Journal of Experimental Botany, Vol. 52, No. 357, pp. 669±679, April 2001
Characterization of plasma membrane domains
enriched in lipid metabolites
Ewa Madey, Linda M. Nowack, Liming Su, Yuwen Hong1, Katalin A. Hudak2 and
John E. Thompson3
Department of Biology, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1
Received 27 July 2000; Accepted 25 September 2000
Abstract
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
Hq-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.
1
Key words: Immunoprecipitation,
membrane domains.
Hq-ATPase,
plasma
Introduction
The lipid and protein constituents of membranes have a
®nite 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-esteri®cation 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).
Present address: Skye Pharmatech Inc., 6354 Viscount Rd., Mississauga, Ontario, Canada L4V 1H3.
Present address: Biotech Center, Cook College, Rutgers University, New Brunswick, New Jersey 08901-8520, USA
3
To whom correspondence should be addressed. Fax: q1 519 746 2543. E-mail: [email protected]
Abbreviations: HEPES, (N-w2-hydroxyethylxpiperazine-N9-w2-ethanesulphonic acidx); EPPS (N-2-hydroxyethylpiperazine-N9-3-propanesulphonic acid;
DTT (dithiothreitol), EDTA (ethylenediaminetetra-acetic acid); EGTA, ethylene glycol-bis(b-aminoethyl ether)-N,N,N,N9-tetra-acetic acid; PVPP
(polyvinylpoly-pyrrolidone); PMSF (phenylmethylsulphonyl fluoride)..
2
ß Society for Experimental Biology 2001
670
Madey et al.
In the present study, vesicles of plasma membrane
origin enriched in lipid metabolites, including free fatty
acids and steryl and wax esters, have been puri®ed 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 Hq-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 HqATPase in turn mediates other secondary transport
systems associated with the cell membrane. Accordingly,
the plasma membrane Hq-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 acidi®cation 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 Hq-ATPase
is encoded by a multigene family containing up to 10
highly conserved isoforms. Several of these have been
identi®ed and characterized in a number of plant species
including tomato (Lycopersicon esculentum, seven isoforms) (Ewing et al., 1990; Ewing and Bennett, 1994),
Arabidopsis thaliana (®ve 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 HqATPase isoforms appears to be cell- and tissue-speci®c,
possibly re¯ecting functional diversity. For example, in
Arabidopsis, the Hq-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 ¯ow of
nutrients to the developing embryo (Harper et al., 1994).
In the present study, polyclonal antibodies raised
against a recombinant fragment of the AHA1 HqATPase isoform were used to immunopurify plasma
membrane vesicles from petals and leaves, and the
properties of these immunopuri®ed vesicles have been
compared with those of corresponding plasma membrane preparations obtained by phase partitioning. The
observations indicate that the immunopuri®ed vesicles
represent lipid metabolite-enriched domains within the
plasma membrane that have been released into the
cytosol, presumably by microvesiculation.
Materials and methods
Plant material
Microsomal and plasma membranes were isolated from petals of
carnation ¯owers (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 mmol m 2 s 1). Day and night temperatures were
25 8C and 18 8C, respectively. The ¯owers were harvested when
the petals were fully expanded with yellow-tinted centres.
Canola seedlings were grown in chambers at 23 8C, 50%
humidity, under 16u8 h dayunight photoperiods. Fully expanded
primary leaves of canola were harvested 16 d after planting.
Microsomal and plasma membrane isolation
Plasma membrane was puri®ed 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 ®ltered
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 ®nal concentration of 6.2% (wuw), 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 puri®ed 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 puri®ed 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
MorreÂ, 1992).
Antibodies
Antibody for immunoprecipitation and Western blotting
(designated antibody A) was raised against a conserved region
(aa 340±650) of the plasma membrane Hq-ATPase AHA1 gene
from Arabidopsis thaliana (Harper et al., 1989). The gene
sequence was ampli®ed 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 Hq-ATPase recombinant fragment
Plasma membrane lipid domains
linked though a Factor Xa proteolytic cleavage site to maltosebinding protein, was puri®ed 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 puri®ed by amylose
column chromatography and used as antigen for the generation
of polyclonal antibodies in rabbit.
A second Hq-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).
671
according to protocols already described (Hodges and Leonard,
1974).
Protein was quanti®ed 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
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 immunopuri®cation 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 ®nal
concentration of 2 mg protein ml 1 and mixed with 200 ml of
Hq-ATPase antibody serum. The mixture was incubated for
30 min at 4 8C 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 ml of microsomal
resuspension buffer in preparation for immunoprecipitation
with the magnetic beads. Prior to incubation with the antigen±
antibody complex, magnetic beads were washed ®ve 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 ml) was then
mixed with the membrane±antibody complex and incubated
for 2 h at 4 8C with continuous mixing. The membranebead±antibody complex was then collected using the DYNAL
magnetic concentrator and washed ®ve times, 4 min each, in
microsomal resuspension buffer containing 0.25 M NaCl.
Following the ®nal wash, the plasma membrane vesicles were
eluted from the bead±antibody complex by incubation
with 200 ml of 100 mM glycine (pH 2.5) for 30 min at 4 8C.
Control experiments were conducted in which microsomal
membrane suspension was immunoprecipitated with magnetic
polystyrene beads alone without the addition of the Hq-ATPase
antibody.
Enzyme assays and protein analysis
Hq-ATPase activity was determined indirectly by measuring
proton pumping using the DpH probe, acridine orange, as
described (de Michelis et al., 1983). NADPH-cytochrome
c reductase and cytochrome c oxidase activities were measured
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 identi®ed 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
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
Hq-ATPase, a marker enzyme for plasma membrane
(Larsson et al., 1994), and rotenone-insensitive NADHcytochrome c reductase and cytochrome c oxidase, marker
enzymes for endoplasmic reticulum and mitochondrial
membranes, respectively (Hodges and Leonard, 1974).
Hq-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 puri®ed by
phase partitioning from both leaves and petals was
enriched in vanadate-sensitive Hq-ATPase activity by
;4-fold by comparison with corresponding microsomal
membranes (Table 1). Plasma membrane and vacuolar
Hq-ATPases are inhibited by vanadate and nitrate,
respectively (Larsson et al., 1994; de Michelis et al.,
1983). In the present study, vanadate-sensitive (200 mM
vanadate) proton transport constituted the major portion
(;75%) of the total measurable Hq-ATPase activity in
plasma membrane vesicles for leaves and petals, whereas
Table 1. Enzyme activities of microsomal membrane and phase-puri®ed plasma membrane fractions from carnation petals and canola
leaves
Values are means"SE for n ˆ 3.
Species
Fraction
Vanadate-sensitive Hq-ATPase
(mOD min 1 mg 1 protein)
NADPH-cytochrome c reductase
(nmol min 1 mg 1 protein)
Cytochrome c oxidase
(nmol min 1 mg 1 protein)
Canola
Microsomes
Plasma membrane
Microsomes
Plasma membrane
20.8"0.83
95.0"4.3
43.3"12.5
186.7"34.0
9.0"0.8
2.3"0.3
8.3"0.4
2.8"0.3
70.2"19.3
16.5"3.0
12.7"2.2
3.6"0.8
Carnation
672
Madey et al.
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-puri®ed membrane preparation is attributable to vesicles of plasma
membrane.
Marker enzymes were used to identify other
contaminating membranes in the phase-puri®ed plasma
membrane preparations. The speci®c 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 puri®ed plasma membrane fraction
than in corresponding microsomal membranes for leaves
and petals (Table 1). These observations together with the
enrichment of vanadate-sensitive Hq-ATPase activity
can be interpreted as indicating that the phase-puri®ed
membrane preparations from leaves and petals are composed of predominantly plasma membrane vesicles.
Plasma membrane vesicles were also puri®ed from
microsomal membrane preparations by immunoprecipitation with antibodies raised against the plasma membrane Hq-ATPase. Two antibodies, one designated
antibody A, which was raised against a recombinant
fragment of the Hq-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 Hq-ATPase polypeptide
(;100 kDa) in Western blots. Antibody A did not
recognize the native Hq-ATPase polypeptide in Western
blots of either microsomal membranes or phase-puri®ed
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 Hq-ATPase including polypeptides at 41, 43 and 68 kDa (Fig. 3). The 41 and
43 kDa polypeptides were clearly resolved in some blots
Protein composition of membrane fractions
The protein compositions of phase-puri®ed plasma membrane and corresponding microsomal membrane preparations were examined by SDS-PAGE. For both leaves
and petals, the polypeptide composition of phase-puri®ed
plasma membrane preparations was clearly distinguishable from that of microsomal membranes (Fig. 1A, B).
This ®nding is consistent with enzyme data (Table 1)
indicating that the phase-puri®ed membrane vesicle
preparations are enriched in plasma membrane vesicles
relative to microsomal membranes.
Fig. 1. SDS-PAGE (12%) of microsomal membranes and phasepuri®ed plasma membrane fractions from carnation petals and canola
leaves. (A) Carnation petals: lane 1, microsomal membranes; lane 2,
phase-puri®ed plasma membrane. (B) Canola leaves: lane 1, microsomal
membranes; lane 2, phase-puri®ed plasma membrane. Each lane was
loaded with 1 mg of protein, and the gels were stained with silver.
Molecular mass markers (kDa) are indicated.
Fig. 2. Diagrammatic representation of the Hq-ATPase polypeptide
denoting the localization and length of the recombinant fragments
(indicated in grey) used to generate polyclonal antibodies. Antibody A
was raised against the hydrophilic mid-section of the protein corresponding to aa 340±650; antibody B was raised against aa 390±662.
Plasma membrane lipid domains
(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 Hq-ATPase protein corresponding to
aa 340 through 650 and does not recognize the native
Hq-ATPase polypeptide, was used for immunoprecipitation in order selectively to purify vesicles of plasma
membrane that contain catabolites of the Hq-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 immunopuri®ed plasma membrane
vesicles proved to be quite distinct from that of corresponding phase-puri®ed 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 steryluwax esters) relative to
phospholipid are enriched in the immunopuri®ed plasma
membrane vesicles by comparison with phase-puri®ed
plasma membrane (Table 2). Indeed, the ratio of free to
esteri®ed fatty acids is ;5-fold higher in immunopuri®ed
vesicles than in corresponding phase-puri®ed plasma
membrane (Table 3). This supports the contention that
the two puri®cation procedures yield different populations of plasma membrane vesicles. It is also clear that the
Fig. 3. Western blots of microsomal and plasma membrane fractions
from carnation petals probed with polyclonal antibodies raised against
recombinant fragments of the Hq-ATPase. Antibody A, raised against
aa 340±650; antibody B, raised against aa 390±662. Lanes 1 and 3,
microsomal membranes; lanes 2 and 4, phase-puri®ed plasma membrane. Each lane was loaded with 5 mg of protein. Molecular masses
(kDa) are indicated.
673
lipid compositions of the immunopuri®ed 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 nonspeci®cally to magnetic beads incubated with microsomal
membranes in the absence of Hq-ATPase antibody was
below the limit of detection.
The fatty acid composition of phase-puri®ed and
immunopuri®ed 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 immunopuri®ed membrane
vesicles from canola leaves and carnation petals are
11-fold and 23-fold higher, respectively, than the corresponding ratios for phase-puri®ed plasma membrane,
indicating that the immunopuri®ed 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-puri®ed plasma membrane all contain high levels of the diunsaturated
fatty acid, linoleic acid (18:2), and the steryluwax ester
fraction contains both linoleic acid and the triunsaturated
fatty acid, linolenic acid (18:3) (Fig. 5). By contrast,
corresponding lipid classes from the immunopuri®ed
Fig. 4. Lipid class composition of microsomal membrane (black solid
bars), phase-puri®ed plasma membrane (hatched bars), immunopuri®ed
plasma membrane (white bars) and rabbit Hq-ATPase antiserum (grey
bars) included for comparison. (A) Membrane fractions from carnation
petals. (B) Membrane fractions from canola leaves. The data are
expressed in terms of fatty acid equivalents as a percentage of total fatty
acid. (PL), phospholipid; (DG), diacylglycerol; (FFA), free fatty acids;
(TAG), triacylglycerol; (SWE), steryl and wax esters. Where indicated
values are means"SE for n ˆ 3.
674
Madey et al.
Table 2. Levels of lipid metabolites in microsomal membrane, phase-puri®ed plasma membrane and immunopuri®ed plasma membrane
vesicles relative to phospholipid
(A) Membrane fractions from carnation petals. (B) Membrane fractions from canola leaves. Fatty acid equivalents for each of diacylglycerol (DG),
free fatty acids (FFA), triacylglycerol (TAG), and steryluwax esters (SWE) were expressed as a proportion of phospholipid fatty acid equivalents for
each membrane fraction.
(A)
Lipid class
Proportion relative to phospholipid
DG
FFA
TAG
SWE
Phase-puri®ed
Immunopuri®ed
Enrichment (immunopuri®eduphase puri®ed)
0.33
0.06
0.03
0.02
1.07
0.83
0.86
0.95
3.23
14.34
33.33
40.59
(B)
Lipid class
Proportion relative to phospholipid
DG
FFA
TAG
SWE
Microsomal
membrane
Phase-puri®ed
Immunopuri®ed
Enrichment
(immunopuri®edu
microsomes)
Enrichment
(immunopuri®edu
phase puri®ed)
0.06
0.11
0.03
0.02
0.10
0.03
0.03
0.03
0.88
0.61
0.64
0.87
14.67
5.55
21.33
43.5
8.81
18.19
19.57
27.60
Table 3. Levels of free and saturated fatty acids in membrane
vesicles from carnation petals and canola leaves
Free fatty acids are expressed as a ratio of esteri®ed fatty acids, and
saturated fatty acids are expressed as a ratio of unsaturated fatty acids.
Species
Fraction
Saturated to
unsaturated
fatty acid
ratio
Free to
esteri®ed
fatty acid
ratio
Canola
Microsomal membrane
Phase-puri®ed
Immunopuri®ed
0.43
0.72
8.12
0.10
0.03
0.18
Carnation
Phase-puri®ed
Immunopuri®ed
0.24
5.47
0.04
0.21
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-puri®ed plasma
membrane were again more unsaturated than the corresponding fractions from immunopuri®ed vesicles (Fig. 6).
However, in canola the fatty acid compositions of the free
fatty acid, triacylglycerol and steryluwax ester fractions
were very similar (Fig. 6).
Discussion
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 MorreÂ, 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-puri®ed membrane fraction was assessed by measuring the levels of vanadatesensitive 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 DpH
in membrane vesicles. Since Hq translocation and ATP
hydrolysis are stoichiometric, the acridine orange assay
serves as a reliable measure of Hq-ATPase activity
(Briskin, 1990). Based on this assay, the vanadatesensitive Hq-ATPase speci®c activity of phase-puri®ed
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 Hq-ATPase activity can be interpreted as
re¯ecting puri®cation of plasma membrane.
Plasma membrane lipid domains
675
Fig. 5. Fatty acid composition of the separated lipid classes for phase-puri®ed plasma membrane (hatched bars) and immunopuri®ed plasma
membrane (solid black bars) from carnation petals. (A) Phospholipid; (B) diacylglycerol; (C) free fatty acid; (D) triacylglycerol; (E) steryl and wax
esters. 16 : 0, Palmitic acid; 16 : 1, palmitoleic acid; 18 : 0, stearic acid; 18 : 1, oleic acid; 18 : 2, linoleic acid; 18 : 3, linolenic acid. Values are means"SE
for n ˆ 3.
Fig. 6. Fatty acid composition of the separated lipid classes for microsomal membrane (black solid bars), phase-puri®ed plasma membrane
(white bars) and immunopuri®ed plasma membrane (grey bars) from canola leaves. (A) Phospholipid; (B) diacylglycerol; (C) free fatty acid;
(D) triacylglycerol; (E) steryl and wax esters. 16 : 0, Palmitic acid; 16 : 1, palmitoleic acid; 18 : 0, stearic acid; 18 : 1, oleic acid; 18 : 2, linoleic acid; 18 : 3,
linolenic acid. Where indicated values are means"SE for n ˆ 3.
676
Madey et al.
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 speci®cally 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-puri®ed membrane preparations. Vanadatesensitive proton transport constituted ;75% of the total
ATP-dependent proton transport in the puri®ed 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 speci®c activities of
both markers were about 4-fold lower in the puri®ed
plasma membrane fraction than in corresponding microsomal membranes. These ®ndings indicate a relatively
low abundance of membrane vesicles originating from
mitochondria and endoplasmic reticulum in the puri®ed
plasma membrane fraction.
Plasma membrane vesicles were also isolated from preparations of microsomal membrane vesicles by immunopuri®cation using a polyclonal antibody (Antibody A)
raised against a recombinant fragment of the HqATPase. Two different antibodies (A and B) were tested
by Western blotting for their ability to cross-react with
the Hq-ATPase polypeptide. Antibody B, which was
raised against a central region of the Hq-ATPase
corresponding to aa 390±662, recognized the native HqATPase 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 Hq-ATPase
corresponding to aa 340±650, recognized three of the
same lower molecular weight catabolites of the HqATPase (68, 43 and 41 kDa), but did not recognize the
native polypeptide. Thus the catabolites of the HqATPase apparently possess epitopes that are not exposed
on the native protein. The ®nding that antibody A did
not recognize the native protein may also re¯ect the fact
that it was raised against a denatured antigen. The
immunopuri®ed plasma membrane vesicles are presumably inside-out since the antibody was raised against the
hydrophilic domain of the Hq-ATPase projecting into
the cytoplasm in intact cells.
The ®nding that antibody A recognizes the same
catabolites as antibody B and no other polypeptides
indicates that it is speci®c for the Hq-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 membraneassociated catabolites of the native Hq-ATPase recognized by antibody A in both puri®ed plasma membrane
and microsomes are polypeptides of cytosolic origin
unrelated to the Hq-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 ®nding in the present study
that Hq-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 Hq-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-puri®ed
plasma membrane and immunopuri®ed 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-puri®ed plasma membrane consisted primarily of
phospholipid. The possibility that the enrichment of lipid
metabolites relative to phospholipid in the immunopuri®ed plasma membrane vesicles simply re¯ects 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 immunopuri®ed fraction are clearly distinguishable
from those in the corresponding phase puri®ed plasma
membrane. Speci®cally, the diacylglycerol, free fatty
acids, steryl and wax esters and triacylglycerol of the
immunopuri®ed vesicles contain much higher proportions of saturated fatty acids than were found in the
corresponding lipid classes of phase-puri®ed plasma
membrane. Indeed, the saturated : unsaturated fatty acid
ratios for immunopuri®ed 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-puri®ed plasma membrane. If the enrichment of lipid metabolites relative
to phospholipid in the immunopuri®ed vesicles were
simply due to phospholipid leakage, there should not
be a difference in the fatty acid pro®les of the lipid
metabolites.
Plasma membrane lipid domains
The presence of triacylglycerol in both phase-puri®ed
and immunopuri®ed 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 re¯ect inclusion of triacylglycerol in the
bilayers of microvesicles originating from the endoplasmic reticulum that serve to target proteins to the
plasmalemma.
The ®nding that the immunopuri®ed 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 immunopuri®ed plasma membrane vesicles, speci®cally free fatty acids and steryluwax
esters, are gel-phase forming lipids (Yao et al., 1991), and
their presence in membranes would thus lead to lipid
phase separations. The immunopuri®ed vesicles from
carnation petals, for example, proved to be enriched
in free fatty acids and steryluwax esters by 14-fold and
40-fold, respectively, by comparison with corresponding
phase-puri®ed plasma membrane.
One likely interpretation of these ®ndings is that
microsomal fractions, while being comprised in the main
of vesicles formed during tissue homogenization, also
contain vesicles originating from lipid metaboliteenriched 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 traf®cking 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 metaboliteenriched 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 suf®cient 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 immunopuri®ed plasma membrane vesicles is attributable to the action of a lipase
released during homogenization. First, plant and animal
677
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 immunopuri®ed
vesicles differs from that of the plasmalemma in ways
that cannot be explained simply by the action of one or
more lipases. Speci®cally, vesicles of plasma membrane
formed during tissue homogenization would have the
same lipid composition as the plasmalemma, whereas the
immunopuri®ed vesicles are enriched in free fatty acids,
in steryluwax esters and in triacylglycerol by comparison
with phase-puri®ed 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 immunopuri®ed 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 steryluwax 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 immunopuri®ed plasma membrane
vesicles from the same tissue in that they have a much
lower buoyant density. Speci®cally, the lipid particles
were isolated by ¯otation centrifugation of a microsomal
supernatant (cytosol fraction) that was made 10% (wuv)
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 immunopuri®ed 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 re¯ects
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
678
Madey et al.
others by blebbing of lipid particles from the membrane
surface.
Acknowledgements
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 Hq-ATPase antibody.
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