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First published online March 2, 2007; 10.1104/pp.106.094102 Plant Physiology 144:402-418 (2007) © 2007 American Society of Plant Biologists
Characterization of Lipid Rafts from Medicago truncatula Root Plasma Membranes: A Proteomic Study Reveals the Presence of a Raft-Associated Redox System1,[W]Laboratoire des Interactions Plantes Micro-organismes, Unité Mixte de Recherche, Centre National de la Recherche Scientifique-Institut National de la Recherche Agronomique 2594/441, 31326 Castanet-Tolosan cedex, France (B.L., J.C.); Laboratoire de Biogenèse Membranaire, Unité Mixte de Recherche 5200-Centre National de la Recherche Scientifique-Université Victor Segalen Bordeaux 2, 33076 Bordeaux cedex, France (F.F., F.S.-B., J.-J.B., S.M.); Plate-forme proteomique, IFR40, 31326 Castanet-Tolosan cedex, France (M.R.); Plateau technique Imagerie/Cytologie, Institut National de la Recherche Agronomique, IFR 103, 33883 Villenave d'Ornon cedex, France (J.-P.C.); Institut de Biologie Moléculaire des Plantes, Unité Propre de Recherche-Centre National de la Recherche Scientifique 2357, 67083 Strasbourg cedex, France (M.-A.H.); and Crop Performance and Improvement Division, Rothamsted Research, Harpenden, Hertfordshire AL5 2JQ, United Kingdom (L.V.M., J.A.N.)
Several studies have provided new insights into the role of sphingolipid/sterol-rich domains so-called lipid rafts of the plasma membrane (PM) from mammalian cells, and more recently from leaves, cell cultures, and seedlings of higher plants. Here we show that lipid raft domains, defined as Triton X-100-insoluble membranes, can also be prepared from Medicago truncatula root PMs. These domains have been extensively characterized by ultrastructural studies as well as by analysis of their content in lipids and proteins. M. truncatula lipid domains are shown to be enriched in sphingolipids and  7-sterols, with spinasterol as the major compound, but also in steryl glycosides and acyl-steryl glycosides. A large number of proteins (i.e. 270) have been identified. Among them, receptor kinases and proteins related to signaling, cellular trafficking, and cell wall functioning were well represented whereas those involved in transport and metabolism were poorly represented. Evidence is also given for the presence of a complete PM redox system in the lipid rafts.
The plasma membrane (PM) of living cells is known to play a key role in the perception and transduction of signals that regulate the communication of cells with their environment. A model of PM structure has emerged in the past decade from biophysical and biological studies on animal PM in which liquid-ordered domains enriched in cholesterol/sphingolipids coexist with liquid-crystalline domains rich in phospholipids containing unsaturated fatty acids (for review, see Simons and Vaz, 2004  ). Sterols are thought to fill the voids between the sphingolipid acyl chains, resulting in an increase of the packing of lipids. Hydrophilic interactions also occur between the hydroxyl groups of the sterols and the sphingolipid acyl chains. Because sphingolipid/cholesterol-rich liposomes were found to be insoluble in mild nonionic detergents such as Triton X-100 (TX100) at 4°C, the analysis of low-density TX100-insoluble membrane fractions were thought to reflect the in vivo composition of lipid microdomains (Brown and Rose, 1992; Schroeder et al., 1994; Simons and Ikonen, 1997; Brown and London, 1998). Consequently, one could assume that the insolubility in cold TX100 and the ability to float on density gradients of a given protein constituted a criterion for its localization or its interaction with these lipid microdomains. These membrane domains were termed lipid rafts by Simons and Ikonen in the late 1990s, a name which rapidly raised their popularity both in mammalian and yeast (Saccharomyces cerevisiae) biology (Rajendran and Simons, 2005). Lipid rafts have been proposed to play biological roles in signal transduction (Simons and Toomre, 2000), exocytosis (Salaün et al., 2004), endocytosis (Parton and Richards, 2003), apoptosis (Garcia et al., 2003), and actin cytoskeleton organization (Wickstrom et al., 2003; Falk et al., 2004). In addition, lipid rafts appear to be used for toxin and virus cellular entry (Rosenberger et al., 2000; Lafont et al., 2004) and for viral budding (Brugger et al., 2006). The in vivo occurrence, size, and extent in the PM of lipid rafts are still matters of debate (Munro, 2003). However, intensive work has been performed in some lipid raft proteins such as the T-cell receptor showing that their detergent-insolubility property correlates with functional properties (for review, see He et al., 2005). The emerging use of microscopic analyses to localize proteins in lipids raft and to deduce their lateral mobility should give new insights into in vivo lipid raft protein composition, size, and dynamics (Kenworthy et al., 2004; Douglass and Vale, 2005).
Despite this rising interest, the literature on membrane domains in plant cells is still poorly documented (for review, see Bhat and Panstruga, 2005 This work is devoted to the complete characterization of PM lipid rafts from Medicago truncatula roots. Roots are responsible for many important plant functions such as water and nutrient acquisition essential for cellular metabolism and plant growth, adaptation to biotic and abiotic stress such as pathogen attacks, or salt stress. They also form agronomically and environmentally mutualistic interactions with soilborne microorganisms, among which the most widespread are the symbioses with mycorrhizal fungi. The symbiosis with rhizobial bacteria, which is restricted to legumes, is also very important because it leads to atmospheric nitrogen fixation and the growth of these plants without an added nitrogen source. The legume M. truncatula has been internationally adopted as a model plant and genomic and EST sequence banks are well developed. The involvement of the root PM in perception, signalization, or exchange with the soil as well as the putative role of lipid rafts as a signaling platform prompted us to characterize lipid rafts from M. truncatula root PM. To our knowledge, this study is the first to report the characterization of PM lipid rafts from root tissue. We isolated a root PM-enriched fraction by phase partition and determined the lipid composition of DIMs compared with the root PM. Sphingolipids, free sterols, and steryl glycosides (SGs) appeared as the main lipid components of these DIMs. A morphological comparison between PM and DIMs was also carried out, by electronic microscopy, showing the high rigidity of DIMs and their strong ability to undergo self adhesion. A proteomic approach allowed us to identify 270 proteins including numerous proteins that might be involved in cell signaling (RLK, Phospholipase C, remorin, etc) and in an electron transfer system (peroxidase, NADP ubiquinone oxidoreductase, cytochrome b561, L-ascorbate oxidase [AO]).
Isolation of PM and DIM from Roots of M. truncatula We used aeroponic cultures of M. truncatula plants and the conventional phase-partition procedure to obtain PM-enriched fraction (approximately 500–700 µg protein per 150 g root fresh weight). The purity of the PM fraction was first checked with marker activities to evaluate the contamination by other membranes. Measurement of the vanadate-sensitive ATPase activity showed a 7-fold enrichment in PM proteins after phase partition compared with the starting material consisting of a crude microsomal fraction; a classical ratio for this technique. The mitochondrial marker activity was depleted by a factor of 7.0 between microsomes and PM, and the tonoplast marker by a factor of 5.6 (see Supplemental Table S1). The absence of monogalactosyldiacylglycerol (data not shown) indicated a lack of contamination by plastid envelopes. The purity of the PM fraction was further confirmed by ultrastructural analysis (see below). These results indicate that we could purify a PM-enriched fraction from Medicago roots, without significant contaminant by other endomembranes and suitable as starting material for extraction of lipid rafts.
The property of insolubility of particular proteins and lipids in nonionic detergents at low temperature has been commonly used to characterize membrane microdomains called lipid rafts. The Medicago root PM fraction was treated for 30 min with 1% cold TX100 with a ratio of detergent-to-protein of 12:1, as previously established (Mongrand et al., 2004
The DIM fraction was used for further studies and compared to the PM fraction from which it originates (Fig. 1). The study consisted of (1) morphological studies by electron microscopy of isolated fractions after chemical or physical fixation, (2) lipid characterization using high performance (HP)-TLC, gas chromatography (GC), and mass spectrometry (MS) analyses, and (3) protein identification by a proteomic-based approach of one-dimensional SDS-PAGE coupled to nano-liquid chromatography (nLC)-electrospray ionization (ESI) analysis.
Membranes (PM and DIMs) were processed either by chemical fixation or by physical fixation (high-pressure cryofixation followed by freeze substitution). Electron microscopy observations showed that the Medicago PM fraction contained mostly membrane vesicles and only a small amount of dense, nonmembranous material. The membrane of vesicles assumed to originate from the PM was thicker (5.5–6 nm) and displayed a heavier electronic contrast. After chemical fixation, the shape and contrast of these vesicles were very variable, some of them appeared to be more or less spherical, whereas others were flattened (Fig. 3, A and C ). By contrast, after freeze fixation and substitution most of the vesicles were distinctly ring shaped, with a high electronic contrast, the membrane leaflet being very clearly delineated (Fig. 3, B and D). It has to be noted that no change was observed in regard to apparent membrane thickness when the two methods were compared.
The DIM pellet (Fig. 4 ) was also processed by chemical fixation or by physical fixation. In both cases, it was found to contain mostly long membrane fragments, some membrane vesicles, and also aggregates of dense material. After chemical fixation, these components were scattered randomly in the pellet (Fig. 4A). Membrane strips could be seen either as transverse sections or flat views (Fig. 4C), they were not straight edged, but with irregular outlines, 40 to 120 nm wide. The thickness (approximately 6.5 nm) and the electronic contrast of the membranes were constant all along the fragments. After high-pressure fixation, the general organization of the DIM pellet was quite different. The pellet components were not loose but tightly packed and consisted of long, straight membrane fragments, either isolated or densely packed, which appear to stack into onion-like structures (Fig. 4, B and F). This aggregation was probably due to the high pressure during the fixation procedure. Unlike chemically fixed ones, freeze-fixed DIMs appeared to be far less tilted and appeared mostly as cross sections, showing even thickness and contrast and more even width, approximately 50 nm (Fig. 4D). A distinct attention was paid to the dense aggregates, which were clearly seen between membrane fragments. High magnifications showed that they consist of two different elements, namely very small 10 to 15 nm long membrane fragments and discrete clusters of seemingly proteinaceous dense material (Fig. 4E). Taken together our observations showed that DIMs are distinct from the PM vesicles and consist largely of long and narrow, highly rigid, membrane strips. These observations also demonstrated that high-pressure freeze fixation of membrane pellets followed by freeze substitution provides a better definition of membrane shape and contrast compared to chemical fixation.
Polar and Neutral Lipids of Medicago PM and DIM Fraction To investigate the lipid content of PM and DIMs, we first extracted lipids by chloroform:methanol (2:1). The various lipid classes were further separated by HP-TLC with a series of different solvent mixtures. Figure 5A displays HP-TLC plates for the separation of phospholipids (polar lipids 1), SGs (polar lipids 2), and neutral lipids. Acyl-containing lipids of PM and DIM were quantified by densitometric scanning after copper sulfate staining, and by GC analysis after methanolic acid hydrolysis, for sterol-containing lipids. Figure 5B shows that glycerolipids (phospholipids and digalactosyldiacylglycerol) represented up to 50.9% of the total polar lipids in the PM but only 23.8% in DIMs. In contrast, we observed in the DIMs, an enrichment in free sterols (1.7-fold, P value = 0.025), SGs (1.6-fold, P value = 0.001), and acyl-SGs (1.8-fold, P value = 0.019), glucosylceramide being not significantly enriched (1.3-fold, P value = 0.297). Neutral lipid analysis of Medicago PM and DIMs showed only two major components, namely free sterols and free fatty acids; sterol esters, triacylglycerol, and monoacylglycerol were not detected (Fig. 5A).
Free phytosterols were 1.7-fold enriched in DIMs compare to PM (Fig. 5B). They were further purified from the plates and analyzed as acetate derivatives by GC. The GC patterns revealed one major component and at least five minor sterols (Fig. 6A
). The Medicago sterols were quantified by GC and identified by GC-MS (Itoh et al., 1981
Sphingolipids and SGs of PM and DIMs were recovered after mild-alkaline hydrolysis of corresponding whole lipid extracts. HP-TLC analysis of the alkaline-stable lipids extracted resolved only three bands (Fig. 7A
): free fatty acids (resulting from glycerolipid hydrolysis), neutral lipids (i.e. free sterols), and glucosyl ceramide (gluCER) mixed with SG. These two latter lipids were separated from each other by HP-TLC using a solvent mixture able to resolve GluCER and SG molecular species into different groups (Fig. 7B). We used as standards the tobacco leaf PM gluCER, previously characterized by matrix-assisted laser-desorption ionization time of flight (Mongrand et al., 2004
Acylated SGs (ASGs) and SG were 1.6-fold (P = 0.001) and 1.8-fold (P = 0.019) enriched in DIMs compare to PM. Their position on the HP-TLC plate was determined by comparison with that of commercially available standards. These lipids were also identified due to their pink color after spraying HP-TLC plates with orcinol/sulfuric acid and to their blue color after copper sulfate staining. We further characterized the hydrophobic moieties of SG and ASG (see experimental procedures described in "Materials and Methods"). SG and ASG sterol moieties were quantified by GC. Figure 7D shows that SG and ASG contained the same sterols as the free forms, but in different relative proportions. Thus, the relative amount of
Proteins copurified with the sphingolipid-sterol-enriched DIM fractions were separated by monodimensional SDS-PAGE. The gel strip was divided into 14 fractions and peptides released by trypsin digestion were analyzed by nLC-ESI using a Q-TRAP mass spectrometer. The peptide sequences deduced from MS were attributed to M. truncatula proteins using the MASCOT search engine against either M. truncatula EST, genomic, or the National Center for Biotechnology Information (NCBI) databases (see experimental procedures). A total of 270 proteins were identified with this method with a probability of misidentification of <0.05, based on the peptide fragmentation qualities. These 270 proteins were classified as follows: 43 signaling proteins (Table I), 65 transport proteins (Table II ), 21 redox proteins (Table III ), 30 cytoskeleton, trafficking, and protein-stability factors (Table IV ), and 111 miscellaneous proteins including 22 with unknown functions (Table V ). For highly related members of multigene families, only the member with the highest mascot score is shown in the tables, with the others being described in Supplemental Table S2. Bioinformatics tools were then used to predict the Mrs, the number of transmembrane domains, the posttranslation addition of a glycosylphosphatidylinositol (GPI) anchor, and the presence of a signal peptide in each of the identified proteins (see experimental procedures), and results are shown in the different tables. When the subcellular location of a protein was experimentally determined in Medicago or other plant species, it is shown in the tables in roman type. If not experimentally determined, the subcellular location of soluble proteins was predicted with Psort and is shown in italics. Predictions of subcellular locations for intrinsic membrane proteins are not reliable and therefore, the putative location of these proteins (shown in italics) is based on suggestions in the literature.
Out of the 43 signaling proteins, 31 of them are RLKs. The proteins have been classified in this family based either on their homology to known RLKs or by the presence of a transmembrane domain linked to either a RLK extracellular or kinase domain. Twenty-one specific RLK proteins were unambigouously identified through having at least one specific peptide (see Table I). Sixteen RLKs belong to the LRR-RLK families but two representatives of the lectin, one of the S-locus, and one of the DUF26 families were also identified. In addition 10 RLK peptides were identified that were common with some of the 21 unambigouously identified RLK proteins but might suggest also the presence of members of the S-locus, DUF26, WAK, and LRR families (shown in green in Supplemental Table S2). Many proteins identified in DIMs are involved in water and nutrient acquisition (Table II). The membrane intrinsic proteins commonly called aquaporins are represented by 17 members (composed of both PIP1 and PIP2 family members). In addition, two phosphate, one nitrate, and one copper transporter have been identified. Other transport proteins include ion pumps and channels (13 H+-ATPases, nine ATP-binding cassette [ABC] transporters, and eight voltage-dependent anion channels). Several probable integral membrane proteins with a redox activity have been identified (an L-AO, five peroxidases, a cytochrome b561, a germin, and a plastocyanin-like protein). For almost all of them, a signal peptide has been predicted by the signalP algorithm, suggesting that these proteins follow the secretory pathway and distinguishing them from related mitochondrial or plastidic redox proteins (Table III).
Several isoforms of Table V regroups all the other identified proteins in the Medicago DIMs among which ribosomal proteins represent a large number (50). Among the other proteins identified, six are involved in lipid and six in saccharide (three glycoside hydrolase, an endoglucanase, and two Suc synthases) metabolism (Table V). No function could be attributed to 22 other proteins, because either homologs found with BLASTp have also an unknown function or domain identification by Prodom or Pfam algorithm was unsuccessful. Therefore a putative raft location could give a first indication in search of the functions of these proteins.
Several of the identified proteins belong either to multigenic families (e.g. 17 aquaporins) or to multisubunit complexes (e.g. tubulin
In this study we have used a variety of techniques to characterize the structure, the lipid, and the protein content of lipid rafts from the root PM of the plant model M. truncatula. Membrane lipids in plants consist of basically three classes of molecules, namely the diacylglycerol-based lipids, the long-chain-sphingoid-based lipids, and the sterols and their conjugates; these lipids play important roles in determining the physicochemical properties of the biological membranes. Sterols and sphingolipids have been detected as the prominent constituents of lipid rafts from tobacco leaf and BY2 cell PM (Mongrand et al., 2004 ) and from Arabidopsis callus microsomes (Borner et al., 2005). The presence of sterols, glycerolipids, and sphingolipids in root PM (Gronewald et al., 1982) suggest that liquid-ordered (raft) and liquid-crystalline (nonraft) phases could exist in roots. Here we show that DIMs can be isolated from Medicago root PM and share some characteristics with the Arabidopsis and tobacco lipid rafts. Microscopic examination of purified Medicago DIMs show they have a different structure to the PM and consist of long, narrow, and rigid membrane strips, with a lipid bilayer slightly thicker than that of the PM. We must emphasize that the length of isolated DIM strips are too large to reflect the size of membrane domains in vivo, and moreover it has been shown that TX100 treatment triggers aggregation of rafts (Giocondi et al., 2000). Therefore, ultrastructural analysis of isolated DIMs, which reveals a broad size range of membrane strips, are likely to represent a coalescence of smaller raft domains (Shogomori and Brown, 2003).
Compared to the PM, Medicago root lipid rafts were found to be greatly enriched in sphingolipids, free sterols, and SGs (up to 73.5% of the total raft lipids) and consequently depleted in phospho and glyco glycerolipids (Fig. 5B). Major free sterols of the Medicago root PM have been identified as
In Medicago root lipid rafts, we have also detected SGs in which the 3
Lipid domain-promoting activity of a membrane component is usually considered as its ability to strengthen close packing with saturated lipids (conversely, compounds that impair lipid packing behave as domain-disrupting compounds). Very little attention has been paid to the effects of SG and ASG on plant membrane properties. One reason is probably the lack of these glycolipids in the PM of mammalian cells. It is expected that SG molecules (Fig. 7F) are oriented in membranes with the sterol moiety embedded in the hydrophobic phase of the bilayer and the sugar in the plane of the polar head groups of the phospholipids. ASGs have probably both the sterol and the fatty acyl chain moieties inside the bilayer and the sugar lying along the hydrophilic surface. In a monolayer system, at a surface pressure of 21 dynes/cm, SG and ASG were found to occupy an area of 38 Å/mol and 54 Å/mol, respectively (40 Å/mol for cholesterol), a result consistent with the hairpin orientation proposed for ASG in the lipid bilayer (Mudd, 1980
Proteins associated with Medicago lipid rafts represent about 12% of the total PM protein (Fig. 2), a percentage similar to that observed in tobacco leaves but 2-fold higher than BY2 cells (Morel et al., 2006
Our analysis of root lipid rafts identified many proteins that have previously been identified in other lipid rafts (see below). Although our proteome analysis does not establish the enrichment of a protein in the lipid-raft fraction, the identification of these known lipid-raft-associated proteins, coupled with the extensive lipid and structural analyses, suggests that the fraction used for proteomic analysis is highly enriched in lipid rafts. This conclusion is supported by studies on the PM H+-ATPases, which was identified in this analysis in addition to previous studies on lipid rafts of both plants (Mongrand et al., 2004
The major proteins found in Medicago root lipid raft are signaling and redox proteins, as well as some subfamilies of pump and channels. In common to other lipid raft studies, we identified aquaporins, ABC transporters, RLKs, remorin, prohibitin, and the PM H+-ATPases (Shahollari et al., 2004
An interesting result of this study is the identification of many proteins belonging to the PM redox system in the Medicago root lipid raft (for review, see Vuletic et al., 2005
Our data show that many RLKs are present in the root DIM fraction (Table I), strengthening the hypothesis that root lipid rafts, as in animals, are involved in signaling. The RLKs is one of the largest families of plant proteins with about 600 members in Arabidopsis and 1,200 members in rice (Oryza sativa; Shiu et al., 2004 In conclusion, the work presented here represents a careful and extensive analysis of the structure, lipid, and protein content of lipid rafts from an important plant organ, roots, that had not been characterized previously. The results reveal several novel features of the lipids in these microdomains, and the presence of a redox system, which may be specific to their function in roots. As M. truncatula is serving as a model organism for studies on root endosymbioses, these studies will also form a base for future work to examine the role of protein recruitment in plant lipid rafts during symbiotic infection, studies that are complicated by the presence of the two organisms.
Materials HP-TLC plates were Silicagel 60 F254 (Merck). Steryl conjugates were purchased from Matreya LLC, other lipid standards and all other reagents were from Sigma.
For each experiment, about 400 plants of Medicago truncatula Gaertn ‘Jemalong’ genotype A17 were grown in an aeroponic chamber for 14 d (22°C; 16 h of light per day) in a nitrogen-rich medium (5 mM NH4NO3), then were starved of this nitrogen source for 4 d, as described in Journet et al. (2001)
The PM was purified essentially according to Rabotti and Zocchi (1994)
TX100 (10%) was added to a ratio of detergent-to-PM proteins of 10 to 12:1 (in all cases, 1% final concentration), and the membranes were solubilized at 4°C for 30 min, then brought to a final concentration of 52% Suc (w/w), overlaid with successive 3 mL steps of 40%, 35%, and 30% Suc in TBS buffer (w/w), and then centrifuged for 16 h at 200,000g in a TST41 rotor (SORVALL). DIMs could be recovered above the 30% to 35% interface as an opaque band and this fraction was washed in 4 mL of TBS buffer to remove residual Suc. The protein concentration was determined with a bicinchoninic acid protein assay to avoid TX100 interference, using bovine serum albumin as a protein standard.
Purified Medicago PM and DIM were pelleted at 100,000g. For chemical fixation, pelleted membranes were sequentially fixed with glutaraldehyde (2.5% in 100 mM phosphate buffer, 2 h at 4°C), osmium tetroxide (1% buffered, 2 h at 4°C), and tannic acid (1% in deionized water, 30 min at 22°C), according to Carde (1987)
Lipids were extracted and purified from the different fractions according to Bligh and Dyer (1959)
Free sterols, SG, and ASG sterol moieties were analyzed by GC as acetate derivatives with a Varian GC model 8300 equipped with a flame ionization detector and a fused capillary column (WCOT 30 m x 0.25 mm i.d.) coated with DB1, with the following program: an initial rise of temperature from 60°C to 220°C at 30°C/min, a rise from 220°C to 280°C at 2°C/min and a plateau at 280°C for 10 min. Cholesterol was used as internal standard. Sterol acetates were identified by GC-MS according to Rahier and Benveniste (1989)
Total sphingolipid levels were determined by quantitative measurement of sphingolipid LCBs, using methods previously described by Borner et al. (2005)
Data are presented in means ± SD from at least three independent experiments from three different biological samples. Comparisons were performed by Student's t test and P < 0.05 was considered statically significant.
Proteins in the Suc density gradient were precipitated in 10% cold TCA for 30 min at 4°C. After centrifugation the pellet was first washed with 10% TCA in water to remove residual Suc, and finally with cold acetone before being resuspended in Laemli buffer for SDS-PAGE and western-blot analysis.
All Antibodies were diluted in TBS-tween supplemented with 1% powdered milk as follows: anti-H+-ATPase (Maudoux et al., 2000
A total of 25 µg of DIMs resuspended in TBS were centrifuged at 100,000g and the pellet was solubilized overnight in 20 µL, 6 M Urea, 2.2 M Thiourea, 5 mM EDTA, 0.1% SDS, 2% N-Octyl-Glucoside, 50 mM Tris-HCl pH 8.8. After addition of 13 µL, 4% SDS, 100 mM dithiothreitol, 10% glycerol, 50 mM Tris-HCl pH 8.8, samples were incubated 30 min at room temperature, centrifuged 5 min at 20,000g before protein separation by 1D SDS-PAGE (separating gel: 7.5% acrylamide, stacking gel: 4%). The gel tracks were each cut into 14 slices and the proteins in each slice were digested overnight by 150 ng trypsin as described in Borderies et al. (2003)
All analysis were performed using a Q-TRAP (LC packings and Applied biosystems/MDS Sciex) nLC-MS/MS system as described in Boudart et al. (2005)
Transmembrane domains were predicted by TMHMM2 (http://www.cbs.dtu.dk/services/TMHMM/), beta-barrel by Pred-TMBB (http://biophysics.biol.uoa.gr/PRED-TMBB/), signal peptide by signalP (http://www.cbs.dtu.dk/services/SignalP-2.0/), myristoylation by the MYR Predictor (http://mendel.imp.ac.at/sat/myristate/SUPLpredictor.htm), GPI anchor by big-PI Plant Predictor (http://mendel.imp.ac.at/sat/gpi/plant_server.html), subcellular location by Psort (http://psort.nibb.ac.jp/form.html) or according to the SUBA database (http://www.plantenergy.uwa.edu.au/applications/suba), and protein functional domains by Prodom (http://prodes.toulouse.inra.fr/prodom/current/html/home.php).
The following materials are available in the online version of this article.
MS analysis was performed at the "Plate-forme proteomique, IFR40" with the help of Gisèle Borderies and Carole Pichereaux. Electron microscopic observations were performed at the "Plateau Technique Imagerie/Cytologie, IFR 103," with the skillful collaboration of Martine Peypelut. We are grateful to Jérome Gouzy and Hélène San Clemente for help with in silico data analysis and to Alain Puppo for helpful discussion and critical reading of the manuscript. We thank Frederic Domergue for GC analyses and Marc Boutry for anti-H+-ATPase antibodies. Received December 4, 2006; accepted February 20, 2007; published March 2, 2007.
1 This work was supported by Conseil Régional d'Aquitaine, France (to J.-J.B., S.M., F.S.-B., F.F.), and Agence Nationale de la Recherche (to J.C., S.M.). B.L. and F.F. are funded by fellowships from the Ministère délégué à l'Enseignement supérieur et à la Recherche, France.  The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Sébastien Mongrand (sebastien.mongrand{at}biomemb.u-bordeaux2.fr).
[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.106.094102 * Corresponding author; e-mail sebastien.mongrand{at}biomemb.u-bordeaux2.fr; fax 033–5–56–51–83–61.
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-D-glucopyranoside (SG) and of spinasterol (6'-O-palmitoyl)-3
and 
