Journal of Experimental Botany, Vol. 52, No. 357, pp. 669-679,
April 15, 2001
© 2001 Oxford University Press
Original Papers |
Characterization of plasma membrane domains enriched in lipid metabolites
Department of Biology, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1
Received 27 July 2000; Accepted 25 September 2000
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and methodsResultsDiscussionReferences |
|---|
A subpopulation of plasma membrane vesicles enriched in membrane lipid metabolites has been isolated from petals of carnation flowers and leaves of canola seedlings. This was achieved by immunopurification from a microsomal membrane preparation using region-specific antibodies raised against a recombinant polypeptide of the plasma membrane H+-ATPase. The properties of this subpopulation of vesicles were compared with those of purified plasma membrane isolated by partitioning in an aqueous dextran-polyethylene glycol two-phase system. The lipid composition of the immunopurified vesicles proved to be clearly distinguishable from that of phase-purified plasma membrane, indicating that they represent a unique subpopulation of plasma membrane vesicles. Specifically, the immunopurified vesicles are highly enriched in lipid metabolites, including free fatty acids, diacylglycerol, triacylglycerol and steryl and wax esters, by comparison with the phase-purified plasma membrane. These findings can be interpreted as indicating that lipid metabolites generated within the plasma membrane effectively phase-separate by moving laterally through the plane of the membrane to form discrete domains within the bilayer. It is also apparent that these domains, once formed, are released as vesicles into the cytosol, presumably by microvesiculation from the surface of the plasmalemma. Such removal may be part of normal membrane turnover.
Key words: Immunoprecipitation, H+-ATPase, plasma membrane domains.
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
The lipid and protein constituents of membranes have a finite lifespan and, like other macromolecules in the cell, are broken down and resynthesized, a process commonly referred to as membrane turnover (Mazliak, 1980
; Steer, 1988; Hare, 1990). Membrane proteins, for example, that become conformationally altered are proteolytically degraded and replaced (Duxbury et al., 1991). Membrane turnover also encompasses continuous phospholipid breakdown (Sandelius and Sommarin, 1990; Wang et al., 1993; Kim et al., 1994) and synthesis (Ohlrogge and Browse, 1995). In particular, there is growing evidence that phospholipases mediating de-esterification of phospholipid fatty acids play a role in membrane turnover as well as in the release of lipid signalling molecules (Chapman, 1998). Little is known, however, about the mechanisms for removal of lipid metabolites from the bilayer. It is likely that lipid breakdown products are released from membranes as part of the turnover process inasmuch as their accumulation in the bilayer would destabilize membrane structure. There is, for example, an accumulation of free fatty acids as well as steryl and wax esters in membranes during senescence, and this leads to lipid phase separations in the bilayer and membrane leakiness (Yao et al., 1991).
In the present study, vesicles of plasma membrane origin enriched in lipid metabolites, including free fatty acids and steryl and wax esters, have been purified from petals and leaves by immunoprecipitation from microsomal fractions. This was achieved using antibodies raised against a recombinant polypeptide corresponding to the central hydrophilic region of the H+-ATPase. This enzyme is an integral protein associated with the plasma membrane that energizes the translocation of protons from the cytosol to the cell exterior (Briskin, 1990). The electrochemical and pH gradient established by the H+-ATPase in turn mediates other secondary transport systems associated with the cell membrane. Accordingly, the plasma membrane H+-ATPase acts as the primary transducer between chemical energy (in the form of ATP) and the generation of potential energy used to drive transport phenomena and as such, plays a central role in the physiology of a living cell. It controls ion and mineral uptake by the plant (Leonard, 1984), cell turgor (Curti et al., 1993), intracellular pH (Kurkdjian and Guern, 1989) and, indirectly, through the acidification of the apoplastic space, cell expansion (Rayle and Cleland, 1992). It is an abundant membrane protein and serves as a reliable marker for the plasmalemma.
In higher plants, the plasma membrane H+-ATPase is encoded by a multigene family containing up to 10 highly conserved isoforms. Several of these have been identified and characterized in a number of plant species including tomato (Lycopersicon esculentum, seven isoforms) (Ewing et al., 1990; Ewing and Bennett, 1994), Arabidopsis thaliana (five isoforms) (Harper et al., 1989, 1990, 1994; Pardo and Serrano, 1989; Houlne and Boutry, 1994), Nicotiana plumbaginifolia (four isoforms) (Boutry et al., 1989; Moriau et al., 1993), Oryza sativa (two isoforms) (Wada et al., 1992), and seagrass (Zostera marina L.) (Toshiyuki et al., 1996). Expression of the H+-ATPase isoforms appears to be cell- and tissue-specific, possibly reflecting functional diversity. For example, in Arabidopsis, the H+-ATPase encoded by AHA3 (Arabidopsis H±-ATPase isoform 3) has been immunolocalized to the plasma membrane of phloem companion cells and is believed to provide energy for active phloem loading (DeWitt and Sussman, 1995). Also in Arabidopsis, the transcripts of AHA9 are expressed mainly in anther tissue and encode the isoform involved in pollen tube growth (Houlne and Boutry, 1994), whereas AHA10 is expressed exclusively in the seed integument and drives the flow of nutrients to the developing embryo (Harper et al., 1994).
In the present study, polyclonal antibodies raised against a recombinant fragment of the AHA1 H+-ATPase isoform were used to immunopurify plasma membrane vesicles from petals and leaves, and the properties of these immunopurified vesicles have been compared with those of corresponding plasma membrane preparations obtained by phase partitioning. The observations indicate that the immunopurified vesicles represent lipid metabolite-enriched domains within the plasma membrane that have been released into the cytosol, presumably by microvesiculation.
|
Materials and methods |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Plant material
Microsomal and plasma membranes were isolated from petals of carnation flowers (Dianthus caryophyllus L. cv. White Sim) and from leaves of canola (Brassica napus L.). Carnations were grown under standard greenhouse conditions and fertilized on a continuous feed schedule with 28-14-14 N-P-K (Plant Products). Lighting was supplemented from dawn to dusk with high pressure sodium lamps (General Electric Lucalox, 400 W, 210 µmol m-2 s-1). Day and night temperatures were 25 °C and 18 °C, respectively. The flowers were harvested when the petals were fully expanded with yellow-tinted centres. Canola seedlings were grown in chambers at 23 °C, 50% humidity, under 16/8 h day/night photoperiods. Fully expanded primary leaves of canola were harvested 16 d after planting.
Microsomal and plasma membrane isolation
Plasma membrane was purified from leaf and petal tissue by partitioning microsomes in an aqueous dextran-polyethylene glycol two-phase system (Kjellbom and Larsson, 1984). Petal or leaf tissue (50 g) was homogenized in 200 ml of homogenization buffer (0.5 M sucrose, 50 mM HEPES-KOH, pH 7.5, 5 mM ascorbate, 3.6 mM cysteine, 0.5 mM PMSF, and 0.6% insoluble PVPP) for 45 s in a Sorvall Omninixer and for an additional minute in a Polytron homogenizer. The homogenate was filtered through four layers of cheesecloth and centrifuged at 10 000 g for 20 min. Microsomal membranes were pelleted from the supernatant by centrifugation at 250 000 g for 1 h. For plasma membrane isolation, the microsomes were resuspended to 9 ml in 0.33 M sucrose, 5 mM potassium phosphate, pH 7.8, and 3 mM KCl (plasma membrane resuspension buffer). This suspension was mixed with Dextran T 500 and polyethylene glycol 3350, both at a final concentration of 6.2% (w/w), to form the aqueous two-phase polymer system. The phase system was vigorously shaken and centrifuged in a swing-out rotor for 5 min at 4000 g. The plasma membrane fraction (upper-phase) was purified by sequential partitioning against fresh lower-phase solution as described earlier (Kjellbom and Larsson, 1984). The upper-phases were pooled, diluted 3-fold with plasma membrane resuspension buffer and centrifuged at 250 000 g for 1 h to pellet the plasma membranes. The purified plasma membrane pellet was resuspended in 1 ml of plasma membrane resuspension buffer and frozen and thawed in order to convert right-side-out vesicles to inside-out vesicles (Brightman and Morré, 1992).
Antibodies
Antibody for immunoprecipitation and Western blotting (designated antibody A) was raised against a conserved region (aa 340–650) of the plasma membrane H+-ATPase AHA1 gene from Arabidopsis thaliana (Harper et al., 1989). The gene sequence was amplified by PCR, subcloned into the fusion protein cloning vector, pMAL-c2 (New England BioLabs) and over-expressed in E. coli BL21 (DE3). The resultant fusion protein, consisting of the H+-ATPase recombinant fragment linked though a Factor Xa proteolytic cleavage site to maltose-binding protein, was purified by amylose column chromatography (New England BioLabs protocol). The fusion protein was cleaved with Factor Xa (New England BioLabs protocol), and the ATPase recombinant fragment was purified by amylose column chromatography and used as antigen for the generation of polyclonal antibodies in rabbit.
A second H+-ATPase antibody used for Western blotting only (designated antibody B), which was raised against aa 390–662 of the protein, was a gift from Dr R Serrano (European Molecular Biology Laboratory, Heidelberg, Germany).
Immunopurification of plasma membrane vesicles
Magnetic, polystyrene beads (Dynabeads M-280, Dynal) with sheep anti-Rabbit IgG covalently bound to the surface were used for immunopurification of plasma membrane vesicles. Microsomal membrane fraction was diluted with microsomal resuspension buffer (50 mM EPPS pH 7.4, 0.25 M sorbitol, 10 mM EDTA, 2 mM EGTA, and 1 mM DTT) to a final concentration of 2 mg protein ml-1 and mixed with 200 µl of H+-ATPase antibody serum. The mixture was incubated for 30 min at 4 °C on a rotating shaker and then washed four times with microsomal resuspension buffer to remove all unbound antibody. The membrane–antibody complex recovered by centrifugation was resuspended in 600 µl of microsomal resuspension buffer in preparation for immunoprecipitation with the magnetic beads. Prior to incubation with the antigen–antibody complex, magnetic beads were washed five times, 4 min each, in microsomal resuspension buffer containing 0.25 M NaCl and collected using the DYNAL magnetic particle concentrator (Dynal MPC). Bead suspension (500 µl) was then mixed with the membrane–antibody complex and incubated for 2 h at 4 °C with continuous mixing. The membrane-bead–antibody complex was then collected using the DYNAL magnetic concentrator and washed five times, 4 min each, in microsomal resuspension buffer containing 0.25 M NaCl. Following the final wash, the plasma membrane vesicles were eluted from the bead–antibody complex by incubation with 200 µl of 100 mM glycine (pH 2.5) for 30 min at 4 °C. Control experiments were conducted in which microsomal membrane suspension was immunoprecipitated with magnetic polystyrene beads alone without the addition of the H+-ATPase antibody.
Enzyme assays and protein analysis
H+-ATPase activity was determined indirectly by measuring proton pumping using the 
pH probe, acridine orange, as described (de Michelis et al., 1983). NADPH-cytochrome c reductase and cytochrome c oxidase activities were measured according to protocols already described (Hodges and Leonard, 1974).
Protein was quantified as described previously (Bradford, 1976) using BSA (bovine serum albumin) as a standard. Polypeptides were fractionated by SDS-PAGE in Mini Protein Dual Slab Cells (Bio-Rad, Mississauga, ON, Canada) using 12% acrylamide. The gels were either stained with silver (Wray et al., 1981) or blotted onto nitrocellulose for Western analysis (Hudak et al., 1997).
Lipid analysis
Lipids were extracted according to Bligh and Dyer (Bligh and Dyer, 1959) and fractionated by thin layer chromatography (Yao et al., 1991). The separated lipids were visualized with iodine vapour and identified using authentic standards. Fatty acids were methylated according to Morrison and Smith (Morrison and Smith, 1964) and analysed by gas-liquid chromatography (Fobel et al., 1987).
|
Results |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Marker enzyme activities
The purity of plasma membrane preparations from carnation petals and canola leaves obtained by phase partitioning was determined by measurements of vanadate-sensitive H+-ATPase, a marker enzyme for plasma membrane (Larsson et al., 1994
), and rotenone-insensitive NADH-cytochrome c reductase and cytochrome c oxidase, marker enzymes for endoplasmic reticulum and mitochondrial membranes, respectively (Hodges and Leonard, 1974). H+-ATPase activity was measured indirectly by determining the accumulation of protons in plasma membrane vesicles using acridine orange, a pH-sensitive dye (de Michelis et al., 1983). Plasma membrane purified by phase partitioning from both leaves and petals was enriched in vanadate-sensitive H+-ATPase activity by 
4-fold by comparison with corresponding microsomal membranes (Table 1). Plasma membrane and vacuolar H+-ATPases are inhibited by vanadate and nitrate, respectively (Larsson et al., 1994; de Michelis et al., 1983). In the present study, vanadate-sensitive (200 µM vanadate) proton transport constituted the major portion (75%) of the total measurable H+-ATPase activity in plasma membrane vesicles for leaves and petals, whereas nitrate-inhibited (100 mM nitrate) proton transport accounted for only 25% of the total proton transport activity (data not shown). Thus most of the proton transport capability of the phase-purified membrane preparation is attributable to vesicles of plasma membrane.
|
Marker enzymes were used to identify other contaminating membranes in the phase-purified plasma membrane preparations. The specific activities of NADPH-cytochrome c reductase and cytochrome c oxidase, marker enzymes for endoplasmic reticulum and mitochondrial membranes, respectively, were about 4-fold lower in the purified plasma membrane fraction than in corresponding microsomal membranes for leaves and petals (Table 1
). These observations together with the enrichment of vanadate-sensitive H+-ATPase activity can be interpreted as indicating that the phase-purified membrane preparations from leaves and petals are composed of predominantly plasma membrane vesicles.
Protein composition of membrane fractions
The protein compositions of phase-purified plasma membrane and corresponding microsomal membrane preparations were examined by SDS-PAGE. For both leaves and petals, the polypeptide composition of phase-purified plasma membrane preparations was clearly distinguishable from that of microsomal membranes (Fig. 1A, B). This finding is consistent with enzyme data (Table 1) indicating that the phase-purified membrane vesicle preparations are enriched in plasma membrane vesicles relative to microsomal membranes.
|
Plasma membrane vesicles were also purified from microsomal membrane preparations by immunoprecipitation with antibodies raised against the plasma membrane H+-ATPase. Two antibodies, one designated antibody A, which was raised against a recombinant fragment of the H+-ATPase corresponding to aa 340–650, and another designated antibody B raised against the aa 390–662 (Fig. 2
), were tested for their reactions against the native H+-ATPase polypeptide (100 kDa) in Western blots. Antibody A did not recognize the native H+-ATPase polypeptide in Western blots of either microsomal membranes or phase-purified plasma membrane, whereas antibody B did react with the native polypeptide (Fig. 3). Thus it is apparent that these antibodies recognize different epitopes. However, this notwithstanding, antibodies A and B both recognized common catabolites of the H+-ATPase including polypeptides at 41, 43 and 68 kDa (Fig. 3). The 41 and 43 kDa polypeptides were clearly resolved in some blots (e.g. Fig. 3B, lane 1), but in others were not well resolved (Fig. 3).
|
|
Lipid composition of immunoprecipitated membrane vesicles
Antibody A, which was raised against a recombinant fragment of the H+-ATPase protein corresponding to aa 340 through 650 and does not recognize the native H+-ATPase polypeptide, was used for immunoprecipitation in order selectively to purify vesicles of plasma membrane that contain catabolites of the H+-ATPase polypeptide rather than its native form. It was reasoned that such vesicles might also be enriched in lipid metabolites. This in fact proved to be the case. The lipid class composition of immunopurified plasma membrane vesicles proved to be quite distinct from that of corresponding phase-purified plasma membrane. They both featured a full spectrum of lipids including phospholipids, diacylglycerol, free fatty acids, triacylglycerol, and a mixture of steryl and wax esters (Fig. 4A, B). There were, however, differences between the two preparations in the relative proportions of these lipid classes. In particular, levels of lipid metabolites (i.e. diacylglycerol, free fatty acids, triacylglycerol, and steryl/wax esters) relative to phospholipid are enriched in the immunopurified plasma membrane vesicles by comparison with phase-purified plasma membrane (Table 2). Indeed, the ratio of free to esterified fatty acids is 5-fold higher in immunopurified vesicles than in corresponding phase-purified plasma membrane (Table 3). This supports the contention that the two purification procedures yield different populations of plasma membrane vesicles. It is also clear that the lipid compositions of the immunopurified plasma membrane fraction from carnation petals and canola leaves are distinguishable from that of the antibody-containing serum (Fig. 4A, B). As well, any lipid adhering non-specifically to magnetic beads incubated with microsomal membranes in the absence of H+-ATPase antibody was below the limit of detection.
|
|
|
The fatty acid composition of phase-purified and immunopurified plasma membrane was also analysed. Here again, there are large differences between the two membrane preparations. The saturated to unsaturated fatty acid ratios for the immunopurified membrane vesicles from canola leaves and carnation petals are 11-fold and 23-fold higher, respectively, than the corresponding ratios for phase-purified plasma membrane, indicating that the immunopurified membrane vesicles contain higher levels of saturated fatty acids (Table 3
). This was further illustrated by an analysis of the fatty acid composition of each lipid class. For carnation petals, the phospholipid, diacylglycerol, free fatty acid, and triacylglycerol fractions of phase-purified plasma membrane all contain high levels of the diunsaturated fatty acid, linoleic acid (18:2), and the steryl/wax ester fraction contains both linoleic acid and the triunsaturated fatty acid, linolenic acid (18:3) (Fig. 5). By contrast, corresponding lipid classes from the immunopurified vesicles contain higher levels of the saturated fatty acids, palmitic acid and stearic acid, and do not contain detectable levels of di- and polyunsaturated fatty acids (Fig. 5). Similarly, for canola leaves the phospholipid and diacylglycerol fractions from phase-purified plasma membrane were again more unsaturated than the corresponding fractions from immunopurified vesicles (Fig. 6). However, in canola the fatty acid compositions of the free fatty acid, triacylglycerol and steryl/wax ester fractions were very similar (Fig. 6).
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Partitioning of microsomal membranes in an aqueous dextran-polyethylene glycol two-phase system is a standard technique for purifying plasma membrane fractions from plant tissues (Larsson et al., 1994
). These polymers are water-soluble, and form separate phases when mixed at concentrations ranging from 5–7%. Phase partitioning relies on differences in surface properties of the vesicles rather than on differences in their size and buoyant density, and is facilitated by the presence of salt (Sandelius and Morré, 1990). During isolation, the plasma membrane vesicles preferentially partition into the polyethylene-glycol-rich upper phase, while all contaminating organellar membranes partition preferentially at the interface or into the lower phase.
In the present study, the relative abundance of plasma membrane vesicles in the phase-purified membrane fraction was assessed by measuring the levels of vanadate-sensitive proton transport using acridine orange (de Michelis et al., 1983). Acridine orange is a small, cationic dye that freely permeates membranes in its unprotonated form and can be used to monitor transmembrane pH in membrane vesicles. Since H+ translocation and ATP hydrolysis are stoichiometric, the acridine orange assay serves as a reliable measure of H+-ATPase activity (Briskin, 1990). Based on this assay, the vanadate-sensitive H+-ATPase specific activity of phase-purified plasma membrane preparations from canola leaves and carnation petals proved to be 4-fold higher than the corresponding activity of microsomal membranes. This enrichment of H+-ATPase activity can be interpreted as reflecting purification of plasma membrane.
There are two P-type proton-translocating ATPases, one localized on the tonoplast and the other on the plasma membrane (de Michelis et al., 1983). Apart from their different subcellular localizations, these two ATPases show differential sensitivity to inhibitors: the plasma membrane ATPase is specifically inhibited by vanadate, whereas the tonoplast ATPase is unaffected by vanadate and strongly inhibited by NO3- (de Michelis et al., 1983). In the present study, these inhibitors were used to distinguish between tonoplast and plasmalemma in the phase-purified membrane preparations. Vanadate-sensitive proton transport constituted 75% of the total ATP-dependent proton transport in the purified membrane fraction, whereas nitrate-inhibited proton transport accounted for only 25%. These data are in accordance with previously published results (Larsson et al., 1994) and indicate that the majority of membrane vesicles obtained by phase-partitioning are of plasma membrane origin.
Marker enzymes were used to identify the presence of other contaminating organellar membranes in the plasma membrane preparation. Cytochrome c oxidase activity was measured to assess contamination by mitochondrial membrane, and NADPH-cytochrome c reductase was assayed as a marker for endoplasmic reticulum (Hodges and Leonard, 1974). The specific activities of both markers were about 4-fold lower in the purified plasma membrane fraction than in corresponding microsomal membranes. These findings indicate a relatively low abundance of membrane vesicles originating from mitochondria and endoplasmic reticulum in the purified plasma membrane fraction.
Plasma membrane vesicles were also isolated from preparations of microsomal membrane vesicles by immunopurification using a polyclonal antibody (Antibody A) raised against a recombinant fragment of the H+-ATPase. Two different antibodies (A and B) were tested by Western blotting for their ability to cross-react with the H+-ATPase polypeptide. Antibody B, which was raised against a central region of the H+-ATPase corresponding to aa 390–662, recognized the native H+-ATPase as well as a number of lower molecular weight catabolites of the protein. By contrast, antibody A, which was raised against the central region of the H+-ATPase corresponding to aa 340–650, recognized three of the same lower molecular weight catabolites of the H+-ATPase (68, 43 and 41 kDa), but did not recognize the native polypeptide. Thus the catabolites of the H+-ATPase apparently possess epitopes that are not exposed on the native protein. The finding that antibody A did not recognize the native protein may also reflect the fact that it was raised against a denatured antigen. The immunopurified plasma membrane vesicles are presumably inside-out since the antibody was raised against the hydrophilic domain of the H+-ATPase projecting into the cytoplasm in intact cells.
The finding that antibody A recognizes the same catabolites as antibody B and no other polypeptides indicates that it is specific for the H+-ATPase. This contention is further supported by the fact that the same antibody has been previously shown to recognize antigens in both the plasma membrane and the cytosol in intact cells of carnation petals (Hudak et al., 2000). This argues against the possibility that the membrane-associated catabolites of the native H+-ATPase recognized by antibody A in both purified plasma membrane and microsomes are polypeptides of cytosolic origin unrelated to the H+-ATPase that become associated with the membranes as a result of tissue homogenization. Indeed, the fact that antibody A does recognize antigens in situ in both the plasma membrane and the cytosol is consistent with the finding in the present study that H+-ATPase catabolites are present in cytosolic microvesicles.
Antibody A was used for immunoprecipitation in an effort to selectively purify vesicles of plasma membrane that are enriched in membrane lipid metabolites. This strategy was based on the assumption that vesicles of plasma membrane containing the proteolytic catabolites of the H+-ATPase rather than the native protein might also be enriched in lipid metabolites. This proved to be the case. Indeed, the lipid compositions of phase-purified plasma membrane and immunopurified plasma membrane were clearly distinguishable. In particular, the immunoprecipitated plasma membrane proved to be enriched in lipid metabolites including free fatty acids, steryl and wax esters and diacylglycerol, whereas the phase-purified plasma membrane consisted primarily of phospholipid. The possibility that the enrichment of lipid metabolites relative to phospholipid in the immunopurified plasma membrane vesicles simply reflects leakage (i.e. selective partitioning) of phospholipid out of the vesicles during homogenization appears not to be the case. The fatty acid composition of lipid metabolites in the immunopurified fraction are clearly distinguishable from those in the corresponding phase purified plasma membrane. Specifically, the diacylglycerol, free fatty acids, steryl and wax esters and triacylglycerol of the immunopurified vesicles contain much higher proportions of saturated fatty acids than were found in the corresponding lipid classes of phase-purified plasma membrane. Indeed, the saturated:unsaturated fatty acid ratios for immunopurified plasma membrane vesicles from canola leaves and carnation petals proved to be 11-fold and 23-fold higher, respectively, than the corresponding ratios for phase-purified plasma membrane. If the enrichment of lipid metabolites relative to phospholipid in the immunopurified vesicles were simply due to phospholipid leakage, there should not be a difference in the fatty acid profiles of the lipid metabolites.
The presence of triacylglycerol in both phase-purified and immunopurified vesicles of plasma membrane is surprising in light of the fact that diacylglycerol acyl transferase, the enzyme catalysing the terminal step in triacylglycerol synthesis, is thought to be associated with the endoplasmic reticulum (Settlage et al., 1995). This may reflect inclusion of triacylglycerol in the bilayers of microvesicles originating from the endoplasmic reticulum that serve to target proteins to the plasmalemma.
The finding that the immunopurified plasma membrane vesicles contain high levels of these lipid metabolites is of interest, for they are all bilayer-perturbing lipids. It is known, for example, that free fatty acids behave as detergents (Thomas, 1982). Diacylglycerols act as membrane-destabilizing agents and promote microvesiculation (Allan et al., 1976). As well, some of the lipids enriched in the immunopurified plasma membrane vesicles, specifically free fatty acids and steryl/wax esters, are gel-phase forming lipids (Yao et al., 1991), and their presence in membranes would thus lead to lipid phase separations. The immunopurified vesicles from carnation petals, for example, proved to be enriched in free fatty acids and steryl/wax esters by 14-fold and 40-fold, respectively, by comparison with corresponding phase-purified plasma membrane.
One likely interpretation of these findings is that microsomal fractions, while being comprised in the main of vesicles formed during tissue homogenization, also contain vesicles originating from lipid metabolite-enriched domains within the plasmalemma that are released in situ by microvesiculation as part of normal membrane turnover. Microvesiculation is well established as an inherent feature of membrane trafficking involved in both targeting of newly synthesized proteins (Robinson et al., 1998) and endocytosis (Mellman, 1996), but has not been previously implicated in membrane turnover. It is also possible that the vesiculation of lipid metabolite-enriched domains from intact plasmalemma is facilitated by tissue homogenization. In any case, the observations constitute evidence for the existence of discrete domains of lipid metabolites within the plane of the plasma membrane. It has been demonstrated that in senescing membranes these lipid metabolites accumulate, forming domains of sufficient size and frequency to be detectable by X-ray diffraction and freeze-fracture electron microscopy (Paliyath and Thompson, 1990; Yao et al., 1991). Such domains, however, are not detectable in membranes from non-senescing tissues suggesting that they are removed from the bilayer during normal membrane turnover.
It is unlikely that the presence of elevated levels of lipid metabolites in the immunopurified plasma membrane vesicles is attributable to the action of a lipase released during homogenization. First, plant and animal lipases appear to be cytosolic enzymes with access to the plasma membrane in the intact cell (Hong et al., 2000). Second, the lipid composition of the immunopurified vesicles differs from that of the plasmalemma in ways that cannot be explained simply by the action of one or more lipases. Specifically, vesicles of plasma membrane formed during tissue homogenization would have the same lipid composition as the plasmalemma, whereas the immunopurified vesicles are enriched in free fatty acids, in steryl/wax esters and in triacylglycerol by comparison with phase-purified plasma membrane, and have a much higher lipid saturation index. Although the action of a lipase on the vesicles during their isolation could give rise to increased levels of free fatty acids, it would not account for the other differences in lipid composition. Rather, the observations are more consistent with the contention that these lipid metabolites once formed move laterally through the plane of the bilayer to form metabolite-enriched domains, which are subsequently released from the membrane as microvesicles.
In an earlier study, lipid particles containing phospholipid and enriched in the same lipid metabolites found in immunopurified plasma membrane vesicles were isolated from the cytosol of carnation petals (Hudak and Thompson, 1996). These cytosolic lipid particles proved to be particularly enriched in free fatty acids and steryl/wax esters by comparison with corresponding microsomal membranes. Similar lipid particles have also been isolated from the stroma of chloroplasts (Ghosh et al., 1994). The lipid particles appear to be formed by blebbing from the surface of membranes in much the same way that oil bodies are released from the endoplasmic reticulum (Huang, 1996), and it has been postulated that their release from membranes allows removal of lipid metabolites that would otherwise destabilize the bilayer (Thompson et al., 1998). However, the carnation cytosolic lipid particles are clearly distinguishable from immunopurified plasma membrane vesicles from the same tissue in that they have a much lower buoyant density. Specifically, the lipid particles were isolated by flotation centrifugation of a microsomal supernatant (cytosol fraction) that was made 10% (w/v) with sucrose to increase its buoyant density and then centrifuged for 12 h at 305 000 g (Hudak and Thompson, 1996). By contrast, carnation plasma membrane vesicles enriched in the same lipid metabolites were immunopurified from microsomal vesicles pelleted by centrifugation of a 10 000 g-20 min supernatant for 1 h at 250 000 g, and their higher buoyant density presumably reflects higher levels of protein than are present in cytosolic lipid particles. These observations collectively suggest that discrete lipid metabolite-enriched domains differing in composition and perhaps also size are formed within membrane bilayers, and that some of these domains are released from the bilayer by microvesiculation and others by blebbing of lipid particles from the membrane surface.
|
Acknowledgments |
|---|
This work was supported by the Natural Sciences and Engineering Research Council of Canada. The authors also thank Dr R Serrano (European Molecular Biology Laboratory, Heidelberg, Germany) for providing the H+-ATPase antibody.
|
Notes |
|---|
1 Present address: Skye Pharmatech Inc., 6354 Viscount Rd., Mississauga, Ontario, Canada L4V 1H3.
2 Present address: Biotech Center, Cook College, Rutgers University, New Brunswick, New Jersey 08901-8520, USA
3 To whom correspondence should be addressed. Fax: +1 519 746 2543. E-mail: JET{at}sciborg.uwaterloo.ca
|
Abbreviations |
|---|
HEPES, (N-[2-hydroxyethyl]piperazine-N'-[2-ethanesulphonic acid]); EPPS (N-2-hydroxyethylpiperazine-N'-3-propanesulphonic acid; DTT (dithiothreitol), EDTA (ethylenediaminetetra-acetic acid); EGTA, ethylene glycol-bis(ß-aminoethyl ether)-N,N,N,N'-tetra-acetic acid; PVPP (polyvinylpoly-pyrrolidone); PMSF (phenylmethylsulphonyl fluoride)..
|
References |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Allan D, Billah MM, Finean JB, Michell RH.1976. Release of diacylglycerol-enriched vesicles from erythrocytes with increased intracellular Ca2+. Nature 261, 58–60.[Medline]
Bligh EG, Dyer WJ.1959. A rapid method of total lipid extraction and purification. Canadian Journal of Biochemistry and Physiology 37, 911–917.
Boutry M, Michelet B, Goffeau A.1989. Molecular cloning of a family of plant genes encoding a protein homologous to plasma membrane H+-translocating ATPases. Biochemical and Biophysical Research Communications 162, 567–574.[Web of Science][Medline]
Bradford MM.1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72, 248–254.[Web of Science][Medline]
Brightman AO, Morré DJ.1992. Sidedness of plant plasma membrane vesicles altered by conditions of preparation. Plant Physiology 98, 183–190.
Briskin DP.1990. The plasma membrane H+-ATPase of higher plant cells: biochemistry and transport function. Biochimica et Biophysica Acta 1019, 95–109.
Chapman KD.1998. Phospholipase activity during plant growth and development and in response to environmental stress. Trends in Plant Science 3, 419–426.
Curti G'massardi F, Lado P.1993. Synergistic activation of plasma membrane H+-ATPase in Arabidopsis thaliana cells by turgor decrease and by fusicoccin. Physiologia Plantarum 87, 592–600.
de Michelis MI, Pugliarello MC, Rasi-Caldogno F.1983. Two distinct proton translocating ATPase are present in membrane vesicles from radish seedlings. FEBS Letters 162, 85–90.
DeWitt ND, Sussman MR.1995. Immunocytological localization of an epitope-tagged plasma membrane proton pump (H+-ATPase) in phloem companion cells. The Plant Cell 7, 2053–2067.[Abstract]
Duxbury CL, Legge RL, Paliyath G, Thompson JE.1991. Lipid breakdown in smooth microsomal membranes from bean cotyledons alters membrane proteins and induces proteolysis. Journal of Experimental Botany 42, 103–112.
Ewing NN, Bennett AB.1994. Assessment of the number and expression of P-type H+-ATPase genes in tomato. Plant Physiology 106, 547–557.[Abstract]
Ewing NN, Wimmers LE, Meyer DJ, Chetelat RT, Bennett AB.1990. Molecular cloning of tomato plasma membrane H+-ATPase. Plant Physiology 94, 1874–1881.
Fobel M, Lynch DV, Thompson JE.1987. Membrane deterioration in senescing carnation flowers. Plant Physiology 85, 204–211.
Ghosh S, Hudak KA, Dumbroff EB, Thompson JE.1994. Release of photosynthetic protein catabolites by blebbing from thylakoids. Plant Physiology 106, 1547–1553.[Abstract]
Hare JF.1990. Mechanisms of membrane turnover. Biochimica et Biophysica Acta 1031, 71–90.[Medline]
Harper JF, Manney L, DeWitt ND, Yoo MH, Sussman MR.1990. The Arabidopsis thaliana plasma membrane H+-ATPase multigene family: genomic sequence and expression of third isoform. Journal of Biological Chemistry 265, 13601–13608.
Harper JF, Manney L, Sussman MR.1994. Evidence for at least ten plasma membrane H+-ATPase genes in Arabidopsis, and complete genomic sequence of AHA 10 which is expressed primarily in developing seeds. Molecular and General Genetics 244, 572–587.
Harper JF, Surowy TK, Sussman MR.1989. Molecular cloning and sequence of cDNA encoding the plasma membrane proton pump (H+-ATPase) of Arabidopsis thaliana. Proceedings of the National Academy of Sciences, USA 86, 1234–1238.
Hodges TK, Leonard RT.1974. Purification of a plasma membrane–bound adenosine triphosphatase from plant roots. Methods in Enzymology 32, 392–406.[Medline]
Hong Y, Wang TW, Hudak KA, Schade F, Froese CD, Thompson JE.2000. An ethylene-induced cDNA encoding a lipase expressed at the onset of senescence. Proceedings of the National Academy of Sciences, USA (in press).
Houlne G, Boutry M.1994. Identification of an Arabidopsis thaliana gene encoding a plasma membrane H+-ATPase whose expression is restricted to anther tissues. Plant Journal 5, 311–317.[Web of Science][Medline]
Huang AHC.1996. Oleosins and oil bodies in seeds and other organs. Plant Physiology 110, 1055–1061.[Web of Science][Medline]
Hudak KA, Ghosh S, Dumbroff DB, Thompson JE.1997. Cytosolic catabolites of H+-ATPase in senescing carnation petals. Phytochemistry 44, 371–376.
Hudak KA, Madey E, Hong Y, Su L, Thompson JE.2000. Immunopurification of H+-ATPase-containing lipid-protein particles from the cytosol of carnation petals. Physiologia Plantarum 109, 304–312.
Hudak KA, Thompson JE.1996. Flotation of lipid-protein particles containing triacylglycerol and phospholipid from the cytosol of carnation petals. Physiologia Plantarum 98, 810–818.
Kim DK, Lee HJ, Lee Y.1994. Detection of two phospholipase A2 (PLA2) activities in leaves of higher plant Vicia faba and comparison with mammalian PLA2's. FEBS Letters 343, 213–218.[Web of Science][Medline]
Kjellbom P, Larsson C.1984. Preparation and polypeptide composition of chlorophyll-free plasma membranes from leaves of light-grown spinach and barley. Physiologia Plantarum 62, 501–509.
Kurkdjian A, Guern J.1989. Intracellular pH: measurement and importance in cell activity. Annual Review of Plant Physiology and Plant Molecular Biology 40, 271–303.[Web of Science]
Larsson C, Sommarin M, Widell S.1994. Isolation of highly purified plant plasma membranes and separation of inside-out and right-side-out vesicles. Methods in Enzymology 228, 451–469.
Leonard RT.1984. Membrane associated ATPases and nutrient absorption by roots. In: Tinker PB, Lauchli A, eds. Advances in plant nutrition. Praeger Scientific Publishers, 209–240.
Mazliak P.1980. Synthesis and turnover of plant membrane phospholipids. Progress in Phytochemistry 6, 49–102.
Mellman I.1996. Endocytosis and molecular sorting. Annual Review in Cellular and Developmental Biology 12, 575–625.[Web of Science][Medline]
Moriau L, Bogaerts P, Jonniaux J-L, Boutry M.1993. Identification and characterization of a second plasma membrane H+-ATPase gene subfamily in Nicotiana plumbaginifolia. Plant Molecular Biology 21, 955–963.[Web of Science][Medline]
Morrison WR, Smith LM.1964. Preparation of fatty acid methyl esters and dimethylacetals from lipids with boron trifluoride-methanol. Journal of Lipid Research 5, 600–608.[Abstract]
Ohlrogge J, Browse J.1995. Lipid biosynthesis. The Plant Cell 7, 957–970.[Web of Science][Medline]
Paliyath G, Thompson JE.1990. Evidence for early changes in membrane structure during postharvest development of cut carnation flowers. New Phytologist 114, 555–562.
Pardo JM, Serrano R.1989. Structure a plasma membrane H+-ATPase gene from the plant Arabidopsis thaliana. Journal of Biological Chemistry 264, 8551–8562.
Rayle DL, Cleland R.1992. The acid growth theory of auxin-induced cell elongation is alive and well. Plant Physiology 99, 1271–1274.
Robinson DG, Hinz G, Holstein SE.1998. The molecular characterization of transport vesicles. Plant Molecular Biology 38, 49–76.[Web of Science][Medline]
Sandelius AS, Morré DJ.1990. Plasma membrane isolation. In: Larsson C, Moller IM, eds. The plant plasma membrane. Berlin: Springer-Verlag, 44–75.
Sandelius AS, Sommarin M.1990. Membrane localized reactions involved in polyphosphoinositide turnover in plants. In: Morré DJ, ed. Inositol metabolism in plants. New York: Wiley-Liss, 139–161.
Settlage SB, Wilson RF, Kwanyuen P.1995. Localization of diacylglycerol acyl transferase to oil body associated endoplasmic reticulum. Plant Physiology and Biochemistry 33, 399–407.
Steer MW.1988. Plasma membrane turnover in plant cells. Journal of Experimental Botany 39, 987–996.
Thomas H.1982. Control of chloroplast demolition during leaf senescence. In: Waring PF, ed. Plant growth substances. London: Academic Press, 559–567.
Thompson JE, Froese CD, Madey E, Smith MD, Hong Y.1998. Lipid metabolism during plant senescence. Progress in Lipid Research 37, 119–141.[Web of Science][Medline]
Toshiyuki F, Pak J-Y, Ohwaki Y, Tsujimura H, Nitta T.1996. Tissue-specific expression of the gene for a putative plasma membrane H+-ATPase in a seagrass. Plant Physiology 110, 35–42.[Abstract]
Wada M, Makoto T, Kasamo K.1992. Nucleotide sequence of a complementary DNA encoding plasma membrane H+-ATPase form rice (Oryza sativa L.). Plant Physiology 99, 794–795.
Wang X, Dyer JH, Zheng L.1993. Purification and immunological analysis of phospholipase D form castor bean endosperm. Archives of Biochemistry and Biophysics 306, 486–494.[Web of Science][Medline]
Wray WW, Bovikas T, Wray VP, Hancock R.1981. Silver staining of proteins in polyacrylamide gels. Analytical Biochemistry 118, 197–203.[Web of Science][Medline]
Yao K, Paliyath G, Thompson JE.1991. Non-sedimentable microvesicles from senescing bean cotyledons contain gel phase-forming phospholipid degradation products. Plant Physiology 97, 502–508.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  W. G. van Doorn and E. J. Woltering Physiology and molecular biology of petal senescence J. Exp. Bot., March 3, 2008; (2008) erm356v2. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Regente, G. C. Monzon, and L. de la Canal Phospholipids are present in extracellular fluids of imbibing sunflower seeds and are modulated by hormonal treatments J. Exp. Bot., February 1, 2008; 59(3): 553 - 562. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. L. Jeong, H. Jiang, H.-S. Chen, C.-J. Tsai, and S. A. Harding Metabolic Profiling of the Sink-to-Source Transition in Developing Leaves of Quaking Aspen Plant Physiology, October 1, 2004; 136(2): 3364 - 3375. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||