From the Department of Molecular and Cellular Biochemistry and the Lucille P. Markey Cancer Center, University of Kentucky College of Medicine, Lexington, Kentucky 40536
Received for publication, January 28, 2003 , and in revised form, April 25, 2003.
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ABSTRACT |
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INTRODUCTION |
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S. cerevisiae V-ATPase contains two components or domains, V1 and V0, which associate to form an active V1V0 complex on the vacuolar membrane (reviewed in Refs. 35). The V1 domain has ATPase activity and is composed of 8 different proteins. It can exist free in the cytoplasm or complexed with V0 on the vacuolar membrane. The V0 domain contains 5 protein subunits and is imbedded in the vacuolar membrane where it serves as a proton pore. The V1 and V0 domains are assembled and associate in the ER to form functional V1V0 complexes, which are then transported from the Golgi to the vacuole (reviewed in Refs. 3 and 5). Alternatively, the two domains assemble independently and then associate once V0 reaches the vacuole.
The ER is also where sphingolipid synthesis begins (reviewed in Ref. 6) with generation of ceramide. Ceramide is transported to the Golgi where it is converted sequentially into the complex sphingolipids inositol-phosphoceramide, mannose-inositol-phosphoceramide, and finally to mannose-(inositol-phospho)2-ceramide. Complex sphingolipids are delivered to cellular compartments, particularly the plasma membrane (7) and to a lesser extent the vacuole (8).
One of the distinguishing features of sphingolipids in S. cerevisiae is the C26 acyl group. Fatty acids with 20 or more carbons, very long chain fatty acids (VLCFAs), are ubiquitous in nature, but little is known about their functions. VLCFAs are mostly found in the ceramide portion of sphingolipids. The importance of the C26 acyl component of S. cerevisiae sphingolipids was demonstrated by the isolation of mutant strains that do not make sphingolipids (9), but instead make a set of novel sphingolipid mimics in which ceramide is replaced by diacyl-glycerol (10). The presence of a C26 acyl group is the unique feature of these novel glycerolipids, which enables them to mimic some sphingolipid functions.
Further evidence for the essentiality of the C26 acyl group in
sphingolipids comes from studies of the FEN1 (ELO2) and
SUR4 (ELO3) genes. In fen1 cells only 29% of
the sphingolipids contain a C26 acyl group, the rest contain C22 and C24 acyl
groups. The sphingolipids in sur4
cells contain only C22 and
C24 acyl groups and no C26s
(11,
12). Fen1p and Sur4p are
components of the enzyme system that elongates C16 and C18 fatty acids to form
VLCFAs. The exact function of Fen1p and Sur4p are unclear because the
elongation system has not been fully characterized in any organism.
sur4
and fen1
cells are viable, although they
have many mutant phenotypes (reviewed in Ref.
13) and deletion of both genes
is lethal (14).
A fraction of sphingolipids in higher eukaryotes also contain VLCFAs
(15) and the mouse genes
Ssc1 and Cig30 complement a sur4 and a
fen1
mutant, respectively
(16). Interestingly, the mice
mutants Quaking and Jimpy, which develop intense tremors at the age of about 2
weeks as a result of severe demyelination of the central nervous system, have
reduced levels of Ssc1 mRNA and reduced fatty acid elongation
activity (17). Cig30
mRNA has the interesting property of being induced in brown adipose tissue
when animals are exposed to cold temperature. These results suggest that
VLCFAs are performing important, but unknown functions in mammals, and that
some of these functions may be evolutionarily conserved.
Recently Kohlwein et al.
(12) reported that
sur4 and fen1
cells contain small vacuoles
called fragmented vacuoles that fail to properly fuse to form larger vacuoles.
This observation suggested to us that sphingolipids with a C26 acyl group are
needed for some vacuolar function(s). Here we show that V1 domains
in sur4
cells lack ATPase activity even though they associate
with V0 domains on the vacuolar membrane. Our data are the first to
implicate sphingolipids with a C26 acyl group in the generation of a fully
functional V1 domain.
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EXPERIMENTAL PROCEDURES |
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Buffered medium was prepared by the addition of 50 mM MES and 50 mM MOPS to YPD, and the pH was adjusted to pH 5.5 with NaOH. YPD plates supplemented with 100 mM CaCl2,4mM CaCl2, or buffered to pH 7.5 with 100 mM Hepes were prepared as described previously (21). YPD medium contained 1% yeast extract, 2% Bacto-peptone, and 2% dextrose.
Quinacrine Staining and Semiquantitative Quinacrine Assay Vacuolar accumulation of quinacrine was assessed by fluorescence microscopy as described by Roberts et al. (22) or by a semiquantitative quinacrine assay (21). Fluorescence measurements of cell suspensions were done in a Beckman spectrofluorometer (excitation = 419 nm, emission = 425 nm) and the OD at 600 nm was monitored in a spectrophotometer.
SDS-PAGE, Immunoblotting, and AntibodiesSDS-PAGE and immunoblotting were performed according to procedures recommended for the Bio-Rad Tray-Blot S.D. Semi-Dry transfer cell (Bio-Rad Inc.). Monoclonal antibodies against Vph1p, Vma1p, Vma2p, CPY, and ALP were from Molecular Probes. Anit-Myc antibodies were from Roach Applied Science. Dr. Patricia Kane provided anti-Vma5p. Nitrocellulose membranes (Bio-Rad) were washed four times after the first and second antibody reactions with 0.1% phosphate-buffered saline containing 0.1% Triton X-100. Secondary antibody was anti-mouse IgG conjugated to alkaline phosphatase (Sigma). Membranes were incubated for 5 min facedown in ECF substrate (Amersham Biosciences) and fluorescent signals were collected by using a Molecular Dynamics Storm Phosphorimager and quantified by using ImageQuant software (version 5.1).
HPLC Analysis of LCBs and LCBPsLipids were extracted from whole cells, converted to fluorescent derivatives and analyzed by HPLC as described previously (23).
Treatment of Purified Vacuoles with PHSA 100x stock
of PHS (10 mM PHS in 95% EtOH) was diluted into a suspension of
purified vacuolar membranes suspended in buffer (10 mM Tris, 10
mM MES, pH 6.9, 5 mM MgCl2, 25 mM
KCl) to give a final concentration of 100 µM. Samples were
incubated on ice for 0, 30, 60, 90, and 120 min followed by centrifugation at
13,000 rpm for 10 min in a microcentrifuge (4 °C). Proteins in the
supernatant fluid were precipitated by incubating with 5% trichloroacetic acid
(final concentration) for 2 h on ice. Pellets and trichloroacetic
acid-precipitated proteins were resuspended in 50 µl of cracking buffer (8
M urea, 5% SDS, 1 mM ethylenediamine tetraacetate, 50
mM Tris-HCl, pH 6.8, 5% -mercaptoethanol, Ref.
24) and equal volumes were
analyzed by SDS-PAGE and immunoblotting.
Miscellaneous ProceduresVacuolar membrane vesicles were purified by centrifugation on Ficoll gradients and Mg-ATPase activity was measured at 23 °C as described previously (22) except that membranes were not homogenized before centrifugation on the second Ficoll gradient.
Vacuolar membrane vesicles were also purified by sucrose density gradient centrifugation as previously described (25), except that the concentration of sucrose was increased to prevent membranes from pelleting on the bottom of the centrifuge tube. For these experiments, 4 ml of the membrane fraction was overlaid onto a 32-ml gradient composed of equal volumes of 10, 30, 50, and 60% (w/v) sucrose. The gradient was centrifuged for 35 min at 100,000 x g in a Sorvall AH629 rotor at 4 °C and then fractionated starting from the top into 9 fractions of 4, 7, 2, 6, 2, 6, 2, 7 ml and the resuspended pellet. Fractions were frozen in liquid nitrogen and stored at 80 °C. V1V0 complexes on vacuolar membranes prepared by sucrose gradient centrifugation were dissociated by treatment with KI as described previously (26, 27).
Cell-free protein extracts used for analysis of Vph1p (Fig. 3), were prepared as described by Kunz et al. (28). Cell-free protein extracts used for the analysis Vma1p, Vma2p, and Vma5p were prepared as described (29).
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To measure the calcium-dependent ATPase activity of cytosolic V1 domains, cells were grown, lysed and a high speed supernatant fraction was prepared as previously described (5). Proteins were precipitated by treatment with 5% trichloroacetic acid as described above, resuspended in and dialyzed against buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 0.5 mM p-methylsulfonylfluoride, 10% glycerol), and centrifuged on a glycerol gradient (30). Fractions (750 µl) were collected from the top of the gradient. Fractions (912) containing V1 domains were located by immunoblotting for Vma1p, Vma2p, Vma5p, and Vph1p. The pooled V1-containing fractions were assayed for calcium-dependent ATPase activity as described previously (5). Values are expressed as the difference between assays performed with and without 1.6 mM CaCl2.
The subunit composition (Fig.
7) of V0 and the V1V0 complex
were analyzed by using a published procedure to cross-link proteins before
immunoprecipitation and SDS-PAGE analysis
(31). The procedure was
modified so that 1 OD of 600-nm units of spheroplasts were incubated with 50
µCi of Trans[35S] label (ICN Inc., 1175 Ci/mmol, 5100607). After
pretreatment of the sample with protein A-Sepharose beads (Sigma Inc.), 400
µl a solution of 5% bovine serum albumin/phosphate-buffered saline
containing 5 µl of antibody solution was added, and the sample was
incubated overnight on ice with mixing. Protein A-Sepharose (40 µl of a 40%
(v/v) suspension) was added to each sample and incubated for2honice with
mixing. Immunoprecipitates were collected by centrifugation at 5,000 rpm for 5
min in a microcentrifuge. Pellets were washed four times in buffer (1% Triton
X-100, 1% deoxycholic acid, 50 mM Tris-HCl, pH 7.5, 150
mM NaCl), and precipitated proteins were eluted from the beads by
incubation for 10 min at 95 °C in 50 µl of 4x SDS-loading buffer
(50 mM Tris base, pH 6.8, 8% glycerol, 1.6% SDS, 4%
-mercaptoethanol, 0.04% bromphenol blue). Half of each precipitate was
subjected to SDS-PAGE and phosphorimager analysis.
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Cells were prepared for indirect immunofluorescent microscopy by using a published procedure (22). The secondary antibody was goat anti-mouse IgG labeled with FluoroLinkTM Cy3TM (Amersham Biosciences). Fluorescent images were obtained with a Nikon Àclipse E800 fluorescence microscope equipped with a Nikon 100X/1.3 plan fluor oil-immersion objective and a Diagnostic instruments Spot camera controlled by Adobe Photoshop software. For Cy3 fluorescence, samples were excited at 510560 nm and viewed with a barrier filter of 570650 nm. Adobe PhotoShop software was used to process images. Protein concentrations were determined with the Bio-Rad DC protein assay kit with bovine serum albumin as a standard.
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RESULTS |
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Another Vma phenotype is failure to grow on YPD plates
containing 4 mM ZnCl2
(34). We found that
sur4 cells have this phenotype whereas fen1
cells do not since they grow almost as well as wild-type cells
(Fig. 1). Other
Vma phenotypes include failure to grow in the presence of
100 mM CaCl2
(35) or on medium containing a
non-fermentable carbon source such as glycerol
(36). We found that 100
mM CaCl2 inhibits growth of sur4
cells
but does not inhibit growth of fen1
cells
(Fig. 1), again supporting the
idea that sur4
cells are less able to acidify their vacuoles
than fen1
cells. In addition, sur4
cells grow
slowly on YP-glycerol while fen1
cells grow slightly faster
and both grow better than vma2
control cells
(Table II). These data show
that the Vma phenotypes of sur4
cells are
nearly as pronounced as those in vma2
cells and that the
phenotypes are less severe in fen1
cells. Based upon these
phenotypes it appears that the V-ATPase is more impaired in
sur4
cells than in fen1
cells.
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To directly examine vacuolar acidification in vivo we used the
lysosomotropic fluorescent dye quinacrine, which is taken up by cells and
concentrated in acidified vacuoles; if vacuoles are not acidified the dye
remains in the cytoplasm and gives a diffuse fluorescent signal
(37). A strong fluorescent
vacuolar signal was observed in wild-type cells
(Fig. 2A). No
fluorescent signal was seen in sur4 cells, which behaved like
vma2
control cells that do not acidify their vacuole. The
fluorescent signal in fen1
cells was lower than in wild-type
cells but greater than in sur4
cells.
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The fluorescent microscopy results were verified by a semiquantitative
spectrofluorometric assay of the relative fluorescence of populations of cells
stained with quinacrine. By this technique, wild-type cells fluoresced
strongly whereas sur4 cells fluoresced near the background
level measured in vma2
cells
(Fig. 2 and
Table II). The fluorescent
signal in fen1
cells was 60% of the wild-type level indicating
that they are able to acidify their vacuoles better than sur4
cells but not as well as wild-type cells. Taken together these data show that
sur4
cells have a unique set of Vma
phenotypes and that they fail to acidify their vacuoles. These phenotypes are
less severe in fen1
cells, which have some ability to acidify
their vacuoles, although they do not acidify as well as wild-type cells.
V-ATPase Activity Is Reduced in sur4 Cells and Ficoll
Dissociates the V1V0
ComplexThe data presented in Figs.
1 and
2 and
Table II suggest that vacuolar
membranes isolated from sur4
cells should have less V-ATPase
activity than vacuolar membranes isolated from fen1
cells and
both should have less activity than those isolated from wild-type cells. In
agreement with this prediction, vacuolar membranes isolated by Ficoll density
gradient centrifugation from sur4
cells had only 10% of the
wild-type V-ATPase activity while those isolated from fen1
cells had 25% of the wild-type activity
(Table III).
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Reduced V-ATPase activity could be due to either a reduced specific
activity or fewer molecules. To differentiate between these alternatives the
concentration of Vph1p, a subunit of the V0 domain, and Vma1p,
Vma2p, and Vma5p, subunits of the V1 domain, were measured by
immunoblotting of whole cell protein extracts and of purified vacuolar
membranes (Fig. 3). Whole cell
protein extracts from wild-type, sur4, and
fen1
mutants contained a similar level of each of the four
proteins, indicating that the steady-state level of the proteins is similar in
mutant and wild-type cells. Likewise, Vph1p was present in similar amounts in
vacuolar membranes purified from the three strains. In contrast, the level of
Vma1p Vma2p, and Vma5p in vacuolar membranes isolated from
sur4
and fen1
cells was greatly reduced
(Fig. 3). These results suggest
that there is a defect in the V1 domain in sur4
and
fen1
cells or that the interaction between V0 and
V1 is abnormal and V1 or some of its subunits dissociate
during vacuolar purification.
The interaction of V1 with V0 has been examined by
treating vacuolar membranes with a low salt buffer containing ethylenediamine
tetraacetate and then determining if particular protein subunits remained in
the vacuolar fraction (high speed pellet) or became soluble
(38). Because Ficoll
purification removed V1 subunits from vacuolar membranes, we used
cell-free extracts in place of vacuolar membranes for these assays. Treatment
with a low salt buffer did not reveal any difference in the level of Vma1p and
Vma2p in the pellet and soluble fractions obtained with sur4,
fen1
, or wild-type cells (data not shown). We also determined
if increasing concentrations of sodium carbonate
(38) preferentially
solubilized Vma1p and Vma2p in cell-free extracts made from
sur4
and fen1
cells compared with wild-type
cells. Vma2p but not Vma1p was more readily solubilized in the
sur4
and fen1
samples (data not shown). Thus,
whatever the nature of the abnormality in the interaction between Vma1p and
Vma2p and V0 in sur4
and fen1
cells, it must be fairly subtle and unique since, as far as we are aware,
sensitivity of the V1V0 complex to Ficoll has not been
reported previously.
Vacuoles have also been partially purified by using sucrose density
gradient centrifugation (25).
We examined this procedure to see if V1 domains remained attached
to vacuolar membranes isolated from sur4 cells. Sucrose
gradient fractions were analyzed by immunoblotting for Vph1p to detect
V0 domains and for Vma1p and Vma2p to detect V1 domains.
The concentration of these three proteins peaked around fractions 5 and 6 in
both the sur4
and wild-type sample
(Fig. 4). Thus, unlike the
results obtained when extracts from sur4
cells were
centrifuged on Ficoll gradients, sucrose gradients yield a fraction in which
the V1 and V0 domains remain associated.
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Fractions 5 and 6 obtained from wild-type cells contain functional
V1V0 complexes because they have V-ATPase activity which
has a slightly higher specific activity than that measured in vacuolar
membranes isolated by Ficoll density gradient centrifugation
(Table III). Even though the
immunoblot of the sur4 sample
(Fig. 4) indicates that
fractions 5 and 6 have both V1 and V0 domains, the
fractions have only 20% as much V-ATPase activity as the wild type.
The data presented in this section show that V1 domains do
associate with V0 on the vacuolar membrane in sur4
cells but the association is abnormal because the Ficoll gradient procedure
causes Vma1p, Vma2p, and Vma5p and possibly the entire V1 domain to
dissociate from V0 domains. In addition, vacuolar membranes
isolated from sur4
cells by either Ficoll or sucrose density
gradients have reduced V-ATPase activity.
V1 Associates with the Vacuolar Membrane
in sur4 and fen1
CellsBoth density
gradient procedures for isolating vacuolar membranes subjects the sample to
non-physiological conditions and could create artifacts such as by
proteolysis. To try and avoid these potential complications, we examined the
association of V1 with V0 on the vacuolar membrane of
intact cells by using indirect immunofluorescence microscopy with anti-Vma1p
or anti-Vma2p antibodies (33).
In wild-type cells both antibodies localized to the vacuolar membrane as
expected (Fig. 5). A similar
localization is seen in sur4
and fen1
cells
except that the vacuoles are fragmented and do not stain as uniformly as do
vacuoles in wild-type cells (Fig.
5). Control cells lacking the vma2 gene show diffuse
staining throughout the cytoplasm with anti-Vma1p antibody (data not shown)
because V1 subunits are not formed and no staining with anti-Vma2p
is observed because the protein is absent
(Fig. 5). These data verify
those obtained by sucrose density gradient centrifugation and together the two
sets of data establish two critical points about the V-ATPase in
sur4
and fen1
cells. First, V1
domains are assembled and, second, at least some of them do associate with
V0 on the vacuolar membrane.
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V0 Domains Are Functional in
sur4 Cells but V1 Domains Lack
ATPase ActivityThe nature of the V-ATPase defect in
sur4
cells was further elucidated by determining whether
V1, V0 or both domains are defective. For these
experiments vacuolar membranes were isolated by sucrose density gradient
centrifugation then treated with KI to dissociate V1
(27). Samples were then
centrifuged to give a vacuolar membrane pellet (P) containing V0
and a supernatant fraction (S) containing V1. It has been shown
previously that upon mixing the P and S fractions and dialyzing away the KI,
V1 and V0 associate, as determined by immunoblotting,
and V-ATPase activity is partially restored (3540%)
(27). We observed very similar
results using the P and S fractions derived from wild-type cells. By
immunoblotting, Vma1p and Vma2p (V1 subunits) were more
concentrated in the P fraction following dialysis compared with the
non-dialyzed sample (Fig.
6A, sample 2) showing that V1
associates with V0. In addition, 35% of ATPase activity was
restored in the dialyzed sample compared with only 9% in the non-dialyzed
sample. The control experiment using wild-type V1V0
complexes that were not treated with KI showed that all of the Vma1p and Vma2p
were in the pellet along with Vph1p (Fig.
6A, sample 1). Dialysis reduced ATPase activity
to 68% of the non-dialyzed sample. Similar mixing experiments with the P
(V0) and the S (V1) fractions from sur4
cells showed that dialysis does promote association of Vma1p and Vma2p with
the P fraction, but it does not restore any ATPase activity
(Fig. 6A, sample
4). The control experiment using sur4
V1V0 complexes that were not treated with KI showed that
all of the Vma1p and Vma2p were in the pellet along with Vph1p and that
dialysis reduced ATPase activity slightly from 18 to 13%
(Fig. 6A, sample
3). Results from other control reactions are shown in samples 7 and 8 of
Fig. 6.
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Mixing the S (V1) fraction from wild-type cells with the P
(V0) fraction from sur4 cells followed by dialysis
resulted in association of Vma1p and Vma2p with the P fraction and restoration
of 36% of the ATPase activity (Fig.
6A, sample 5). A reciprocal mixing experiment
with the P (V0) fraction from wild-type cells the S (V1)
fraction from sur4
cells showed that Vma1p and Vma2p associate
with the P fraction but there was no restoration of ATPase activity
(Fig. 6A, sample
6). These experiments demonstrate that V0 domains in
sur4
cells are fully functional and can associate with
wild-type V1 domains to generate ATPase activity whereas the
V1 domains are capable of association with V0 on the
vacuolar membrane but they do not have ATPase activity.
V1 Domains from sur4 Cells
Lack Ca-ATPase Activity Free V1 domains do not show
ATPase activity when assayed in the presence of Mg2+,
but do show activity when assayed in the presence of
Ca2+
(20). Thus, to verify that
V1 domains in sur4
cells lack ATPase activity we
partially purified V1 complexes by velocity centrifugation on
glycerol gradients and assayed them for ATPase activity in the presence
Ca2+. To remove background ATPase activity that was not
stimulated by Ca2+, the assay was also done in the
absence of Ca2+ and this value was subtracted from the
value obtained in the presence of Ca2+ to give the
Ca-stimulated activity. In preliminary experiments increasing concentrations
of protein in the pooled V1-containing sucrose gradient fractions
were assayed for Ca-stimulated ATPase activity (release of Pi).
Activity was linear with increasing protein concentration (data not shown). A
protein concentration that gave easily measured activity using V1
domains from wild-type cells was chosen for kinetic analyzes. The kinetics of
Ca-stimulated ATP hydrolysis for wild-type V1 domains were linear
up to about 5 min and then reached a plateau whereas the V1 domains
from sur4
cells showed no ATP hydrolysis over the entire 30
min incubation period (Fig.
6B). These results verify those shown in
Fig. 6A and based upon
the combined data we conclude that V1 domains in
sur4
cells lack ATPase activity.
The Subunit Composition of the V1 and
V0 Domains Appears Normal in sur4
CellsV1 domains in sur4
cells may
have reduced ATPase activity and dissociate from the vacuolar membrane in the
presence of Ficoll because a subunit is missing. To examine the subunit
composition of V1 and V0 domains, cells were converted
to spheroplasts, metabolically labeled with [35S]amino acids,
gently lysed, and then proteins were cross-linked with a reversible
cross-linker. Samples were immunoprecipitated with monoclonal antibodies
specific for Vph1p, Vma1p, or Vma2p and radioactive proteins in the
immunoprecipitate were analyzed by SDS-PAGE and phosphorimaging.
The anti-Vph1p antibody used in these experiments only recognizes
V0 domains that are not complexed with V1
(24) and is, therefore, useful
for examining the subunit composition of V0 domains. Radiolabeled
proteins of 100 (Vph1p), 36 (Vma6p), 19 (identity unknown, Refs.
30 and
31) and 17 (Vma11p) kDa were
immunoprecipitated by the Vph1p antibody in wild-type sur4,
fen1
, and vma2
cells
(Fig. 7A). We observed
small variations in the relative intensity of bands from experiment to
experiment, and the samples shown in Fig.
7 were chosen to represent the average of the results of three
independent experiments. The data shown in
Fig. 7A are in
agreement with published results (e.g.
Refs.30 and
31) and indicate that the
subunit composition of the V0 domain is normal in
sur4
and fen1
cells.
The anti-Vma1p and anti-Vma2p antibody used in these experiments recognize
free V1 and V1V0 complexes
(24,
30). Radioactive proteins
immunoprecipitated by anti-Vma1p and anit-Vma2p are shown in
Fig. 7, B and
C, respectively. Again, there were small variations is
the intensity of some radioactive bands from experiment to experiment, but
overall our results indicate that the subunit composition of free
V1 and the V1V0 complex in
sur4 and fen1
cells is similar if not
identical to that in wild-type cells and to published data. The
vma2
cells served as a control for cells lacking V1
and V1V0 complexes
(30).
A limitation of these data is that not all V1 subunits are
readily detected including Vma7p, Vma10p, and Vma13p. Vma7p and Vma10p are
probably present in the V1 domains we examined because if either
protein were absence then V1 and V0 would not associate
(3942).
Vma13p is not necessary for V1-V0 association and could
be missing. To determine if it was present in vacuolar membranes isolated from
sur4 cells, sur4
vma13
, and
vma13
cells were transformed with a vector carrying a
VMA13 allele having a Myc epitope inserted immediately downstream of
the methionine start codon
(20). Vacuolar membranes were
isolated by sucrose density gradient centrifugation and fractions from the
gradient were immunoblotted with anti-Myc antibody. The concentration of
Myc-Vma13p in the peak fraction containing vacuolar membranes
(Fig. 7D, fraction
5) was similar in the sur4
vma13
and
vma13
samples, and the overall distribution of Myc-Vma13p was
very similar in the two gradients. Immunoblots of the total cell-free extracts
showed that the concentration of Myc-Vma13p was the same in the two strains as
were the other Vma subunits examined (data not shown). We conclude that Vma13p
associates with V1V0 complexes in sur4
cells.
Elevated Levels of LCBs and LCBPs Do Not Correlate with Reduced
V-ATPase ActivityIt has been noted previously that
sur4 and fen1
cells accumulate LCBs but the
levels have not been quantified nor have the species that accumulate been
determined (11,
12,
43). Likewise, the
concentration of long chain base phosphates (LCBPs) has not been measured.
We quantified LCBs and LCBPs by tagging them after extraction with a
fluorescent reagent followed by HPLC
(23). The analysis was done in
two different strain backgrounds to see how similar or different they might be
and if any differences correlated with mutant phenotypes. The five species of
LCBs are at nearly identical levels in the two wild-type strains
(Table IV). In the two
sur4 strains all five species are elevated and their levels
are similar in the two strain backgrounds except for C18-DHS and C20-DHS,
which are less elevated in strain RCD410 (the W303 background). All five
species are also elevated in the two fen1
strains but there is
more variability between strains and only the C16-DHS species have similar
values.
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LCBP values are also very similar in the two wild-type strains
(Table V) and are quite low as
we have reported for a wild-type strain related to RCD390
(44). All LCBPs show similar
elevations in the two sur4 strains except for C18-DHSP and
C20-DHSP. All LCBPs are also elevated in the two fen1
strains
but the values vary between the strains.
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The elevated levels of LCBs and LCBPs could be responsible for the lack of
ATPase activity. However, if this were the case we would expect
fen1 cells (RCD393), not sur4
cells (RCD389),
to have a greater loss of V-ATPase activity because they have a higher total
level of LCBs and LCBPs (Tables
IV and
V).
To determine directly if LCBs dissociate Vma1p and Vma2p from V0, vacuolar membranes purified from wild-type cells were incubated with increasing concentrations of PHS. After incubation, samples were centrifuged to give a pellet and a supernatant fraction, which where analyzed by immunoblotting for Vma1p and Vma2p. Because PHS has detergent-like properties and could release proteases from vacuoles, we also immunoblotted for CPY to control for proteolysis. In the presence, but not in the absence of PHS, the level of Vma1p and Vma2p in the pellet fraction gradually decreased over the 120-min incubation period (Fig. 8). However, the two proteins did not appear in the supernatant as would be expected if they were dissociating from V0. This result plus the fact that they disappeared (Fig. 8) at the same rate as CPY indicates that they are being digested by proteases. We conclude that LCBs do not dissociate Vma1p and Vma2p from purified vacuolar membranes.
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DISCUSSION |
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To understand why sphingolipids with a C26 acyl group are needed for
V-ATPase activity we determined if V1 domains could associate with
V0 domains on the vacuolar membrane. Vacuolar membranes isolated on
a Ficoll gradient were examined by immunoblotting to determine if
V0 and V1 domains were present. The results showed that
vacuolar membranes from sur4 and fen1
cells
had very low, barely detectable levels of Vma1p, Vma2p, and Vma5p, indicating
that the V1 domain was not associated with the V0 domain
(Fig. 3). Since these three
V1 subunits are present in cell extracts
(Fig. 3) their absence from
purified vacuoles suggests that they mislocalize, do not assemble into a
functional V1 domain or V1 associates abnormally with
V0. The finding that V1 and V0 are found
together in fractions of a sucrose density gradient where vacuolar membranes
are located (Fig. 4, fractions 5 and 6), supports either of the latter two
possibilities. In addition, indirect immunofluorescent microscopy on intact
sur4
and fen1
cells showed that the
V1 subunit Vma2p was bound to vacuoles
(Fig. 5). Together these data
show that V1 is able to associate with V0 on the
cytoplasmic face of the vacuolar membrane to form V1V0
complexes in sur4
cells, but the complexes are not stable in
Ficoll (Fig. 3), have low
ATPase activity (Table III),
and do not acidify vacuoles (Figs.
1 and
2).
Reduced ATPase activity in sur4 cells suggested that the
V1 domain was defective. To test this hypothesis, V1 and
V0 domains were isolated and tested in vitro for
association and for restoration of ATPase activity. Wild-type V1
and sur4
V0 associated to produce
V1V0 complexes with ATPase activity. Wild-type
V0 and sur4
V1 also associated but the
V1V0 complexes had no ATPase activity
(Fig. 6). We also partially
purified cytosolic V1 domains from wild-type and
sur4
cells and assayed them for calcium-dependent ATPase
activity. V1 domains from sur4
cells completely
lacked activity (Fig.
6B). These results establish that V0 domains
in sur4
cells are normal, but the V1 domains are
defective and lack ATPase activity.
To understand why V1 domains lack ATPase activity, we examined
the subunit composition of V1 and V0 and found that they
are the same in sur4, fen1
, and wild-type
cells (Fig. 7). Separate
analysis of Myc-tagged Vma13p showed that it was present in V1
domains in sur4
cells (Fig.
7). The procedures used by us would not have detected the
V1 subunits Vma7p or Vma10p, although we infer that they are
present because if either protein was missing from cells, V1
domains would not associate with V0
(3942).
Thus, lack of a protein subunit is not likely to be the cause of the defect in
ATPase activity in V1 domains present in sur4
cells.
Others have reported that sur4 and fen1
cells contain high levels of LCBs
(11,
12,
43), but the levels have not
been quantified nor have the species been identified. We were concerned that
the high level of LCBs might impair ATPase activity. It was recently shown
that the reduced level of glucan synthase activity in sur4
and
fen1
cells is due to elevated levels of PHS and DHS
(43). Our analysis of LCBs
shows that all five species are elevated in both sur4
and
fen1
mutants in two different strain backgrounds, W303 and
JK9-3d (Table IV). In the
JK9-3d strain background, which we used for the majority of our experiments,
there is a 44-fold increase in total LCBs in sur4
cells and a
63-fold increase in fen1
cells. The same trends hold for the
mutants in the W303 background but the increases are smaller. If elevated LCBs
were responsible for disruption of V-ATPase activity, then we would expect the
activity to be reduced more in fen1
than in
sur4
cells because fen1
cells have a higher
level of LCBs. Our results are just the opposite of this prediction and argue
that elevated LCBs are not responsible for reduced ATPase activity. However,
such arguments cannot eliminate the possibility that LCBs are interfering with
V1 function.
We also compared the total LCB content of fractions from a Ficoll gradient
to see if there was any correlation between reduced V-ATPase activity in
sur4 and fen1
cells and the level of these
compounds. The vacuolar membrane fraction of wild-type cells had a very low,
barely detectable level of LCBs, as did the two fractions below the membrane
fraction (data not shown). The pellet at the bottom of the gradient had the
highest level of LCBs, but the concentration was still very low. The level of
LCBs was higher in the gradient fractions obtained from sur4
and fen1
cells. As with the total LCB values
(Table IV), the levels in the
Ficoll gradient fractions are higher in fen1
cells, suggesting
that it is not the LCBs that are responsible for reduced V-ATPase activity.
LCBPs were not detectable in any of the Ficoll gradient fractions nor in the
cell-free extracts. Most likely they were degraded during the incubation
period when cells were converted to spheroplasts.
We also determined if PHS added in vitro to purified vacuolar
membranes could mimic what occurs in sur4 and
fen1
cells and selectively release Vma1p and Vma2p from
membranes. We found that neither protein was selectively released. In fact,
the membrane V-ATPase was quite resistant to disruption by PHS and only at
higher concentrations did the two proteins start to disappear from the
membrane pellet (Fig. 8). Their
disappearance, however, was not accompanied by their appearance in the soluble
fraction. Rather their concentration decreased as a function of increasing PHS
at the same rate as the luminal vacuolar protein CPY, indicating that protein
loss was probably due to disruption of vacuolar membrane integrity and
subsequent degradation by vacuolar proteases. Thus, wild-type
V1V0 complexes are very resistant to dissociation by
treatment in vitro with PHS.
We also measured LCBPs, which have not been measured, and found that all
five species were elevated in sur4 and fen1
cells. The total level is similar in both wild-type strains and in both
sur4
mutants, but is different in the fen1
strains (Table V). The
differences may be due to strain-specific variation in the activity of
metabolic pathways that make and degrade LCBPs. Again, it seems unlikely that
LCBPs are disrupting V-ATPase activity because in the W303 strain background
their level is higher in fen1
than in sur4
cells, yet the Vma phenotypes are less serve in
fen1
than in sur4
cells
(Table II). However, further
work will be necessary to eliminate the possibility that elevated LCBPs are
responsible for the defective V1 domain in sur4
cells.
How might sphingolipids with a C26 acyl group affect the ATPase activity of
V1? One possibility is that ceramides, which are made in the ER
(reviewed in Ref. 6), play a
role in the assembly of V1 in the ER
(45,
46). Another possibility is
that complex sphingolipids in the Golgi influence maturation of V1
as it transits to the vacuolar membrane. In sur4 cells the
ceramides and sphingolipids with C22 and C24 acyl groups would not substitute
for the normal C26 groups and V1 domains would assemble
incorrectly. The ATPase defect would be less severe in fen1
cells because about one-third of the ceramides and complex sphingolipids have
C26 acyl groups. Alternatively, the RAVE protein complex
(21) has recently been shown
to be necessary for assembly of the V-ATPase
(47) and some step in the
action of RAVE may require sphingolipids with a C26 acyl group. Ficoll, a
polymer of sucrose, may dissociate V1 from V0 by
interacting with one or more V1 subunits that are not correctly
folded or assembled in sur4
cells.
The results presented here are the first to indicate a role for C26 acyl
groups and for sphingolipids in V-ATPase function. Their exact role will
require further characterization of V1 domains in
sur4 cells. Our results suggest that sphingolipids may be
important for the activity of V-ATPases and related ATPases in other
organisms. S. cerevisiae contains another type of V-ATPase located in
the Golgi/endosomal compartments that is identical to the V-ATPase except that
the V0 domain contains Stv1p in place of Vph1p
(48). Our results suggest that
the functionality of this V-ATPase may also require sphingolipids with a C26
acyl group.
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FOOTNOTES |
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To whom correspondence should be addressed: Dept. of Molecular and Cellular
Biochemistry, University of Kentucky College of Medicine, 800 Rose St.,
Lexington, KY 40536. Tel.: 859-323-6052; Fax: 859-257-8940; E-mail:
bobd{at}uky.edu.
1 The abbreviations used are: V-ATPase, vacuolar ATPase; DHS,
dihydrosphingosine; LCB, long chain base; LCBP, long chain base phosphate;
PHS, phytosphingosine; VLCFA, very long chain fatty acid; V1,
peripheral domain of V-ATPase; Vma, vacuolar membrane ATPase; V0,
membrane domain of V-ATPase; ER, endoplasmic reticulum; MES,
4-morpholineethanesulfonic acid; MOPS, 4-morpholinepropanesulfonic acid; HPLC,
high pressure liquid chromatography; FITC, fluorescein isothiocyanate.
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ACKNOWLEDGMENTS |
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REFERENCES |
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