Identification and characterization of nonsedimentable lipid-protein ...

Proc. Nati. Acad. Sci. USA Vol. 88, pp. 2269-2273, March 1991 Botany

Identification and characterization of nonsedimentable lipid-protein microvesicles K. YAO*, G. PALIYATH*, R. W. HUMPHREYt, F. R. HALLETTt,

AND

J. E. THOMPSON*

Departments of *Horticultural Science, tMicrobiology, and tPhysics, University of Guelph, Guelph, ON NIG 2W1, Canada

Communicated by Kenneth V. Thimann, December 6, 1990

ABSTRACT Previously uncharacterized lipid-protein microvesicles have been isolated from young and senescing bean cotyledon tissue. The microvesicles are nonsedimentable and enriched in phospholipid degradation products (free fatty acids, long-chain aldehydes, and long-chain hydrocarbons). They range from 70 to 170 nm (radius) with a mean radius of 132 nm, and it is clear from freeze-fracture electron micrographs that they are bilayered in nature. Nonsedimentable lipid-protein microvesicles containing the same products of phospholipid degradation but smaller were also formed in vitro when smooth microsomal membranes from young cotyledon tissue were treated with Ca2' to stimulate enzymatic degradation of phospholipids. The data suggest that these microvesices comprise an intermediate stage of membrane lipid deterioration. They appear to serve as a vehicle for moving phospholipid degradation products out of membranes into the cytosol during senescence and perhaps also during normal membrane lipid turnover.

established, although there have been some reports of microvesiculation from membranes under conditions of phospholipid catabolism. It has been established, for example, that treatment of erythrocytes with exogenous phospholipase C or calcium ionophore results in the formation of microvesicles that are enriched in diacylglycerol (20, 21). Vesicle formation has also been visualized ultrastructurally in senescing cowpea cotyledons (4) and in wheat leaf cells exposed to drought (3). In the present study, we have isolated nonsedimentable lipid-protein microvesicles from bean cotyledon tissue that appear to be an intermediate stage of membrane deterioration inasmuch as they are enriched in phospholipid degradation products. They can also be formed in vitro from isolated membranes under conditions in which phospholipid degradation in the membranes is stimulated by c2+. Ca2~

Membrane lipid deterioration is an inherent feature of plant senescence and of capitulation of plant tissues to certain types of stress including drought and freezing (1-4). One of the clearest manifestations of this is a progressive decline in phospholipid phosphate and fatty acids resulting in an increase in the sterol/esterified fatty acid ratio in membranes and a corresponding decrease in membrane bulk lipid fluidity (1). There is also an accumulation of free fatty acids and peroxidized lipids in senescing membranes that alter the phase properties of membrane lipids and introduce packing perturbations in the bilayer that are thought to facilitate enzymatic degradation of lipids (1, 5-7). In addition, membrane phospholipids have been shown to have relatively rapid turnover rates ranging in half-life from 1 to 10 hr (8). Several lipolytic enzymes that could participate in membrane lipid turnover and in the net degradation of membrane lipids that accompanies senescence and capitulation to stress have been found associated with plant membranes. Phospholipase D activity has been detected in isolated fractions of endoplasmic reticulum, Golgi membranes, tonoplast, and plasmalemma (9-12), and it has been proposed that the membrane-associated form of the enzyme participates in lipid turnover (13). Phosphatidic acid phosphatase has been found on endoplasmic reticulum and in association with other isolated membrane fractions (14-16), and there are reports of lipolytic acyl hydrolase being associated with microsomal membranes (17, 18). Moreover, a recent study has indicated that phospholipase D, phosphatidic acid phosphatase, and lipolytic acyl hydrolase are all enriched in the sedimentable material obtained after partial solubilization of microsomes in Triton X-100, suggesting that they are tightly associated with the membranes (19). The precise mechanism by which phospholipid degradation products are removed from membranes has not been

Plant Material and Isolation of Microvesicles. Bean seeds (Phaseolus vulgaris L. cv. Kinghorn) were germinated in vermiculite at 290C and 90o relative humidity under conditions of etiolation. The cotyledons were harvested 2, 4, and 7 days after planting and were homogenized (17%; wt/vol) in 50 mM NaHCO3/0.3 M sucrose, pH 7.0, for 45 s in an Omnimixer and for an additional 1 min in a Polytron homogenizer at 40C. The homogenate was filtered through four layers of cheesecloth and centrifuged at 10,000 x g for 20 min. The resulting supernatant was centrifuged at 130,000 X g for 1 hr to yield a postmicrosomal supernatant. Nonsedimentable lipid-protein microvesicles were isolated by centrifuging the postmicrosomal supernatant at 250,000 x g for 12 hr and concentrating the resulting supernatant to 15 ml by filtration through a Pharmacia 300-kDa cut-off filter. The resulting suspension of microvesicles was washed three times with 30 ml of homogenizing buffer using the same filter. In Vitro Formation of Nonsedimentable Microvesides. Nonsedimentable lipid-protein microvesicles were formed in vitro from smooth microsomal membranes of2-day-old cotyledons. The smooth microsomal membranes were isolated by centrifugation of a 10,000 x g 20-min supernatant (obtained from a homogenate of 2-day-old cotyledons as described above) through an 8-ml cushion of 1.5 M sucrose/50 mM NaHCO3, pH 7.0, at 140,000 x g for 1.5 hr with a Beckman SW 28 rotor. The smooth microsomal membranes formed a band at the interface, and after centrifugation they were removed, diluted four times with homogenizing buffer, and pelleted by centrifugation at 130,000 x g for 1 hr. The membrane pellet was resuspended in homogenizing buffer (0.2 mg of protein per ml) and incubated with gentle shaking at 230C for specified periods in the presence of CaCl2 (30 ,uM or 1 mM) and in some cases streptomycin (25 ptg/ml) or 0.2 mM EGTA/2 mM EDTA/25 ,4M calmidazolium to form microvesicles. At the end of the incubation period, the reaction mixture was centrifuged at 130,000 x g for 1 hr to sediment the membranes, and the resulting supernatant was centrifuged at 250,000 x g for 12 hr. This final supernatant, which contained the microvesicles,

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MATERIALS AND METHODS

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was concentrated to 10 ml by filtration through a Pharmacia 300-kDa cut-off filter and washed three times with 20 ml of homogenizing buffer using the same filter. Analytical Procedures. Lipid extracts from microvesicles or membranes were analyzed by thin-layer chromatography. Lipids were extracted as described (22) and spotted onto silica gel 60 plates (Mandel, Guelph, Ontario). The plates were developed halfway in chloroform/acetic acid/methanol/water (70:25:5:2; vol/vol), dried, and completely developed in petroleum ether/diethyl ether/acetic acid (70:30:1; vol/vol). The separated lipids were visualized with iodine vapor or by sulfuric acid charring, and the plates were then photographed. Free fatty acids were identified by using linoleic acid, longchain aldehydes with octadecanal, and long-chain hydrocarbons with octadecane. Phospholipids and diacylglycerol were identified by using authentic standards. Gel chromatography on Superose-12 was carried out with a 40 x 2.5 cm column. Concentrated, washed microvesicles (10-mg protein equivalents) were loaded on the column and eluted with 50 mM NaHCO3/0.3 M sucrose, pH 7.0. Proteins were quantified according to Bradford (23) and fractionated by SDS/PAGE on 8-16% linear gradients as described by Laemmli (24). The protein gels were stained with silver (25). Light Scattering Measurements and Electron Microscopy. The isolated microvesicles were sized as described (26) by dynamic light scattering measurements made at 230C using light from a helium/neon laser (model 125; Spectra-Physics) that was focused into a thermally jacketed scattering chamber. Freeze fracture was performed in a Balzers model BA360M freeze-etching unit operating at -100'C and 1.33 x 10-8 Pa (1 x 10-6 torr). The etching time was 30 s, and Pt-shadowed, C-coated replicas were produced by a standard procedure. For negative staining, a droplet of microvesicle suspension was placed on a nickel grid that had been glowdischarged to make it wettable, and the sample was dried with the edge of a piece of filter paper and stained for 5 s with 2% uranyl acetate. All samples were examined with a Philips EM300 operating at 60 kV with the anticontamination trap in place.

RESULTS Isolation and Chemical Characterization of Nonsedimentable Lipid-Protein Microvesicles. Cytosolic fractions (postmicrosomal supernatants) isolated from 2-, 4-, and 7-day-old bean cotyledon tissue contained lipid phosphate that was not sedimented during subsequent protracted high-speed centrifugation (250,000 x g for 12 hr) designed to remove residual membrane. When the 250,000 X g 12-hr supernatant was concentrated to 15 ml and washed three times with 2 vol of homogenizing buffer by filtration through a 300-kDa cut-off filter, the lipid phosphate did not pass through the filter, suggesting that it was assembled in lipid-protein microvesicles. This was further substantiated by the finding that nonsedimentable lipid and protein in the concentrated, washed supernatant coeluted in the void volume during chromatographic fractionation on a Superose-12 column (Fig. 1A). Thin-layer chromatography of the pooled lipid extracts for fractions 11-16 and 17-27, respectively, obtained from the Superose-12 column indicated that the microvesicles were enriched in free fatty acids, long-chain hydrocarbons, and to a lesser extent long-chain aldehydes (Fig. 2A), which are phospholipid degradation products. Free fatty acids and long-chain hydrocarbons were evident in both of the pooled microvesicle fractions (Fig. 2A, lanes 1 and 2), but, as indicated in Fig. 1A, it was clear that the microvesicles do not have a uniform lipid composition. Specifically, microvesicles in fractions 17-27 from the Superose-12 column contained more lipid phosphate than those in fractions 11-16 from the

Proc. Natl. Acad. Sci. USA 88 (1991)

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FIG. 1. Chromatographic fractionation of nonsedimentable lipidprotein microvesicles on Superose-12. (A) Microvesicles isolated from 7-day-old cotyledons. (B) Microvesicles formed in vitro in the presence of 1 mM Ca2+ from smooth microsomal membranes of 2-day-old cotyledons. Ei Protein; *, lipid phosphate. Data are from one of four separate experiments showing the same results.

column (Fig. 2A). The data in Fig. 2A are for nonsedimentable microvesicles from 7-day-old cotyledons, but the same lipid degradation products are also present in microvesicles isolated from 2- and 4-day-old cotyledons (data not shown). Values for the ratio of microvesicle phospholipid to corresponding microsomal phospholipid ranged from 0.11 ± 0.03 (SE; n = 3) for young 2-day-old cotyledons to 0.14 ± 0.03 (SE; n = 3) for senescent 7-day-old cotyledons. This suggests that the microvesicles do not accumulate in the tissue with advancing senescence and may be degraded. However, from densitometer scans of the thin-layer chromatograms of lipid extracts from microvesicles it is apparent that the ratio of degradation products (free fatty acids, long-chain hydrocarbons, and long-chain aldehydes) to phospholipid is =3-fold higher in microvesicles from senescent 7-day-old cotyledon tissue than in those from young 2-day-old cotyledon tissue. SDS/PAGE indicated that, notwithstanding these differences in lipid composition, the polypeptide profiles of the two pooled chromatographic fractions of 7-day-old microvesicles are similar to each other and to that of the unfractionated microvesicles (Fig. 3A, lanes 2-4). There were, however, clear differences in protein composition between the microvesicles and smooth microsomal membranes isolated from the same tissue. In particular, a number of large proteins (>90 kDa) detectable in gels for the membranes were not evident in gels for corresponding microvesicles, and a number of intense smaller molecular mass bands in the membrane gels were either not present or were much less intense in the microvesicle gels (Fig. 3A). There was also smearing in the lower region of the microvesicle gels and protein stain reflecting discrete small polypeptide fragments that were not evident in corresponding membrane gels (Fig. 3A). These features suggest that the microvesicles contain partially degraded proteins. In Viro Formation of Nonwdimentable Lipid-Protein Microvesicles. The lipid composition of these microvesicles raised the possibility that they originate from membranes and may be an intermediate stage of membrane deterioration. This was examined by incubating smooth microsomal membranes isolated from young 2-day-old cotyledons for various periods of time in the presence of exogenous Ca2+ (30 ,uM to 1 mM). This treatment induced release of lipid phosphate and

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FIG. 2. Thin-layer chromatography of lipid extracts from nonsedimentable lipid-protein microvesicles. (A) Microvesicles isolated from 7-day-old cotyledons and fractionated on Superose-12. Lanes: 1, pooled lipid extract from fractions 11-16 from the Superose-12 column (lipid equivalents from 0.80 mg of protein); 2, pooled lipid extract for fractions 17-27 from the Superose-12 column (lipid equivalents from 0.80 mg of protein). (B) Microvesicles formed in vitro in the presence of 1 mM Ca2+ from smooth microsomal membranes of 2-day-old cotyledons. Lanes: 1, lipid extract from 1.6-mg protein equivalents of membrane at time 0; 2, lipid extract from 1.6-mg protein equivalents of membrane after 1 hr of Ca2+ treatment; 3-5, lipid extracts from 0.3-mg protein equivalents of microvesicles after 1, 2, and 9 hr, respectively, of Ca2+ treatment. HC, long-chain hydrocarbons; ALD, long-chain aldehydes; FFA, free fatty acids; PA, phosphatidic acid; PE/PG, phosphatidylethanolamine/ phosphatidylglycerol; PC, phosphatidylcholine; PI, phosphatidylinositol; DG1,3, 1,3-diacylglycerol; DG1,2, 1,2-diacylglycerol. Asterisks denote unidentified compounds. Data are from one of four separate experiments showing the same results. The separated lipids were visualized with iodine vapor. (Similar chromatograms were obtained when the separated lipids were visualized by sulfuric acid charring.)

protein into the supernatant that were not sedimented by centrifugation at 250,000 x g for 12 hr and did not pass

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-Ij14 FIG. 3. SDS/PAGE of nonsedimentable lipid-protein microvesicles. Each lane contained 5 ,ug of protein, and the gels were stained with silver. (A) Microvesicles isolated from 7-day-old cotyledons. Lanes: 1, smooth microsomal membranes from 7-day-old cotyledons; 2, unfractionated microvesicles; 3, pooled microvesicles of fractions 11-16 from the Superose-12 column; 4, pooled microvesicles of fractions 17-27 from the Superose-12 column. (B) Microvesicles formed in vitro in the presence of 1 mM Ca2+ from smooth microsomal membranes of 2-day-old cotyledons. Lanes: 1 and 2, smooth microsomal membranes from 2-day-old cotyledons before and after Ca2+ treatment, respectively; 3, microvesicles. Data are from one of three separate experiments showing the same results. Molecular size markers (kDa) are indicated.

through a 300-kDa cut-off filter. The nonsedimentable lipid phosphate and protein released from the membranes also coeluted in the void volume (fractions 11-22) during chromatographic fractionation on Superose-12, suggesting that they were assembled in microvesicles (Fig. 1B). Thin-layer chromatography of lipid extracts from the putative microvesicles formed in vitro indicated that they are enriched in free fatty acids, long-chain aldehydes, and longchain hydrocarbons relative to the membranes from which they were derived. These phospholipid degradation products were not detectable in lipid extracts from 1.6-mg protein equivalents of membrane at time 0 or after 1 hr of treatment with 1 mM Ca2+ (Fig. 2B, lanes 1 and 2), whereas the degradation products were clearly evident in the lipid extracts of 0.3-mg protein equivalents of nonsedimentable microvesicles (lanes 3-5). Levels of microvesicular lipid phosphate released from the membranes in the presence of 1 mM Ca2+ increased during the first 2 hr of treatment and thereafter remained unchanged for up to 9 hr (Fig. 4, bars A-C), were unaffected by inclusion of streptomycin (25 ,g/ml) in the incubation medium (bars C and D), were essentially similar whether 30 ,uM or 1 mM Ca2+ was used (bars C and E), and were inhibited by heat denaturation of the membranes, omission of Ca2 , or inclusion of an inhibitor mixture comprising Ca2+ chelators (EDTA, EGTA) and calmidazolium, an inhibitor of calmodulin function (bars C and F-I). When the Ca2+ buffer (0.2 mM EGTA and 250 ,uM added Ca2+), which generates 40 ,uM free Ca2+ (19, 27), was used, the release of microvesicular lipid phosphate was comparable to that obtained when 30 ,uM or 1 mM Ca2+ was added to the reaction mixture without a Ca2+ buffer. In

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Treatment FIG. 4. Effects of Ca2' on the formation of nonsedimentable lipid-protein microvesicles from smooth microsomal membranes of 2-day-old cotyledons. Microvesicle formation is expressed in terms of lipid phosphate equivalents. Bars: A, 1-hr treatment with 1 mM Ca2+; B, 2-hr treatment with 1 mM Ca2+; C, 9-hr treatment with 1 mM Ca2+; D, 9-hr treatment with 1 mM Ca2+ and 25 jtg of streptomycin per ml; E, 9-hr treatment with 30 AM Ca2+; F, 9-hr incubation in the absence of added Ca2+; G, 9-hr incubation in the presence of 0.2 mM EGTA, 2 mM EDTA, and 25 AtM calmidazolium; H, 9-hr incubation of heat-denatured membranes in the absence of added Ca2+; I, 9-hr incubation of heat-denatured membranes in the presence of 1 mM Ca2+. Bars D, E, H, and I are single experiments. For A-C, F, and G, SEM is shown for n = 3.

addition, the lipid composition ofthe microvesicles formed in vitro in the presence of Ca2+ did not change with time up to 9 hr (Fig. 2B, lanes 3-5). Microvesicles with a similar lipid composition were also formed in vitro in the presence of Ca2+ from smooth microsomal membranes isolated from 4- and 7-day-old cotyledons (data not shown). The polypeptide pattern of membranes that had been treated for 9 hr with 1 mM Ca2+ was similar to that for control membranes (Fig. 3B, lanes 1 and 2). In the microvesicle gels, there was some smearing indicative of protein degradation, and there was also an accumulation of smaller polypeptides at the bottom of the gels that was not evident in the gels for membranes from which the microvesicles had been formed (Fig. 3B). The larger proteins present in the membrane gels were also less evident in the microvesicle gels (Fig. 3B). Dynamic Light Scattering and Electron Microscopy. Dynamic light scattering data for microvesicles isolated directly from tissue and for those formed in vitro in the presence of Ca2+ were analyzed as described (26) to obtain histograms of size distribution. Microvesicles isolated from 7-day-old cotyledons ranged from 70 to 170 nm in radius, with a mean radius of 132 nm, and those formed in vitro from smooth microsomal membranes of 2-day-old cotyledons ranged from 40 to 75 nm in radius with a mean radius of 52 nm (Fig. 5). The spherical nature of the microvesicles was evident by negative staining (Fig. 6 A and B) and by freeze-fracture electron microscopy (Fig. 6C). In addition, freeze fracturing confirmed that the microvesicles are bilayered since both convex and concave fractured faces were observed. The bilayer was particularly apparent in replicas of convex surfaces of the fractured microvesicles where the exposed face of the inner leaflet and the cross-sectioned profile of the outer leaflet were evident (Fig. 6C, arrow). Low relief particulate fine structure was distinguishable in concave exposed surfaces (corresponding to the outer leaflet of the bilayer) and convex exposed surfaces (corresponding to the inner leaflet of the bilayer) (Fig. 6C). These putative intramembranous particles are not well resolved, however, and this may be related to the finding by SDS/PAGE that the microvesicles contain partially degraded proteins and are largely devoid of

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the larger proteins found in membranes. In addition, the microvesicles were difficult to fracture, perhaps because of the high neutral lipid content, and well-resolved replicas were not routinely obtained.

DISCUSSION Nonsedimentable lipid-protein microvesicles enriched in phospholipid degradation products have been isolated from senescing as well as young bean cotyledon tissue and can also be formed in vitro from smooth microsomal membranes isolated from cotyledons. The nonsedimentable lipid and protein do not pass through a 300-kDa cut-off filter, which is consistent with their assembly in microvesicles, and they also coelute in the void volume during fractionation on Superose12. The microvesicles have a mean radius of 132 nm for those isolated directly from tissue and 52 nm for those formed in

FIG. 6. Electron microscopy of nonsedimentable lipid-protein microvesicles. (A) Negative stain of microvesicles isolated from 7-day-old cotyledons. (B) Negative stain of microvesicles formed in vitro in the presence of 1 mM Ca2' from smooth microsomal membranes of 2-day-old cotyledons. (C) Freeze-fracture replica for microvesicles formed in vitro in the presence of 1 mM Ca2+ from smooth microsomal membranes of 2-day-old cotyledons. Thick arrow, direction of shadowing; thin arrow, outer leaflet of the bilayer. (Freeze-fracture replicas of lesser quality but showing the same information were obtained for microvesicles isolated from 7-day-old cotyledons.) (A and B, x232,200; C, x162,000.)

Botany: Yao et A vitro from isolated membranes, and their bilayer nature is evident in freeze-fracture electron micrographs. The nonsedimentable nature of the microvesicles can presumably be attributed to the fact that, in comparison with membranes, they are enriched in neutral lipids (specifically, free fatty acids, long-chain aldehydes, and long-chain hydrocarbons). This is evident from the fact that these neutral lipids were not detectable by thin-layer chromatography in the lipid extract from 1.6-mg protein equivalents of membrane but were clearly evident in the lipid extracts of only 0.3-mg protein equivalents of microvesicles. Several lines of evidence suggest that these nonsedimentable microvesicles are formed in response to enzymatically mediated degradation of phospholipids within membranes. First, their formation in vitro is time dependent and inhibited by prior heat denaturation of the membranes. Second, they are formed in the presence of Ca2", which is known to stimulate the activities of membrane-associated, phospholipid-degrading enzymes leading to the formation of free fatty acids. Three lipid-degrading enzymes (phospholipase D, phosphatidic acid phosphatase, and lipolytic acyl hydrolase) are known to be associated with microsomal membranes from bean cotyledons and to be enriched in the sedimentable material obtained after partial solubilization of these membranes with Triton X-100 (19). Their collective activities generate free fatty acids, which are a dominant lipid component of the microvesicles, and two of the enzymes, phospholipase D and phosphatidic acid phosphatase, are stimulated by concentrations of Ca2" (30 uM to 1 mM) (19) that have been shown in the present study to promote microvesiculation. Finally, the microvesicles are also enriched in long-chain aldehydes and long-chain alkanes, and plant microsomes are known to possess enzymes that are capable of converting free fatty acids through (n-i) aldehydes to (n-2) alkanes (28). Thus, the membranes from which the nonsedimentable microvesicles are formed in vitro, and presumably in vivo as well, possess all of the enzymes required to form those neutral lipid catabolites of phospholipids that are the dominant lipid components of the microvesicles. It is noteworthy that these phospholipid degradation products apparently do not accumulate in the membranes since in the experiments in vitro they were not detectable in the membranes in which they were formed but were clearly evident in the resulting microvesicles. This suggests that blebbing of these microvesicles may be a means of removing phospholipid degradation products from membrane bilayers. This has been observed previously for erythrocytes under conditions in which blebbing attributable to the formation of diacylglycerol was induced by treatment with Ca2+ ionophore or exogenous phospholipase C (20, 21); however, the resulting microvesicles were sedimentable and enriched in diacylglycerol (20, 21). The nonsedimentable microvesicles isolated in the present study also appeared to contain partially degraded protein, but products of protein degradation are less likely to prompt the physical process of microvesiculation than are lipid degradation products, particularly those that have a propensity to form non-bilayer lipid configurations. It is conceivable that these nonsedimentable microvesicles are an intermediate stage of membrane deterioration in that they serve as a vehicle for moving degraded molecular components of membranes, in particular products of phospholipid degradation, into the cytoplasm for further catabolism by cytosolic or digestive body enzymes. The term "deteriosome" connotes this putative function and would


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serve to distinguish these microvesicles from other cytoplasmic microvesicles that are involved in membrane translocation unrelated to deterioration. There is clearly an abundance of these deteriosome-type microvesicles in young as well as senescing cotyledon tissue, and although their origin remains unclear, it is possible that they are a feature of normal membrane lipid turnover as well as net catabolism of membrane lipids. The authors are indebted to Dr. T. J. Beveridge for helpful discussions and to C. A. Flemming for assistance with the electron microscopy. The electron microscopy was carried out in the Natural Sciences and Engineering Research Council of Canada (NSERC), Guelph Regional Scanning Transmission Electron Microscopy Facility of the Department of Microbiology, University of Guelph, which receives NSERC infrastructure funding. This research was funded by a grant-in-aid from the NSERC. 1. Fobel, M., Lynch, D. V. & Thompson, J. E. (1987) Plant Physiol. 85, 204-211. 2. Gordon-Kamm, W. J. & Steponkus, P. L. (1984) Proc. Natl. Acad. Sci. USA 81, 6373-6377. 3. Pearce, R. S. (1985) Planta 166, 1-14. 4. Platt-Aloia, K. A. & Thomson, W. W. (1985) Planta 163, 360-369. 5. Pauls, K. P. & Thompson, J. E. (1984) Plant Physiol. 75, 1152-1157. 6. Barber, R. F. & Thompson, J. E. (1983) J. Exp. Bot. 34, 268-276. 7. Thompson, J. E., Legge, R. L. & Barber, R. F. (1987) New Phytol. 105, 317-344. 8. Mudd, J. B. (1980) in Biochemistry ofPlants: A Comprehensive Treatise, ed. Stumpf, P. K. (Academic, New York), Vol. 4, pp. 250-280. 9. Roughan, P. G. & Slack, C. R. (1976) Biochim. Biophys. Acta 431, 86-95. 10. Yoshida, S. (1978) in Plant Cold Hardiness and Freezing Stress, eds. Li, P. H. & Sakai, A. (Academic, New York), pp. 117-135. 11. Yoshida, S. (1979) Plant Physiol. 64, 241-246. 12. Diesperger, H., Muller, C. R. & Sanderman, H., Jr. (1974) FEBS Lett. 43, 155-158. 13. Yoshida, S. (1979) Plant Physiol. 64, 247-251. 14. Moore, T., Lord, J., Kagawa, T. & Beevers, H. (1973) Plant Physiol. 52, 50-53. 15. Herman, E. M. & Chrispeels, M. J. (1980) Plant Physiol. 66, 1001-1007. 16. Ichihara, K., Norikura, S. & Fujii, S. (1989) Plant Physiol. 90, 413-419. 17. Norman, H. A. & Thompson, G. A. (1986) Biochim. Biophys. Acta 875, 262-269. 18. Paliyath, G., Lynch, D. V. & Thompson, J. E. (1987) Physiol. Plant. 71, 503-511. 19. Paliyath, G. & Thompson, J. E. (1987) Plant Physiol. 83, 63-68. 20. Allan, D., Low, M. G., Finean, J. B. & Michell, R. H. (1975) Biochim. Biophys. Acta 413, 309-316. 21. Allan, D., Billah, M. M., Finean, J. B. & Michell, R. H. (1976) Nature (London) 261, 58-60. 22. Bligh, E. G. & Dyer, W. J. (1959) Can. J. Biochem. Physiol. 37, 911-917. 23. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254. 24. Laemmli, U. K. (1970) Nature (London) 227, 680-685. 25. Wray, V. P., Wray, W. W., Bovikas, T. & Hancock, R. (1981) Anal. Biochem. 118, 197-203. 26. Hallett, F. R., Craig, T., Marsh, J. & Nickel, B. (1989) Can. J. Spectrosc. 34, 63-70. 27. Blinks, J. R., Wier, W. G., Hess, P. & Prendergast, F. G. (1982) Prog. Biophys. Mol. Biol. 40, 1-114. 28. Bognar, A. L., Paliyath, G., Rogers, L. & Kolattukudy, P. E. (1984) Arch. Biochem. Biophys. 235, 8-17.