A role for lipid trafficking in chloroplast biogenesis

Progress in Lipid Research 47 (2008) 381–389

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Review

A role for lipid trafficking in chloroplast biogenesis Christoph Benning * Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48823-1319, United States

a r t i c l e

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Keywords: ABC transporter Fatty acid Flippase Glycerolipid Phospholipid Photosynthetic membrane

a b s t r a c t Chloroplasts are the defining plant organelle carrying out photosynthesis. Photosynthetic complexes are embedded into the thylakoid membrane which forms an intricate system of membrane lamellae and cisternae. The chloroplast boundary consists of two envelope membranes controlling the exchange of metabolites between the plastid and the extraplastidic compartments of the cell. The plastid internal matrix (stroma) is the primary location for fatty acid biosynthesis in plants. Fatty acids can be assembled into glycerolipids at the envelope membranes of plastids or they can be exported and assembled into lipids at the endoplasmic reticulum (ER) to provide building blocks for extraplastidic membranes. Some of these glycerolipids, assembled at the ER, return to the plastid where they are remodeled into the plastid typical glycerolipids. As a result of this cooperation of different subcellular membrane systems, a rich complement of lipid trafficking phenomena contributes to the biogenesis of chloroplasts. Considerable progress has been made in recent years towards a better mechanistic understanding of lipid transport across plastid envelopes. Lipid transporters of bacteria and plants have been discovered and their study begins to provide detailed mechanistic insights into lipid trafficking phenomena relevant to chloroplast biogenesis. Ó 2008 Elsevier Ltd. All rights reserved.

Contents 1. 2. 3.

4. 5. 6.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Interaction of the ER and chloroplast during plastid development. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Basic polar lipid transfer mechanisms relevant to ER-plastid lipid transfer. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Transmembrane transport and leaflet-to-leaflet flipping of polar lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Intermembrane transport of polar lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Vesicular lipid transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatty acid export from plastids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genetic analysis of ER-to-plastid lipid trafficking in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Plants carry out photosynthesis which provides the oxygen in the atmosphere and chemical energy to heterotrophic organisms in the form of reduced carbon. Photosynthesis is essential to most modern life forms, perhaps with the exception of organisms living in extreme environments, e.g. in the vicinity of deep sea hydrothermal vents. The conversion of sun light into chemical energy by photosynthesis requires an extensive membrane system (thylakoids), which in * Tel.: +1 517 355 1609; fax: +1 517 353 9334. E-mail address: [email protected]. 0163-7827/$ - see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.plipres.2008.04.001

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plants is localized in chloroplasts. Photosynthetic protein pigment complexes are embedded into a unique matrix of polar glycerolipids consisting of the non-phosphorous glycoglycerolipids, mono- and digalactosyldiacylglycerol (MGDG and DGDG) and sulfoquinovosyldiacylglycerol (SQDG), and the phospholipid phosphatidylglycerol (PtdGro). The two galactoglycerolipids are predominant in chloroplasts [1,2] and for simplicity much of this review focuses on these two lipids. The two monohexosyldiacylglycerols MGDG and SQDG are exclusively located in plastid membranes, while DGDG and PtdGro can also be found in extraplastidic membranes. The chloroplast is enclosed by two envelope membranes. The outer contains in addition to the already mentioned glycerolipids

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phosphatidylcholine (PtdCho) [3]. Extraplastidic membranes, e.g. the endoplasmic reticulum (ER), contain specific glycerolipids such as phosphatidylethanolamine (PtdEtn) that are not found in the plastid. Overall, there is a great diversity of lipids distinguishing the different subcellular membranes of a plant [4–6]. While we have a basic understanding of the biochemical reactions of glycerolipid biosynthesis in plants [7–11], we still do not understand the mechanisms and regulatory processes on which this lipid diversity of subcellular membranes is based. Moreover, lipid biosynthetic enzymes are not necessarily located in the membrane in which a specific lipid resides and, therefore, lipid trafficking is essential for the biogenesis of organelles. As an example, in plants fatty acids (FAs) found in membrane lipids of all membranes or in storage lipids, which are assembled at the ER, are nearly exclusively synthesized in plastids [12]. As a result of this organization of lipid metabolism in plants, FAs have to be exported from plastids. However, the basic mechanisms by which fatty acids leave the plastid are not understood. As will be discussed in detail below, the ER and the plastid cooperate in the assembly of the thylakoid membrane lipids, which requires the extensive exchange of lipid precursors. In addition, it was recently discovered that the glycoglycerolipid DGDG, which is assembled at the outer plastid envelope membrane, can be exported to extraplastidic membranes following phosphate deprivation [13]. Thus, the plastid exchanges complex lipids in addition to FAs with extraplastidic membranes. As the biochemical power house of the plant cell, plastids continuously import or export many different biochemical precursors and intermediates, or products in addition to lipids [14]. However, this review focuses on lipid trafficking phenomena involved in plastid development. For additional information the reader is also referred to recent reviews on this topic [5,15,16]. 2. Interaction of the ER and chloroplast during plastid development Two parallel lipid assembly pathways contribute to plastid membrane biogenesis in many plants, including the genetic model plant Arabidopsis, on which much of this review focuses. The current hypothesis of galactoglycerolipid biosynthesis in Arabidopsis is shown in Fig. 1. Following the synthesis of FAs, a portion is incorporated directly into glycerolipids catalyzed by plastid acyl-acyl carrier protein (ACP): glycerol 3-phosphate acyltransferase en-

coded by ATS1 [17,18] and plastid acyl-ACP: lyso-phosphatidic acid (-PtdOH) acyltransferase encoded by ATS2 [19,20]. The PtdOH generated in the plastid is dephosphorylated to diacylglycerol (DAG). A candidate gene of Arabidopsis encoding the plastid PtdOH phosphatase of leaves was recently described [21], but definitive proof that this enzyme is involved is still pending. The produced DAG serves along with UDP-galactose as substrate for the formation of MGDG by MGDG synthase. The enzyme responsible for the bulk of MGDG biosynthesis in Arabidopsis is encoded by MGD1 [22,23]. Galactosylation of MGDG by DGD1 using UDP-galactose as substrate leads to the formation of DGDG under normal growth conditions [24,25]. In Arabidopsis, MGD1 is associated with the outer face of the inner envelope membrane [22,26], while DGD1 is inserted into the outer envelope membrane but exposed to the cytosol [27]. The sequence of events described so far is the plastid pathway of galactoglycerolipid biosynthesis. Alternatively, FAs are exported from the plastid and are assembled into glycerolipids at the ER by ER-specific acyltransferase isoforms. Recently, the classic assumption that nascent FAs following their export are first incorporated into PtdOH has been challenged. Rapid pulse-chase labeling experiments with pea or rape seed leaves have shown that newly exported fatty acids first appear in PtdCho presumably due to a very active acyl editing mechanism [28,29]. Apparently, de novo PtdCho biosynthesis occurs from glycerol 3-phosphate and recycled acyl groups at the ER [28]. It is not clear whether the proposed acyl editing happens at the ER or the outer plastid envelope where acyl editing could be part of the FA export mechanism. In any case, the first glycerolipid formed outside the plastid from newly synthesized fatty acids is PtdCho. The DAG moieties of PtdCho find their way back into the plastid by mechanisms discussed below, where they are incorporated into galactoglycerolipids by MGD1 and DGD1. This second series of events constitutes the ER pathway of galactoglycerolipid biosynthesis. Thus the origin of the DAG moieties differs in the two pathways, but not the final assembly of galactoglycerolipids by MGD1 and DGD1. In Arabidopsis, approximately equal fluxes through both pathway contribute to galactoglycerolipid biosynthesis [30]. The nature of the lipid transferred from the ER to the outer envelope as part of the ER pathway of galactoglycerolipid biosynthesis is still unclear. The current working hypothesis (Fig. 1) does not distinguish between direct transfer of PtdOH from the ER, or the transfer of PtdCho and its conversion to PtdOH by a cytosolic phospholipase D recruited to the outer plastid envelope mem-

Fig. 1. Assembly of galactoglycerolipids and PtdGro from ER- and plastid-derived precursors. Membranes: ER, endoplasmic reticulum; iE, inner plastid envelope; oE, outer plastid envelope; Thy, thylakoid membrane. Proteins: ATS1, plastid acyl-ACP: glycerol 3-phosphate acyltransferase; CDS CDP-diacylglycerol synthetase; DGD1, digalactosyldiacylglycerol synthase; FAS, fatty acid synthase complex; MGD1, monogalactosyldiacylglycerol synthase; PAP, plastid phosphatidic acid phosphatase; PGP1, phosphatidylglycerol phosphate synthase; PGPP, phosphatidylglycerol phosphate phosphatase; TGD1, 2, 3, components of the proposed phosphatidic acid transporter. Lipids: CDAG, CDP-diacylglycerol; DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; FA, fatty acid; MGDG, monogalactosyldiacylglycerol; PA, phosphatidic acid; PC, phosphatidylcholine; PG, phosphatidylglycerol; PGP, phosphatidylglycerol 3-phosphate. Arrows indicate movement of lipids. A simplified scheme is shown indicating only lipids and lipid precursors.

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Fig. 2. Pulse-chase labeling of monogalactosyldiacylglycerol (MGDG, solid line) and phosphatidylcholine (PtdCho, dashed line) by application of [14C]-acetate to Arabidopsis leaves. Wild-type, tgd1 mutant, and ats1 mutant results are shown from left to right. The two components of the time courses indicated in the wild-type panel are discussed in the text. The data were originally published in [17,36] and the panels were redrawn from the original data.

brane. An involvement of cytosolic lipases was concluded based on the observation that the conversion of labeled PtdCho into galactoglycerolipids in isolated chloroplasts is stimulated by cytosolic fractions [31]. Because the plastid envelope is widely believed to lack the capability to synthesize PtdCho [10] the plastid needs to import PtdCho from the ER, or lyso-PtdCho as previously suggested [32]. However, this does not necessarily mean that these two lipids after they arrived at the outer envelope membrane will also give rise to the ER-derived DAG moieties found in galactoglycerolipids. ER-to-plastid lipid trafficking is a dynamic process. The distinct subcellular localization and membrane association of the proteins involved in galactoglycerolipid biosynthesis as summarized above (Fig. 1) mandates the movement of lipid precursors between the different biosynthetic enzymes. Classic pulse-chase labeling experiments permit a time-resolved analysis of different lipid pools in intact leaves and directly convey the dynamics of lipid movement between the organelles. In combination with genetic mutants inactivated in specific proteins involved in the process, pulse-chase experiments reveal the dynamic nature of ER-to-plastid lipid trafficking. Indeed, the two-pathway hypothesis for thylakoid lipid assembly was first postulated by Roughan et al. [33] following a series of classic labeling experiment during the 1970s and 1980s as summarized by Frentzen [34] and Roughan and Slack [35]. This approach is illustrated in Fig. 2. The precursor [14C]-acetate is readily taken up when applied to intact leaves and is rapidly incorporated into FAs in the plastid. During the ‘‘cold” chase, which is simply due to the photosynthetic fixation of atmospheric carbon dioxide and incorporation into FAs during continuing growth of the leaf, a biphasic time course of label abundance in MGDG is observed (Fig. 2). A more complex time course is observed for PtdCho. Recall that PtdCho is assembled at the ER but not at the plastid envelope membranes, although it is present in the outer envelope membrane. Furthermore, MGDG is exclusively present in the plastid where it is also assembled. The currently accepted explanation for the biphasic labeling time course of MGDG is that acetate enters the plastid and is converted to acyl-ACPs, which are immediately incorporated into MGDG by the plastid pathway representing component 1 evident in the wild-type time course (Fig. 2). As FAs are exported from the plastid, PtdCho is labeled with a delay. During the chase the rapidly labeled MGDG is converted in part to DGDG decreasing the label in the MGDG pool. Label also rapidly passes through the PtdCho pool as this lipid is turned over and its DAG moiety appears in other glycerolipids in extraplastidic membranes (explaining its more complex labeling time course), but also in MGDG present in plastid membranes. As a consequence, label in the MGDG pool increases during the second phase of the time course (Fig. 2), which is interpreted as the ER pathway of galactoglycerolipid biosynthesis.

Mutants of Arabidopsis are available that lack either of the two components apparent during the pulse-chase labeling analysis of wild-type leaves. The tgd1 mutant of Arabidopsis is deficient in the permease component of a lipid transporter of the inner chloroplast envelope that will be discussed further below [26,36]. This mutant only shows component 1 of MGDG labeling with rapid and increased overall labeling of MGDG (Fig. 2). According to the interpretation for the wild-type given above, MGDG in this mutant is primarily made by the plastid pathway consistent with a disruption of the ER pathway. The ats1 mutant of Arabidopsis (formerly designated act1) is deficient in the plastid glycerol 3-phosphate: acyl-ACP acyltransferase [17,18]. It is missing component 1 of the labeling time course (Fig. 2) consistent with a disruption of the plastid pathway of galactoglycerolipid biosynthesis. It makes MGDG exclusively from ER-derived precursors. The contribution of the two pathways in different plants can be determined based on the specific molecular species composition of lipids derived from either pathway. Indeed, the biochemical analysis of Arabidopsis mutants, which are altered in FA composition of their glycerolipids and many of which are deficient in specific FA desaturases in the ER and the plastid [9,37,38], has elegantly corroborated the two-pathway hypothesis for thylakoid lipid biosynthesis outlined above. Lipids derived from the plastid pathway contain 16-carbon fatty acids in the sn-2 position of the DAG backbone, and lipids derived from the ER pathway have an 18-carbon fatty acid in this position [39]. This difference is due to distinct substrate specificities of the acyltransferase or acyl exchange enzymes in the ER or the plastid outer envelope, and the plastid inner envelope. Because the FA composition of plastid pathway-derived thylakoid lipids resembles that of cyanobacteria [40], which are considered the ancestral precursors of plastids, the plastid pathway is often also referred to as the ‘‘prokaryotic pathway”. In contrast, the ER pathway of thylakoid lipid biosynthesis is referred to as ‘‘eukaryotic” pathway. Many seed plants are known that lack the plastid pathway and exclusively rely on the ER pathway for galactoglycerolipid biosynthesis [41] requiring extensive lipid trafficking between the ER and the plastid. On the other hand, the unicellular alga Chlamydomonas reinhardtii appears to lack the ER pathway for galactoglycerolipid biosynthesis [42]. It should be noted that this organisms also lacks PtdCho, which might implicate this lipid in the mechanism of lipid transfer from the ER to the plastid.

3. Basic polar lipid transfer mechanisms relevant to ER-plastid lipid transfer Consistent with the hypothesis that plastids originated from ancestral endosymbiotic Gram-negative photosynthetic bacteria

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Fig. 3. Postulated lipid transfer processes involved in ER-plastid lipid trafficking in plants. The endoplasmic reticulum (ER) and the outer (oE) and inner envelope membranes (iE) of the plastid are depicted. Bilayer membranes with their two leaflets are drawn. Arrows indicate different lipid transfer processes: (1) polar lipid flipping across the outer envelope membrane; (2) polar lipid flipping across the inner envelope membrane; (3) transmembrane movement of polar lipids through the proposed TGD123 complex; (4) transfer of lipids between the two envelope membranes; (5) lipid transfer mediated by lipid transfer proteins; (6) lipid transfer through ER-outer envelope membrane contact zones (PLAMs); and (7) vesicle formation at the inner envelope membrane. Only proteins of known identity that were implicated in plants are shown: TGD1, 2, 3; LTP, general lipid transfer protein; VIPP1 as discussed in the text.

[43], remnants of the bacterial lipid transfer machineries might still be present in plastids and plant orthologs might have been recruited for the transport of lipids during the assembly of the photosynthetic membrane. This assumption is certainly supported by our increasing understanding of the bacterial origin of the chloroplast protein import apparatus showing that many cyanobacterial components in their cell membranes have orthologs in plastids [44,45]. Whenever appropriate, potential parallels will be drawn between less well studied plant phenomena and well characterized related processes in bacteria and yeast to stimulate the thinking about lipid trafficking in plants. The complexity of photosynthetic membrane assembly in plants highlighted above, demands a rich repertoire of lipid transfer mechanisms which are summarized in Fig. 3 and which are discussed below. Conceptually they can be divided into (1) transmembrane transport and related leaflet-to-leaflet flipping mechanisms and (2) mechanisms of transport between two membranes, and (3) vesicular mechanisms. 3.1. Transmembrane transport and leaflet-to-leaflet flipping of polar lipids For the following discussion it is important to recall that biological membranes generally have a bilayer structure at life-supporting ambient temperatures, consisting of two leaflets of polar glycerolipids. The polar head groups of the glycerolipids face the aqueous environment while the acyl groups point towards the center of the membrane bilayer. A polar lipid or lipid precursor arriving from the ER at the cytosolic leaflet of the outer envelope membrane or synthesized by an enzyme associated with the cytosolic face of the outer envelope membrane (e.g. DGD1, Fig. 1) needs to be exchanged with the leaflet facing the intermembrane space (Fig. 3, process 1) to maintain the bilayer structure of the membrane. Likewise, asymmetric synthesis of the bulk of MGDG on the outside (intermembrane face) of the inner envelope membrane by MGD1 (Fig. 1) requires lipid transfer to the stroma-facing (inside) leaflet of the inner envelope membrane (Fig. 3, process 2). The flipping of polar lipids across a bilayer membrane in general does not occur spontaneously on a time scale relevant for cell division, but is catalyzed by lipid transporters or flippases as recently summarized in [46,47]. These proteins fall into different classes. Some require ATP and are involved in the maintenance of membrane asymmetry or the transport of lipids through a membrane

out of the cell or into an organelle. Others do not require ATP as they facilitate the equilibration of polar lipids between the two membrane leaflets during asymmetric lipid synthesis in biogenic (actively lipid synthesizing) membranes. ATP-requiring lipid transporters can be divided into ABC transporters and P-type ATPases and our current knowledge of ATPrequiring lipid transporters in plants can be summarized as follows: of the family of P-ATPase type lipid transporters one member of Arabidopsis has been described that is involved in cold tolerance and might affect plasma membrane lipid asymmetry [48]. Plant ABC lipid transporters include the exporters of precursors for cuticle or wax biosynthesis in the plant epidermis [49–52]. A fairly well studied plant lipid ABC transporter, PAX1/CTS/PED3, is localized in peroxisomes where it is believed to import FAs or acyl-CoAs destined for b-oxidation [53–55]. This transporter was also shown to be involved in the peroxisomal import of a jasmonate precursor [56]. Jasmonate is a plant growth regulator and oxidized FA derivative (oxylipin). Recently, based on labeling experiments of pax1/cts/ ped3 mutant seedlings with acetate, it was proposed that this ABC transporter could mediate the import of acetate into glyoxysomes [57]. Glyoxysomes are specialized peroxisomes that play a role during seed germination, when fatty acids are converted to carbohydrates by the glyoxylate cycle. The current data on the PAX1/CTS/ PED3 transporter exemplify a recurring dilemma for all systems studied thus far: it is exceedingly difficult to determine the substrates and substrate specificity of lipid transporters because no reliable in vitro assays are available, because the respective transporter is practically embedded into its substrate, and because these proteins currently elude functional reconstitution. The latter issue poses an even greater hurdle for the analysis of multi-component transporters assembled from different peptides. All plant ABC lipid transporters described above consist of a homodimer of a single peptide with a multiple-membrane-spanning permease domain and a membrane-peripheral ATP-binding cassette (ABC) domain. However, the only currently identified Arabidopsis ABC lipid transporter proposed to be relevant for ER-to-plastid lipid trafficking consists of three components, TGD1, 2, 3, similar to bacterial-type multi-component ABC transporters [26,36,58,59]. The TGD1, 2, 3 transporter is localized in the inner plastid envelope of Arabidopsis and is proposed to mediate the transfer of polar lipids through this membrane as will be discussed in detail below (Fig. 3, process 3). In plants DAG derived from the plastid pathway is produced from PtdOH on the stroma side of the inner envelope membrane [60], but has to be made available as substrate to MGD1, which is associated with the intermembrane face of the inner envelope membrane [22,26] (Fig. 1). Contrary to glycerolipids with large polar head groups, spontaneous transbilayer movement of DAG occurs on a time scale of 70 ms [61]. Therefore, it seems possible that this high rate of spontaneous DAG flipping is sufficient to alleviate the need for a protein facilitator. 3.2. Intermembrane transport of polar lipids Intermembrane lipid transfer plays a critical role in the development of the chloroplast and the assembly of photosynthetic membranes. For example, lipids have to be transferred between the two envelope membranes (Fig. 3, process 4). Although the molecular mechanism remains unclear at this time, clues might be gleaned from components and mechanisms proposed to be involved in the transport of nascent lipopolysaccharide (LPS) from the inner membrane of Gram-negative bacteria to the outer membrane as recently summarized in [62,63]. Recalling the bacterial endosymbiont hypothesis for the origin of chloroplasts, it might be instructive to briefly summarize key findings about lipid transfer to the outer membrane in Gram-negative bacteria as similar mechanism are involved in chloroplast development: genetic and biochemical analy-

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sis suggests that MsbA of Escherichia coli is an ABC transporter required for flipping of the LPS core synthesized at the cytoplasmic face of the inner membrane to the periplasmic face of the membrane [64,65]. Two peripheral proteins associated with the inner membrane, LptA and LptB, and possibly interacting with the membrane-spanning protein YrbK are proposed to be involved in the transfer of LPS from the inner to the outer membrane [66]. Mutant analysis also suggests that two outer membrane proteins of E. coli, Imp and RlpB, are involved in chaperoning LPS to the outer membrane [67–69]. It is possible that all these proteins assemble into a complex in contact zones between the outer and the inner membrane that have been proposed based on LPS inner-to-outer membrane transport experiments with spheroplasts [70]. The mechanism of lipid transfer between the ER and the outer chloroplast envelope remains to be identified. Previously, non-specific lipid transfer proteins as catalysts for intermembrane transfer (Fig. 3, mechanism 5) seemed most plausible based on their ability to extract lipids from a donor membrane and delivering it to an acceptor membrane in vitro [71]. However, at least some of these are extracellular proteins and are generally found to be involved in a variety of cellular processes, for example cell defense signaling mechanisms [72,73], in ways that cast doubt on their possible role in ER-to-plastid lipid transfer. More attractive seems to be the possibility of direct contact sites between the ER and the outer envelope membrane (Fig. 3, mechanism 6) that can be isolated in the form of ‘‘plastid associated membranes” (PLAMs) [74]. Aside from transmission electron microscopy evidence scattered throughout the literature, biochemical studies suggest that a non-bulk ER-fraction remains associated with the plastid envelopes during pea chloroplast isolation [75]. In addition, strong attracting forces at membrane contact sites between the ER and the plastid were demonstrated by in vivo optical manipulation [74]. However, it must be emphasized that the apparent existence of PLAMs by itself does not provide a mechanism of ER-plastid envelope membrane lipid exchange. Furthermore, it is not clear what the physical arrangement of polar lipids in such contact sites is. Presumably proteins are required to form these contact sites and to mediate the transfer of lipids. Contrary to PLAMs, analogous mitochondria associated membranes (MAMs) discovered in mammals and yeast [76,77] are well studied, and have been implicated in the transfer of phospholipids from the ER to the mitochondrion. The spatial separation of aminoglycerophospholipid biosynthetic enzymes in yeast with phosphatidylserine (PtdSer) synthase localized in the ER membrane, two distinct PtdSer decarboxylase isoforms localized in the mitochondria and Golgi, respectively, and PtdEtn N-methylases localized in the ER, requires the transfer of PtdSer to the mitochondrion or Golgi, and PtdEtn back to the ER [78]. Using mutant strains selectively inactivated in the PtdSer decarboxylase of mitochondria or the Golgi, enthanolamine auxotrophs were isolated that were impaired in the regulation of aminoglycerophospholipid biosynthesis or in components involved in lipid trafficking between these organelles [79–81]. In addition, based on biochemical analyses of the Golgi resident PtdSer decarboxylase 2 [82,83], a picture of macromolecular protein assemblies in contact zones between donor and acceptor membranes of different organelles in yeasts cells is emerging [84,85]. While the yeast system is not directly transferable to plants, it nevertheless shows the possibilities of applying a genetic approach to the problem. Recently, a genetic screen in Arabidopsis has led to the isolation of mutants disrupted in the transfer of lipids from the ER to the plastid (see below). Mapping of the mutant genes might provide components involved in the formation of PLAMs in the near future. 3.3. Vesicular lipid transport All mechanisms of lipid trafficking discussed above are nonvesicular. On the other hand, membrane protein trafficking between many organelles occurs via vesicles which bud from a donor

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membrane and fuse with an acceptor membrane, a process requiring an elaborate set of protein factors, e.g. [86]. Inherently, membrane protein trafficking by vesicles also provides a mechanism for the interorganelle transfer of glycerolipids. While vesicular transport appears not to be relevant for ER-to-plastid lipid trafficking in plants as discussed above, emerging evidence is consistent with a vesicular lipid transfer system inside plastids involved in thylakoid biogenesis by transferring lipids (and presumably proteins) from the inner envelope to thylakoids (Fig. 3, mechanism 7). Initial evidence for vesicular transport inside plastids was based on ultra-structural analyses [87] and inhibitor studies with isolated chloroplasts [88]. More recently, an Arabidopsis mutant with chloroplasts devoid of internal thylakoid membranes implicated the VIPP1 protein (vesicle-inducing protein in plastids) [89,90]. The protein had been previously proposed to be involved in lipid transfer [91], and inactivation of the respective ortholog in a cyanobacterium also abolished the formation of thylakoids [92]. Cyanobacterial Vipp1 and the chloroplast VIPP1 proteins form high molecular weight ring-like complexes and can be visualized in discrete locations at the inner envelope membrane [93]. Chaperone proteins HSB70B-CDJ2-CGE1 were found in tight association with VIPP1 in Chlamydomonas [94,95]. They are involved in the assembly and disassembly of oligomers of VIPP1. Taken together, the current evidence suggests that VIPP1 is a component of a complex potentially required for vesicular lipid trafficking from the inner plastid envelope to the thylakoids. Additional components of a vesicular transport system within plastids have been proposed based on bioinformatics analysis of the Arabidopsis genome [96]. Whether any of these candidates is involved in thylakoid formation, will have to be shown by genetic and biochemical analyses.

4. Fatty acid export from plastids Plastids such as photosynthetic chloroplasts are the biosynthetic factories of plant cells and the bulk of the FA precursors of lipids is synthesized in these organelles [12,97]. Export of FAs from plastids to the ER is likely the highest-flux lipid trafficking phenomenon in plants, yet we have no clear insights into the underlying molecular or biochemical mechanisms. Yield and quality of lipids (acyl composition) could be impacted by properties of the FA export machinery due to limited capacity or substrate selectivity, assuming that protein catalysts are involved (see discussion below). Knowledge about FA export seems crucial if we want to engineer lipid or oil biosynthesis in plants. In the following, the current state of knowledge and the basic facts are summarized: fatty acids in plastids are released from the fatty acid synthase complex as acyl-ACPs. Elegant labeling experiments with spinach leaves using [18O]-acetate clearly established that free FAs are formed by hydrolysis prior to their incorporation into ER lipids [98]. Furthermore, careful analysis of the kinetics of incorporation of [14C]-acetate into free FAs, acyl-CoAs and complex lipids using spinach and pea leaves identified a small, rapidly turned over pool of free FAs [99]. For both plants steady-state labeling of free FAs was reached within 2 min and the half-life of this pool was estimated to be less than 1 s. This transfer pool of free FAs is proposed to arise from the action of thioesterases on the inside of the inner envelope and reincorporation of free FAs by long chain acyl-CoA synthetase associated with the outside of the outer envelope membrane. Measuring the activity of this enzyme in isolated pea chloroplast using in situ formed FAs, distinct kinetic pools of free FAs were observed by varying the incubation conditions mid-time course. However, it was suggested that only the smallest pool, which was inaccessible to bovine serum albumin, was equivalent to the observed transfer pool in vivo. Moreover, the kinetic analysis did not support a model in which FAs move by simple diffusion

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across the envelope membrane. Instead, it was consistent with a small, channeled pool of FAs that most likely reflects protein-mediated transport of nascent FAs released on the inside by thioesterases to long chain acyl-CoA synthetases on the outside of the chloroplast [99]. The exact identity of the enzyme(s) that activates FAs during FA export is still uncertain. Inactivation of the Arabidopsis LACS9 gene encoding the enzyme responsible for 90% of the in vitro acyl-CoA synthetase activity associated with the chloroplast envelope membranes had no major effect on growth of the mutant [100]. One explanation for this result could be the redundancy of genes in Arabidopsis encoding long chain acyl-CoA synthetases [101]. Overall the findings made in plants on FA export from plastids may be related to the long-standing debate about whether a FA transporter is truly required for the transmembrane movement of FAs [102], because protonated FAs can readily cross a bilayer membrane with estimates of flip rates as high as 15 s 1 [103]. Based on a wealth of data derived from bacterial, yeast, and animal cells as summarized in [104] it seems clear that proteins are involved in the delivery of FAs to membranes on one face and can extract or metabolize FAs on the opposite face. Vectorial acylation, i.e. the conversion of free fatty acids to acyl-CoAs by acyl-CoA synthetases associated with the receiving face of the membrane can drive the transmembrane movement of FAs [104]. Presumably a similar vectorial acylation mechanism is involved in the export of FAs from plastids but as noted above, the exact components remain to be identified. Looking for alternative or additional players, e.g. a bona fide FA transporter, the intrinsic membrane protein FadL present in the outer membrane of gram-negative bacteria might serve as a reference [105]. Repeating the argument made above that plastids carry remnants of their cyanobacterial progenitor [43], it seems possible that plastids have a protein with a similar function. However, when comparing all genome encoded protein sequences of Arabidopsis with bacterial FadL sequences using BLASTP [106], no promising putative ortholog is detected. Thus, identifying the mechanism of FA export from plastids and the potentially involved protein components still presents a formidable challenge, but also an excellent opportunity.

5. Genetic analysis of ER-to-plastid lipid trafficking in Arabidopsis Recently, mutants of Arabidopsis directly affected in ER-to-plastid lipid trafficking have been isolated and have already aided in the identification of proteins involved in this process. These mutants were designated trigalactosyldiacylglycerol (tgd) [26,36,58,59] for the diagnostic accumulation of oligogalactoglycerolipids, which typically are absent from wild-type lipid extracts. The tgd mutants were initially isolated during a suppressor screen in the dgd1 mutant background [36]. The dgd1 mutant lacks 90% of the digalactoglycerolipid and is deficient in the DGD1 protein (Fig. 1) [24,107]. The screen was targeted at the isolation of mutants constitutively expressing the alternative galactoglycerolipid biosynthetic pathway involving MGD2, MGD3, and DGD2 [13,22,108]. While a mutant meeting the criteria for the disruption of a regulator of the alternative galactoglycerolipid pathway was identified, which was affected in a mitochondrial outer membrane protein [109], a substantial number of additional mutants were isolated that accumulated digalactoglycerolipids and oligogalactoglycerolipids. In a subsequent screen tgd mutants were also isolated in the wild-type background (C. Xu and C. Benning unpublished). The structure of the di- and trigalactoglycerolipids accumulating in the mutants was sufficiently different from the regular galactoglycerolipids in their glycosidic linkage [36] that they could not have been produced by the MGD1 or DGD1 galac-

tosyltransferases (see Fig. 1). Instead a distinct processive galactolipid:galactolipid galactosyltransferase was induced in the mutants, which is also active in isolated chloroplasts [110–113] and is still present in dgd1 dgd2 homozygous double mutants [25]. The gene for this enzyme and its role in the wild-type plant remain elusive. The tgd1 mutant has been analyzed in greatest depth [26,36]. Its complex phenotype can be summarized as follows: (1) it accumulates oligogalactoglycerolipids; (2) it also accumulates triacylglycerols in leaves; (3) the PtdOH level is increased in the mutant; (4) molecular species of thylakoid lipids derived from the ER pathway are underrepresented; (5) pulse-chase labeling experiments with acetate (which labels fatty acids in the plastid first; see Fig. 2) and oleic acid (which labels ER-lipids first) strongly indicate an impairment in the import of ER-derived lipids into the plastid; (6) isolated tgd1 chloroplasts show reduced labeling of galactoglycerolipids from PtdOH; (7) loss of TGD1 is embryo-lethal. Additional clues towards TGD1 function were derived from its similarity to integral membrane permease components of bacterial ABC transporters and its predominant location in the inner chloroplast envelope membrane, with a possible sub-fraction located in the outer envelope membrane. The other two tgd mutants studied in greater detail at this time, tgd2 and tgd3, have the same phenotype as tgd1. The molecular identity of the affected proteins is known: TGD2 is anchored with a single membrane-spanning domain in the inner chloroplast envelope with a globular domain facing the intermembrane space [58]. It contains an MCE (mycobacterial cell entry) domain [114] and specifically binds PtdOH. TGD3 was identified based on its similarity to non-intrinsic bacterial ATPases (ATP-binding cassette, ABC proteins) found adjacent in operons to bacterial homologs of TGD1 and TGD2 [59]. TGD3 is a soluble protein imported into the plastid where it could associate with TGD1 on the inside of the inner envelope membrane. Recombinant TGD3 protein fused to maltose binding protein has ATPase activity [59]. Based on these data it was proposed that the TGD1, 2, 3 proteins form an ABC transporter complex in the inner chloroplast envelope membrane (see Fig. 3, process 3). In light of the accumulation of PtdOH in the mutants, the reduced incorporation of PtdOH into galactoglycerolipids by isolated tgd1 chloroplast, and the specific binding of PtdOH by TGD2, which is considered the substrate binding protein of the presumed TGD1, 2, 3 complex, it is proposed that PtdOH is a transported substrate. This transport system seems to be required for the transfer of PtdOH formed at the outer chloroplast envelope from PtdCho to the inside of the inner envelope membrane where the PtdOH phosphatase resides [60], which produces DAG for galactoglycerolipid biosynthesis (see Fig. 1). In the mutants, PtdOH accumulates at the ER and the outer envelope membrane. It presumably activates the enzymes of TAG biosynthesis at the ER, and of oligogalactoglycerolipid biosynthesis at the outer chloroplast envelope membrane. Like the enzyme(s) involved in oligogalactoglycerolipid biosynthesis, the enzyme(s) of TAG biosynthesis activated in the mutants is not known at this time. As a central lipid metabolite, PtdOH also serves as precursor for plastid PtdGro biosynthesis (Fig. 1). In this pathway PtdOH and CTP are converted to CDP-DAG. This intermediate and glycerol 3-phosphate are converted to phosphatidylglycerolphosphate by PGP1 [115–117]. Subsequent dephosphorylation produces PtdGro. One prediction of the proposed PtdOH transport activity of TGD1, 2, 3 proteins is that PtdOH derived from the ER pathway should be available for PtdGro biosynthesis as well as galactoglycerolipid biosynthesis in plastids [118]. However, the FA composition of PtdGro suggests that it is exclusively derived from the plastid pathway as depicted in Fig. 1. Expression of a DAG kinase gene from E. coli in tobacco plant cells and targeting the respective protein to the plastid resulted in the interception of DAG derived from the ER pathway and its channeling into PtdGro in these lines [118].

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Unfortunately, the association of bacterial DAG kinase with one of the two plastid envelope membranes, and the topology of the enzyme were not determined making the interpretation of these data difficult with regard to the proposed TGD1, 2, 3 transporter. Most likely, the biosynthetic enzymes for PtdGro and galactoglycerolipid are subject to substrate channeling [118]. Consistent with this latter hypothesis, PtdGro biosynthesis is only slightly diminished in mutants impaired in the plastid glycerol 3-phosphate acyltransferase, while galactoglycerolipid biosynthesis in these mutants proceeds nearly exclusively by the ER pathway [17,18]. Another classic set of labeling data possibly at odds with the proposed function of the TGD1, 2, 3 complex is the observation that plants in which galactoglycerolipid biosynthesis is exclusively formed from ER-derived precursors appear to have a reduced activity of the plastid PtdOH phosphatase [39,119]. According to the current model shown in Fig. 1, PtdOH phosphatase plays a crucial role in both pathways of galactoglycerolipid biosynthesis. With a promising candidate for chloroplast PtdOH phosphatase now identified [21], the abundance and biochemical properties of orthologs in plants with exclusive ER pathway or with both pathways active can be directly compared. Moreover, the postulated TGD1, 2, 3 complex has yet to be studied in plants with an exclusive ER pathway of galactoglycerolipid biosynthesis. Recently, a novel Arabidopsis TGD protein, TGD4, was identified by mapping of the tgd4-1 mutant allele (C. Xu and C. Benning, unpublished). The tgd4-1 mutant has all the hallmarks of the tgd1, 2, 3 mutants suggesting that it is also disrupted in ER-to-plastid lipid trafficking. Because the TGD4 protein is not in the plastid, but possibly associated with the ER, the exciting possibility arises that TGD4 is required for the transfer of PtdCho or a related lipid to the outer envelope. It is also possible that TGD4 is directly involved in PLAM formation.

6. Perspectives Substantial progress has been made in recent years in identifying proteins and mechanisms involved in ER-to-plastid lipid trafficking and related phenomena in other organisms. Some digressions into non-plant model organisms were made in this review to provide inspiration to the plant scientists interested in these problems. However, they also highlight the fact that our general understanding of non-vesicular interorganelle lipid trafficking or the mechanism of lipid and FA transmembrane movement is still very basic even in the best studied model organisms. To some this dearth of knowledge presents itself as a great opportunity. As more lipid transporters/flippases and other proteins involved in lipid trafficking are discovered in different organisms, the momentum in the field is building and the pace of discovery is increasing. ER-to-plastid lipid trafficking in plants encompasses nearly all concepts and principles (see Fig. 3) that need to be addressed for a broad mechanistic understanding of interorganelle lipid trafficking. Genetic analysis in Arabidopsis has uncovered a set of proteins, TGD1, 2, 3, that could represent a PtdOH transporter, the first described for any organism. However, at this time a direct interaction of the three TGD proteins has not been demonstrated, the transporter has not been reconstituted in vitro, and its substrate specificity has not yet been determined. As for most of the other lipid transporters, addressing these issues remains very challenging. The discovery of a new component involved in ER-to-plastid lipid trafficking, TGD4, that is associated with the ER creates exciting new opportunities. Functional analysis of this novel protein is currently ongoing (C. Xu and C. Benning). The availability of this and other proteins involved in ER-to-plastid lipid trafficking opens new avenues to identify interacting partners, to develop functional assays and to establish an in vitro ER-to-plastid transport system

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based on comparison of mutant and wild-type ER and plastid fractions that will allow the identification of additional compounds by biochemical means. Exciting also is the in vivo visualization of PLAMs [74] using optical tweezers. Applying this technique to Arabidopsis mutants like tgd4 in comparison to wild-type should provide an avenue for the identification of proteins directly involved in PLAM formation. An ongoing quest is the search for the mechanism of FA export from plastids. Whether this problem can be addressed by genetics in plants remains to be determined. Based on the general discussion of FA transport through membranes it is not even clear whether a protein is required for the transport process. However, it is clear that acylation of free FAs on the outside of plastids must occur [98] which makes vectorial acylation an attractive model for directionally controlling fatty acid efflux. However, the abundance of gene families encoding long chain acyl-CoA synthetase isoforms in plants reduces the likelihood that unique components will be identified by genetic analysis. Furthermore, the process must be essential and finding conditional mutants in plants has not been an easy task. One possible approach could be screening for FA auxotrophs in the unicellular microalgae C. reinhardtii. Lipid transfer processes involving the plastid might rival in complexity the chloroplast protein import machinery. Even if this turns out to be only partially correct, one would expect that the tgd mutant collection is not yet saturated. At this time approximately 25 tgd mutants are available and mapping of the different mutant loci is currently underway (C. Xu, J. Gao, and C. Benning, unpublished). Additional proteins involved in ER-to-plastid trafficking could include ER-to-outer plastid envelope docking proteins, lipid flippases in the plastid outer envelope membrane, specific PtdCho lipases producing PtdOH at the outer envelope membrane, the PtdOH phosphatase associated with the inner envelope membrane, acyltransferases coupled to a lyso-PtdCho transporter as recently identified in yeast [120] if lyso-PtdCho is involved, and proteins required for the transfer of galactoglycerolipids from the envelopes to the thylakoids. Just as the yeast sec mutants revolutionized the study of the secretory pathway in eukaryotes [121,122], the tgd mutants of Arabidopsis hold the promise as crucial tools for the exploration of the mechanism of lipid import into plastids in plants. Acknowledgements I would like to thank all current and previous members of my lab, especially Changcheng Xu, Jilian Fan, Koichiro Awai, Jinpeng Gao, and Binbin Lu who contributed to our current understanding of lipid trafficking in Arabidopsis. Work on Arabidopsis chloroplast lipid metabolism in my lab is supported in parts by grants from the US Department of Energy (DE-FG02-98ER20305) and the US National Science Foundation (MCB 0453858). References [1] Dörmann P, Benning C. Galactolipids rule in seed plants. Trends Plant Sci 2002;7:112–8. [2] Joyard J, Marechal E, Miege C, Block MA, Dorne AJ, Douce R. Structure distribution and biosynthesis of glycerolipids from higher plant chloroplasts. In: Siegenthaler P-A, Murata N, editors. Lipids in photosynthesis: structure, function and genetics. Dordrecht, The Netherlands: Kluwer Acad. Publ.; 1998. [3] Dorne A-J, Joyard J, Douce R. Do thylakoids really contain phosphatidylcholine? Proc Natl Acad Sci USA 1990;87:71–4. [4] Browse J, Somerville CR. Glycerolipids. In: Meyerowitz EM, Somerville CR, editors. Arabidopsis. Cold Spring Harbor: Cold Spring Harbor Press; 1994. [5] Jouhet J, Marechal E, Block MA. Glycerolipid transfer for the building of membranes in plant cells. Prog Lipid Res 2007;46:37–55. [6] Webb MS, Green BR. Biochemical and biophysical properties of thylakoid acyl lipids. Biochim Biophys Acta 1991;1060:133–58. [7] Benning C, Garavito MR, Shimojima M. Sulfolipid biosynthesis and function in plants. In: Hell R, Dahl C, Knaff D, Leustek T, editors. Sulfur metabolism in phototrophic organisms. Doordrecht, The Netherlands: Springer; 2008.

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