. J. Cell. Biol. 147, 1223-1236. Choi, Y. H., Lee, J. K., and Cho, S. H. (2001). Structure and expression of a CTP:phosphocholine cytidylyltransferase gene from Arabidopsis thaliana. Mol. Cells 11, 95-99. Cleves, A. E., McGee, T. P., Whitters, E. A., Champion, K. M., Aitken, J. R., Dowhan, W., Goebl, M., and Bankaitis, V. A. (1991). Mutations in the CDP-choline pathway for phospholipid biosynthesis bypass the requirement for an essential phospholipid transfer protein. Cell 64, 789-800.

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