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INTRODUCTION |
PtdCho1 is the major
membrane phospholipid in mammalian cells, and regulation of its
biosynthesis and turnover is critical to maintaining membrane structure
and function. CCT is a key regulatory step in PtdCho biosynthesis in
mammalian cells (for reviews, see Refs. 1 and 2). CCT activity is
regulated primarily through the interaction of lipid modulators with
its amphipathic
-helical domain, which stimulate (1) or inhibit
(3-5) activity by modulating the Km for CTP (6).
CCT is also a phosphoprotein (1) that is phosphorylated on multiple
serine residues within its carboxyl-terminal domain (7). The presence
of the highly phosphorylated carboxyl-terminal domain attenuates enzyme
activation by lipid modulators (8).
Enforced CCT
(9) or CCT
(10) expression results in an
acceleration of PtdCho biosynthesis measured by the incorporation of
radiolabeled Cho into PtdCho. However, transient overexpression of COS
cells with CCT
did not lead to a significant increase in the amount
of PtdCho per cell despite the increased biosynthetic rate (9). In
transiently transfected COS cells, there was a concomitant increase in
GPC and PCho, suggesting that elevated PtdCho degradation compensated
for increased synthesis (9). An indication of a balanced relationship
between PtdCho synthesis and degradation was also found in
ras-transformed cell lines. Transformation with
ras oncogenes accelerated both PtdCho synthesis (11-13) and
PtdCho turnover (14-17). Elevated pools of two PtdCho breakdown
products, PCho and GPC, were observed in these experiments (11-13,
15-18), although the activation of Cho kinase also makes a significant
contribution to the PCho pool (12). Another example of balanced PtdCho
synthesis and degradation was found during the G1 stage of
the cell cycle (19). PtdCho synthesis increased in G1;
however, the total phospholipid mass was maintained as a result of a
concomitant increase in PtdCho degradation.
Taken together, these experiments suggest the existence of a system
that coordinates PtdCho biosynthesis and degradation to maintain a
constant membrane phospholipid content. The goals of this study were to
determine if the coupling between CCT activity and PtdCho turnover is a
manifestation of a general mechanism for phospholipid homeostasis and
to investigate the phospholipase(s) responsible for regulated PtdCho degradation.
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EXPERIMENTAL PROCEDURES |
Materials--
[5,6,8,9,11,12,14,15-3H]Arachidonic
acid (specific activity 100 Ci/mmol),
n-[4-3H]butanol (specific activity 10 Ci/mmol), [1-3H]ethanolamine (specific activity 30 Ci/mmol), [methyl-3H]LPC (specific activity 80 Ci/mmol), [methyl-3H]Cho (specific activity 85 Ci/mmol), and phospho[methyl-14C]choline
(specific activity 56 mCi/mmol) were from American Radiolabeled Chemicals. LysoPAF
(1-O-[3H]octadecyl-sn-glycero-3-phosphocholine
(specific activity 170 Ci/mmol) was from Amersham Pharmacia Biotech.
Preadsorbent Silica Gel G and Silica Gel H thin layer chromatography
plates were from Analtech. LPC, LPE, lysoPAF, phospholipid, and neutral
lipid standards were from Avanti Polar Lipids. Brefeldin A and monensin
were from Calbiochem. Bromoenol lactone was from Cayman Chemical.
[2-3H]Glycerol (specific activity 200 mCi/mmol) was from
NEN Life Science Products. Doxycycline and choline metabolite standards were from Sigma. All other materials were reagent grade or better.
Cell Lines and Growth Conditions--
The CCT.12 cell line was
established in our laboratory by stable transfection of HeLa
Tet-OnTM cells (CLONTECH) using the
pTRE vector carrying a CCT
cDNA (20). Vector control cells
(CCT.00) were isolated following stable transfection of HeLa
Tet-OnTM cells using empty pTRE vector and selection in
medium with hygromycin. Cells were routinely grown in Dulbecco's
modified Eagle's medium (BioWhittaker) containing 10% fetal bovine
serum (Summit Biotechnology) at 37 °C in 5% CO2, 95%
air. Induction of CCT
overexpression in CCT.12 cells did not alter
the proliferative rate of the cultures or the final cell density at
100% confluence. Cell density was determined using a hemocytometer,
and viability was determined by trypan blue exclusion.
CCT Activity--
Cells were washed twice with
phosphate-buffered saline on ice and harvested by scraping into 1 ml of
the same buffer followed by centrifugation. The cell pellets were
resuspended in lysis buffer (10 mM HEPES, pH 7.4, 10 mM NaCl, 1 mM EDTA, 2 mM
dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, 2%
aprotinin, 1 µg/ml leupeptin, 50 mM NaF, 100 mM Na3VO4) and sonicated three
times for 30 s in a cup-horn sonicator (model W225-R, Heat
Systems, Ultrasonics, Inc.) at 80% duty cycle and maintained at
4 °C. The standard CCT activity assay contained an aliquot of the
cell lysate mixed with 125 mM bis-Tris-HCl, pH 6.5, 0.5 µCi of phospho[methyl-14C]Cho, 1 mM phosphocholine, 2 mM CTP, 20 mM
MgCl2, 50 µM PtdCho/18:1 (1:1) in a final
volume of 40 µl. The incubations were for 10 min at 37 °C and were
stopped by placing the samples on ice and adding 5 µl of 0.5 M EDTA. CDP-[14C]choline formation was
determined by thin layer chromatography (21). CCT-specific activity was
calculated from a series of assays that were linear with time and
protein. Protein concentration was determined according to the method
of Bradford (22) using
-globulin as a standard.
Phospholipase D Activity--
Cells were grown in 100-mm dishes
to 20% confluence and either incubated with doxycycline (2 µg/ml)
for 48 h or left untreated. At 48 h, phospholipase D activity
was assayed by measuring phosphatidylbutanol formation
(transphosphatidylation) following the addition of 0.5% [3H]butanol (10 µCi/ml) to the cultures and incubation
for 30 min at 37 °C. After harvesting the cells, the phospholipids
were extracted as described below and phosphatidylbutanol was
quantitated following fractionation of the radiolabeled lipids by thin
layer chromatography on Silica Gel H layers developed with
chloroform/methanol/acetic acid (90:10:10, v/v). Phosphatidylbutanol
was identified by co-migration (RF = 0.8) with a
standard. As a positive control, cells were treated with 100 µM phorbol myristate acetate 60 min prior to the addition
of [3H]butanol.
Phospholipid Mass Determination--
Cells were grown to 80%
confluence, washed twice with phosphate-buffered saline, and harvested
by scraping followed by centrifugation. Aliquots of a cell suspension
were counted with a hemocytometer, and the bulk of the cells were
collected by centrifugation for quantitation of the phospholipid mass.
Cell pellets were extracted using the method of Bligh and Dyer (23),
and the phospholipid in the lower phase was quantitated as described by
Charles and Stewart (24) using PtdCho to construct a standard curve.
Metabolic Labeling--
Cultures were grown in 60-mm dishes to
80% confluence and pretreated with 2 µg/ml doxycycline for 48 h
during exponential growth to induce the overexpression of CCT
as
indicated. Cells were incubated with 5 µCi/ml [3H]Cho,
5 µCi/ml [3H]glycerol, 2 or 1 µCi/ml
[methyl-3H]LPC, 5 µCi/ml
1-O-[3H]lysoPAF, or 1 µCi/ml
[3H]arachidonic acid for indicated times. In the
experiments with 50 µM additional exogenous unlabeled
lysophospholipids, cells were radiolabeled with 1 µCi/ml
[methyl-3H]LPC or 5 µCi/ml
[3H]ethanolamine for 2 h prior to the start of the
experiment and the medium was then removed. Fresh medium with or
without 50 µM of the indicated lysophospholipid was added
to the cells for 6 h. At the indicated times, cells were washed
twice with phosphate-buffered saline on ice and harvested by scraping
into 1 ml of the same buffer followed by centrifugation. Medium from
the cells was also collected prior to washing, and non-adherent cells
and cell debris were removed from the medium by centrifugation. Cells
or medium were extracted using a two-phase system (23) to separate the water-soluble and chloroform-soluble metabolites, and the amount of
radiolabel incorporated into each phase was determined by scintillation counting. The volume of the media samples was reduced in a Speed Vac
concentrator (Savant) prior to lipid extraction. The distribution of
the radiolabel among the PtdCho or PtdEtn precursors or metabolites in
the soluble fraction was determined by thin layer chromatography on
Silica Gel G preadsorbent plates developed in 95% ethanol, 2%
ammonium hydroxide (1:1, v/v). Migration of radiolabeled CDP-choline (RF = 0.68), Cho (RF = 0.2), ethanolamine (RF = 0.18), GPC
(RF = 0.5), GPE (RF = 0.85), PCho (RF = 0.4), or
phosphoethanolamine (RF = 0.8) was confirmed
with standards. The chloroform-soluble phospholipids and neutral lipids
were separated on Silica Gel H thin layer chromatography plates
developed with chloroform/methanol/acetic acid/water (50:25:8:4, v/v).
The major radiolabeled product after incubation with
[methyl-3H]Cho,
[methyl-3H]LPC, or
1-O-[3H]lysoPAF was PtdCho
(RF = 0.45). Fractionation and identification of
neutral lipids was performed using Silica Gel H plates developed with
chloroform/methanol/acetic acid (98:2:1, v/v) where all phospholipids migrated with an RF of 0.1, diacylglycerol with
an RF of 0.85, and triacylglycerol with an
RF of 0.93. The migrations of LPC, PtdCho,
PtdEtn, phosphatidylserine, phosphatidylinositol, neutral lipids
(triacylglycerol, diacylglycerol), and water-soluble phospholipid precursors or metabolites were confirmed with standards.
Drug Treatments--
Brefeldin A (1 mg/ml) and monensin (1 mM) stocks were prepared in ethanol. Cells were incubated
with doxycycline for 48 h and radiolabeled with 5 µCi/ml
[methyl-3H]Cho for the last 2 h prior to
addition of the indicated drug. The label remained in the cells for the
duration of the drug treatment. Brefeldin A (5 µg/ml) was added, and
both cells and media were harvested on ice at 15, 30, 60, 120, and 240 min. Alternatively, monensin (10 µM) was added, and both
cells and media were harvested on ice at 1, 2, and 15 h. In a
separate series of experiments, cells were prelabeled with 2 µCi/ml
[methyl-3H]LPC for 2 h, the label was
removed from the cells, and unlabeled LPC (50 µM final)
plus brefeldin A (5 µg/ml) or unlabeled LPC (50 µM)
only was subsequently added. Cells and media were harvested at 15, 30, 60, 120, and 240 min. Extraction and thin layer fractionation of
radiolabeled Cho metabolites was performed on both cell and media
samples as described above. The disruption of the Golgi apparatus by
brefeldin A was confirmed with confocal fluorescent microscopy using
Texas RedTM-labeled wheat germ agglutinin as a marker for
the Golgi apparatus.
Confocal Fluorescent Microscopy--
Cells were grown on
Lab-Tek®II Chamber SlidesTM to 60% confluence
and treated with 5 µg/ml brefeldin A for 15 min or left untreated. Cells were washed and fixed with 3.7% formaldehyde for 20 min and
permeabilized with 0.2% Triton X-100 for 10 min. Nonspecific binding
was blocked with 1% bovine serum albumin in phosphate-buffered saline
for 60 min. Cells were treated with 50 µg/ml Texas
RedTM-conjugated wheat germ agglutinin that reacts with
N-acetylglucosamine glucoconjugates in the Golgi apparatus
(25). Cells were visualized after staining using a Leica DM-IRBE
confocal microscope equipped with a Leica TCS NT scanning laser.
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RESULTS |
PtdCho Biosynthesis in HeLa Cells Overexpressing CCT
--
The
human HeLa cell line, CCT.12 (20), overexpressed the rodent CCT
in
response to doxycycline (Fig. 1). The
maximum level of CCT
overexpression was about 25-fold higher than
the endogenous activity prior to addition of 2 µg/ml doxycycline.
Both [methyl-3H]Cho (Fig.
2A) and
[3H]glycerol (Fig. 2B) were incorporated into
PtdCho at a higher rate following CCT
overexpression. The rate of
[3H]glycerol incorporation into PtdCho was about 2.2 times higher when the CCT
level was maximal. These data showed that
the rate of PtdCho biosynthesis increased in response to elevated CCT
activity and verified the observations made in COS cells transiently
transfected with either CCT
(9) or CCT
(10) in a stable cell
line. The enhanced rate of PtdCho biosynthesis was not associated with a faster rate of proliferation or a change in morphology when cells
were examined by transmission electron microscopy (data not shown).

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Fig. 1.
CCT activity following induction of
CCT expression with doxycycline. CCT.12
cultures were grown to 20% confluence. and incubations were continued
in the presence ( ) or absence ( ) of 2 µg/ml doxycycline. Cell
lysates were prepared and assayed for CCT activity at the indicated
times after doxycycline addition as described under "Experimental
Procedures." The data points are the average of duplicate
determinations from a representative experiment.
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Fig. 2.
Enforced CCT
expression stimulated the incorporation of Cho and glycerol into
PtdCho but did not increase the cellular phospholipid mass.
Panel A, enhanced radiolabeling of CDP-choline and PtdCho in
cells overexpressing CCT . CCT.12 or vector control cell cultures
were grown to a final density of 1.5 × 106
cells/60-mm dish in the presence (+) or absence ( ) of 2 µg/ml
doxycycline for 48 h. [methyl-3H]Cho (5 µCi/ml) was then added to duplicate cultures for 6 h, and the
cells were harvested. Extraction and quantitation of the radiolabeled
choline metabolites by thin layer chromatography were performed as
described under "Experimental Procedures." Panel B,
overexpression of CCT stimulates incorporation of
[3H]glycerol into PtdCho. CCT.12 cells were incubated
with ( ) or without ( ) 2 µg/ml doxycycline for 48 h. Cells
reached a density of 5.4 × 105/dish, and
[3H]glycerol (5 µCi/ml) was added to duplicate
cultures. Cells were harvested at the indicated times thereafter.
Extraction, fractionation, and quantitation of the radiolabeled
glycerolipids were performed as described under "Experimental
Procedures." Panel C, cellular content of phospholipid
remains constant following CCT overexpression. CCT.12 cell cultures
were incubated with (+) or without ( ) 2 µg/ml doxycycline for
48 h and compared with vector control cultures. Phospholipid mass
was quantitated in duplicate dishes as described under "Experimental
Procedures" and normalized to cell number. The results from two
independent experiments were combined. Standard errors were calculated
from quadruplicate determinations.
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PtdCho constituted 70% of the total membrane phospholipid in HeLa
cells both with and without CCT
overexpression as determined by
equilibrium radiolabeling with [32P]orthophosphate (data
not shown). The cellular phospholipid content was quantitated in cells
after 48 h of induction of CCT
overexpression with doxycycline
and compared with control cells with basal CCT activity (Fig.
2C). The HeLa CCT.12 cells did not exhibit a substantive increase in membrane phospholipid content in response to CCT
overexpression. These data suggested that the elevated rate of PtdCho
biosynthesis was accompanied by a compensatory level of PtdCho
degradation, resulting in a PtdCho pool of the same size with a higher
radiochemical specific activity. These data, together with the data in
Fig. 2 (A and B), were consistent with an
accelerated turnover of both the Cho headgroup and the glycerol
backbone from the PtdCho pool.
PtdCho Degradation--
Our results indicating the metabolic loss
of the glycerol backbone from the phospholipid pool suggested that
phosphatidic acid was not a likely degradation product. We verified
this prediction experimentally by direct assay of cellular
phospholipase D activity before and after CCT
overexpression in the
CCT.12 HeLa cultures (Fig. 3).
Phosphatidylbutanol formation was linear over 60 min and increased as a
function of increasing butanol concentration in the medium up to 0.7%.
The data showed that phospholipase D activity did not increase in
response to enhanced PtdCho biosynthesis following induction of CCT
activity with doxycycline. Phorbol myristate acetate at 100 µM was added as a positive control to stimulate
phospholipase D activity under the same conditions (Fig. 3). These
results demonstrated the existence of a typical phospholipase D
response in the HeLa cells and confirmed the absence of elevated phospholipase D activity in CCT overexpression cells. Diacylglycerol and phosphocholine are the products of phospholipase C degradation, and
diacylglycerol can be converted to phosphatidic acid or triacylglycerol in vivo. Overexpression of CCT in cells labeled with
[3H]glycerol did not result in enhanced formation of
diacylglycerol or triacylglycerol (data not shown). These experiments
were consistent with the conclusion that neither phospholipase D nor C
was involved in PtdCho turnover in response to accelerated PtdCho
biosynthesis.

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Fig. 3.
Enforced CCT
expression did not elevate phospholipase D activity. CCT.12
cell cultures were incubated with or without 2 µg/ml doxycycline to
induce CCT expression for 48 h prior to the assay. Phorbol
myristate acetate (100 µM; TPA) was added as a
positive control to half of the cultures. [3H]Butanol (10 µCi/ml) was added to start the transphosphatidylation assay;
following 30 min of incubation, the cells were washed and harvested.
Radiolabeled phosphatidylbutanol formation was quantitated following
thin layer chromatography as described under "Experimental
Procedures." Standard errors were calculated from quadruplicate
determinations from a representative experiment. The experiment was
performed twice.
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The route for PtdCho breakdown was determined by radiolabeling the
PtdCho pool in cells with a tracer amount of
[3H-methyl]LPC. This approach avoided the
complications in the interpretation of results that arise from high
background labeling of the PCho and Cho precursor pools when using
[3H-methyl]Cho to prelabel PtdCho.
[3H-methyl]LPC was readily acylated, and
[3H]PtdCho constituted >98% of the total cellular
radiolabel (data not shown). The [3H]PtdCho was
subsequently degraded, and two radiolabeled metabolic products, PCho
and GPC, were detected inside the cells (Fig.
4). GPC release from PtdCho was
dramatically elevated in cells induced to overexpress CCT
(Fig.
4A). In contrast, PCho was not affected by CCT
overproduction (Fig. 4B). The lack of an effect on PCho release provided an internal control that underscored the specificity of the GPC response to enhanced PtdCho biosynthesis. The accumulation of GPC suggested that GPC was not readily metabolized to Cho or PCho.

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Fig. 4.
CCT overexpression
stimulates PtdCho degradation to GPC. Panel A,
stimulated GPC formation associated with CCT overexpression.
Panel B, the rate of PtdCho degradation to PCho remains the
same with CCT overexpression. CCT.12 cell cultures were incubated
with ( ) or without ( ) 2 µg/ml doxycycline to induce CCT
overexpression for 48 h prior to radiolabeling with
[methyl-3H]LPC (2 µCi/ml), which was added
at the start of the experiment. Cells were washed and harvested at the
times indicated. The radiolabeled LPC, PtdCho, and water-soluble
degradation products from duplicate cultures were fractionated,
identified by thin layer chromatography, and quantitated as described
under "Experimental Procedures." Standard errors were calculated
from quadruplicate determinations and from two independent
experiments.
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GPC Is Released into the Medium--
The amount of GPC produced as
a result of CCT
overexpression was measured in the cells and in the
medium. HeLa CCT.12 cells treated with doxycycline for 48 h were
labeled with [3H-methyl]Cho for the last
24 h, the label was removed, and the radiolabeled GPC was measured
inside and outside the cells 24 h later in the continued absence
or presence of doxycycline (Fig. 5).
Almost all of the radiolabeled GPC (
90%) accumulated in the medium
of cells under conditions of either CCT
overexpression or basal
activity (Fig. 5). Radiolabeled Cho and PCho were also identified in
the medium, but it is difficult to determine whether these metabolites
were derived directly from PtdCho or from the degradation of a portion
of the extracellular GPC. These data indicated that the GPC arising
from the degradation of PtdCho accumulated in both inside and outside
the cells.

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Fig. 5.
GPC exits from cells. CCT.12 cultures
were preincubated with or without doxycycline (2 µg/ml) as indicated
for 48 h. [methyl-3H]Cho (5 µCi/ml) was
added for the latter 24 h of the preincubation. The radioactive
medium was removed at 48 h, the cells were washed, fresh medium
without radiolabel was added, and the cultures were incubated for
an additional 24 h. Both cells and medium were harvested and
extracted from duplicate cultures. Cho-derived metabolites were
fractionated by thin layer chromatography and analyzed as
described under "Experimental Procedures." Standard errors
were calculated from quadruplicate determinations and from two
independent experiments.
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The possibility that GPC was released by a secretion mechanism was
investigated using either brefeldin A or monensin to block secretion.
Brefeldin A block secretion by causing the redistribution of the Golgi
apparatus into the endoplasmic reticulum (26), and monensin inhibits
the function of the Golgi and lysosomes through the disruption of
H+, Na+, and K+ gradients across
cellular membranes (27). The efficacy of 5 µg/ml brefeldin A was
verified by immunofluorescent staining of the cells using Texas
RedTM-conjugated wheat germ agglutinin. After 15 min of
incubation with the drug, the juxtanuclear staining of the Golgi
apparatus completely disappeared (data not shown). Cells were incubated with monensin (10 µM) for times ranging from 2 to 14 h prior to the measurement of GPC release. HeLa CCT.12 cells were
induced to overexpress CCT
and prelabeled prior to the addition of
either inhibitor. The accumulation of radiolabeled GPC in the medium was not reduced by the drugs (data not shown). These results showed that GPC did not exit the cells by a Golgi-mediated pathway.
iPLA2 Mediates the GPC Response--
The generation of
GPC in response to phospholipid overproduction suggested the
involvement of a regulated PLA activity. An earlier report from our
laboratory concluded that the cytoplasmic PLA2 was a growth
factor-responsive activity and probably did not mediate the regulated
phospholipid turnover observed during G1 phase of the cell
cycle (28). The group VI iPLA2 enzymes are not responsive
to extracellular stimuli and are speculated to have basal housekeeping
activity that is visible through its involvement in phospholipid
remodeling (29). Therefore, we tested whether the iPLA2 was
a participant in the pathway of GPC formation in response to PtdCho
overproduction (Fig. 6). BEL is a
specific inhibitor of iPLA2 and is diagnostic for the
identification of iPLA2 in contrast to other cellular
phospholipases A2 (29). The efficacy of BEL was first
examined in the HeLa cells by monitoring the incorporation of
[3H]arachidonic acid into phospholipid by the remodeling
pathway (Fig. 6A). BEL inhibition of arachidonic acid
incorporation was dose-dependent, and incorporation was
reduced to 50% of control cells without inhibitor. These data are in
complete agreement with the results reported previously for the BEL
inhibition of arachidonic acid incorporation mediated by
iPLA2 in intact cells (17). We then investigated the effect
of 25 µM BEL on GPC production in cells by radiolabeling
with [3H-methyl]LPC. Overproduction of PtdCho
was induced by doxycycline-dependent overexpression of
CCT
in the CCT.12 cells 48 h prior to the start of the
experiment. The formation of GPC in the doxycycline-treated cultures
was dramatically reduced by BEL (Fig. 6B). In a separate experiment shown in Fig. 6C, BEL reduced the level of GPC
produced in doxycycline-treated cultures and also reduced the GPC
levels in control cultures in a dose-dependent manner. In
these experiments total GPC (cells + medium) was quantitated. These
results clearly pointed to a role for the iPLA2 in
regulating the membrane phospholipid content, either by initiating
cleavage of the sn-2 acyl moiety of PtdCho, which would be
further degraded by a lysophospholipase activity, or perhaps by
hydrolyzing both the sn-1 and sn-2 acyl chains to
yield GPC.

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Fig. 6.
BEL inhibition of GPC formation.
Panel A, BEL was added to CCT.12 cultures at the indicated
concentrations for 30 min prior to the addition of
[5,6,8,9,11,12,14,15-3H]arachidonic acid (1 µCi/ml) at
the start of the experiment. The cells were washed, harvested, and
extracted after 2 h of incubation with the radiolabel, and the
incorporation of [3H]arachidonic acid into phospholipids
was determined as described under "Experimental Procedures."
Panel B, CCT.12 cells were grown for 48 h with ( ,
) or without ( , ) 2 µg/ml doxycycline. BEL (25 µM) was added to the indicated cultures (- - -) for 30 min prior to the addition of [methyl-3H]LPC (1 µCi/ml) to start of the experiment. At the indicated times, duplicate
cultures were harvested and the cells and media extracted. Radiolabeled
GPC was quantitated following thin layer chromatography as described
under "Experimental Procedures." Panel C, CCT.12 cells
were grown for 48 h with ( ) or without ( ) 2 µg/ml
doxycycline. BEL was added at the indicated concentrations to the
cultures for 30 min prior to the addition of
[methyl-3H]LPC (1 µCi/ml) at the start of
the experiment. Duplicate cultures were harvested and extracted after
2 h of further incubation. Radiolabeled GPC was isolated from
cells and medium and quantitated following thin layer chromatography as
described under "Experimental Procedures." Standard errors were
calculated from quadruplicate determinations in a representative
experiment, which was independently performed twice, except in
panel B, where the error bars
represent the range of duplicate determinations in a single
experiment.
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PtdCho Degradation to GPC Is a General Response to Excess
Phospholipid--
GPC release from PtdCho could be singularly coupled
to CCT activity, or it could represent a more generalized response to excess PtdCho formation. Stimulating excess PtdCho production by
providing cells with a large amount of exogenous LPC tested this
hypothesis. The PtdCho pool was prelabeled with a tracer amount of
[3H-methyl]LPC for 2 h. The label was
removed, and the cells were treated with 50 µM unlabeled
LPC. This amount of LPC does not lyse cells and is not toxic within the
time frame of this experiment (3, 4). The HeLa cells were not treated
with doxycycline; therefore, CCT activity was at the basal level. GPC
production was stimulated in cells following the addition of exogenous
50 µM LPC, and radiolabeled GPC accumulated with time as
the cells degraded the prelabeled PtdCho (Fig.
7). These data clearly showed that the
stimulation of GPC formation from PtdCho was a generalized response to
an elevated amount of cellular PtdCho and not specifically coupled to
CCT activity.

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Fig. 7.
Synthesis of excess PtdCho by acylation of
LPC stimulates GPC production. CCT.12 cultures were incubated with
[methyl-3H]LPC (2 µCi/ml) for 2 h. The
medium containing the radiolabeled LPC was removed, the cells were
washed, and fresh medium either with no additions ( ) or with 50 µM unlabeled LPC ( ) was added at the start of the
experiment. Cells (8 × 105/60-mm dish) and medium
were harvested from duplicate dishes at the indicated times thereafter.
Extraction and fractionation of radiolabeled LPC-derived metabolites
was performed as described under "Experimental Procedures."
Standard errors were calculated from quadruplicate determinations and
two independent experiments.
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PtdEtn constitutes
24% of the membrane phospholipid pool in the
HeLa cells (data not shown), and we investigated whether the cells
responded to overproduction of PtdEtn as well as PtdCho. Cells were
prelabeled with [3H-methyl]LPC for 2 h,
the medium containing the radiolabel was removed, and the cells were
incubated with 50 µM LPC, LPE, or lysoPAF for 6 h.
LysoPAF is quickly converted to 1-alkyl-2-acyl-PtdCho (data not shown),
a molecular species different from the diacyl-PtdCho derived from LPC.
We postulated that an excessive amount of exogenous lysoPAF would also
trigger the breakdown of PtdCho to GPC if the response was not specific
for diacyl-PtdCho. The radiolabeled GPC in the cells and medium was
elevated following incubation with all of the lysophospholipids (Fig.
8A). These data demonstrated that excess PtdEtn and alkyl phospholipids also triggered the GPC
response.

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Fig. 8.
Synthesis of excess PtdCho or PtdEtn
stimulates GPC and GPE formation. Panel A, overloading
with different lysophospholipids stimulates GPC levels. CCT.12 cultures
were incubated with [methyl-3H]LPC (1 µCi/ml) for 2 h. The medium containing the radiolabeled LPC was
removed, the cells were washed, and fresh medium either with no
additions or with 50 µM unlabeled lysophospholipid was
added as indicated. Cells (1 × 106/60-mm dish) and
medium were harvested 6 h after addition of the fresh medium. The
radiolabeled GPC in the cells and medium was isolated and quantitated
as described under "Experimental Procedures." Panel B,
PtdEtn degradation to GPE responds to phospholipid overload. CCT.12
cultures were incubated with [1-3H]ethanolamine (5 µCi/ml) for 2 h. The medium containing the radiolabeled
ethanolamine was removed, the cells were washed, and fresh medium
either without lysophospholipid or with 50 µM unlabeled
lysophospholipid was added as indicated. Cells (8 × 105/60-mm dish) and medium were harvested 6 h after
addition of the fresh medium. The radiolabeled GPE in the cells and
medium was isolated and quantitated as described under "Experimental
Procedures." Standard errors were calculated from quadruplicate
determinations of representative experiments for each panel.
Experiments in each panel were independently performed twice.
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The ability of excess PtdEtn to stimulate GPC production led us to
investigate whether the PtdEtn pool also responded to the excess
phospholipid by degradation to GPE. Cells were incubated with
[3H]ethanolamine for 2 h which efficiently
radiolabeled PtdEtn, but GPE was not detectable at that time. The
radiolabeling medium was removed, and the cells were incubated
with 50 µM LPC or LPE for 6 h. Radiolabeled GPE was
recovered from both cells and medium, and GPE levels were significantly
elevated in cultures treated with either lysophospholipid (Fig.
8B). These data indicated that the amount of cellular PtdEtn
was also regulated in a manner similar to PtdCho and that PtdEtn
degradation to GPE was a component of a general cellular response to
excess membrane phospholipid.
Consequences of PtdCho Overload--
The identification of
GPC (or GPE) as the major breakdown product in response to excess
phospholipid suggested that removal of the acyl moieties from PtdCho
was an essential event in the cellular regulatory response. LysoPAF
differs from LPC in that an alkyl group rather than an acyl group
occupies the sn-1 position of the glycerol backbone, and the
ether linkage is resistant to cleavage by phospholipases. HeLa CCT.12
cells were treated with doxycycline for 48 h to overexpress CCT
and synthesize PtdCho at a stimulated rate. Control, untreated cells
and induced cells were incubated with a tracer amount of
[3H-alkyl]lysoPAF alone or in the presence of
10 µM unlabeled lysoPAF for 3 days (Fig.
9A). Ninety percent of the
radiolabel was incorporated into cellular lipids within 24 h and
remained associated with the cellular lipids for the entire experiment.
At 10 h, alkylacyl-PtdCho was the major metabolite and was
converted to alkylacyl-PtdEtn with continued incubation (Fig.
9B). Less than 10% of the alkylacyl-phospholipid was
converted to alkylacyl- or alkyldiacylglycerol, indicating the loss of
the phosphocholine headgroup. Radiolabeled water-soluble degradation
products were not detected, and a very small level of lysoPAF (<1%)
was detected throughout the experiment (data not shown). These results
showed that the alkylacyl-phospholipids accumulated in the cell
membranes and suggested that perhaps the cells would have a greater
phospholipid content. However, when the cellular phospholipid mass was
determined (described under "Experimental Procedures") following
incubation with up to 30 µM lysoPAF, the phospholipid did
not significantly increase above the normal value of
80
µg/106 cells (data not shown).

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Fig. 9.
Inability to degrade excess PtdCho to GPC
inhibits cellular proliferation. Panel A,
alkylacylphospholipids accumulate in cells. CCT.12 cultures were
incubated with lysoPAF
(1-O-[3H]octadecyl-glycerophosphocholine,
5 µCi/ml) with ( ) or without ( ) 10 µM unlabeled
lysoPAF added at the start of the experiment. At indicated times, cells
and medium were harvested and extracted, and the radioactivity in each
compartment was quantitated as described under "Experimental
Procedures." Standard errors were calculated from quadruplicate
determinations in a representative experiment. The experiments with or
without 10 µM LPAF were performed independently.
Panel B, cell-associated radiolabeled lysoPAF and its
metabolites in the lipid fraction from the experiment described in
panel A, were fractionated and quantitated by thin-layer
chromatography as described under "Experimental Procedures." Lipid
profiles were calculated using the mean data points from panel
A. Standard errors were not calculated. Panel C,
lysophospholipid at the indicated concentrations was added to cell
cultures in two doses 24 h apart during exponential growth. Cells
were harvested and counted 24 h after the second addition of
lysophospholipid. Viability was 85% in all cultures except following
treatment with 50 µM lysophospholipid. Standard errors
were calculated from quadruplicate determinations in a representative
experiment. The experiment was independently performed twice.
Panel D, lysoPAF was added to duplicate cultures at the
indicated concentrations daily on days 1, 2, and 3. Cells were
harvested and counted on day 4. Viability was 90% except on day 4 following treatment with the 30 µM dose, where it dropped
to 70%. The ranges of data from duplicate determinations of the cell
number were smaller than the points on the graph. The data were from a
representative experiment.
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Previous work suggested that lysoPAF (>20 µM) had a
detrimental effect on cell growth (30). Therefore, we investigated
whether the accumulation of alkyl phospholipids reduced the rate of
cell proliferation during extended incubation. Cells were incubated for
48 h during exponential growth with two doses up to 50 µM of either LPC or lysoPAF at 24-h intervals (Fig.
9C). The total number of cells counted after treatments with
lysoPAF were dramatically reduced compared with the control cultures
treated with LPC. Viability was high (
85%) at lysophospholipid
concentrations <50 µM for both LPC and LPAF. Cell growth
was only moderately affected by the 48-h incubation with 2 doses of 50 µM LPC (1.21 × 106 total cells).
Viability was slightly reduced but remained high (70%) following
treatment at the 50 µM dosage of either LPC or LPAF. The
growth rate and viability count for the 50 µM LPC
condition after a single dose and 24 h of culture was identical to
that for the control condition.
The data suggested that accumulation of the cellular
alkyl-phospholipids would correlate with the inhibition of cell
proliferation. This point was tested directly by determining the growth
rates of cultures incubated with lysoPAF up to 30 µM
added daily (Fig. 9D). The number of viable cells was
reduced as a function of increasing lysoPAF concentration; however,
significant cell death (30%) was only evident after 72 h of
treatment and the third dose of 30 µM lysoPAF. These data
demonstrated that accumulation of the alkyl-phospholipids interfered
with cell proliferation. Flow cytometric analysis of arrested cultures
did not reveal that the cells accumulated at a specific phase of cell
cycle progression (data not shown), and TdT-dependent TUNEL
assays of arrested cultures did not indicate the presence of the
specific DNA fragmentation associated with apoptosis (data not shown).
 |
DISCUSSION |
A large body of work has identified CCT as a key regulator of
PtdCho synthesis. Both the positive and negative regulation of CCT
activity by lipid modulators and phosphorylation exert control over
phospholipid formation (1, 2, 6, 8); however, regulation of
biosynthesis at the CCT step is not the only mechanism to control
PtdCho content. Enforced expression of CCT
significantly accelerates
PtdCho biosynthesis, but does not increase cellular phospholipid
content (9) (Fig. 2). This is surprising when one considers that PtdCho
constitutes 72% of the phospholipid in HeLa cells (data not shown).
Another method to deregulate PtdCho synthesis is to overload cells with
exogenous LPC. LPC is a potent inhibitor of CCT but circumvents the
de novo PtdCho biosynthetic pathway by being directly
acylated to form PtdCho (3). Cells balance increased PtdCho production
with an increased rate of phospholipid degradation to GPC, thereby maintaining the size of the PtdCho pool. The exit and accumulation of
GPC in the culture medium suggests that the glycerol and choline moieties of PtdCho are not readily reutilized during exponential growth
of cultured cells.
The perturbation of cell physiology with enforced CCT expression and
overloading with exogenous LPC reveals the regulated deacylation of
PtdCho to GPC as a key process in membrane phospholipid homeostasis.
PtdCho is the predominant membrane phospholipid and the precursor to
the other two major phospholipids, sphingomyelin (31) and PtdEtn (32),
in cultured animal cells. The metabolic interconnection between the
major phospholipids means that the synthesis and degradation of PtdEtn
(and likely sphingomyelin) reflects that of PtdCho. Overloading the
PtdCho pool with exogenous lysoPAF (a molecular species of LPC) also
increases the PtdEtn pool as alkylacyl-PtdCho is converted to
alkylacyl-PtdEtn (Fig. 9B). Excess PtdCho can lead not only
to stimulated GPC production but also to excess PtdEtn, and this is
reflected by the increased production of GPE (Fig. 8B).
Conversely, the stimulated production of GPC in response to overloading
cells with LPE (Fig. 8A) can be explained if PtdCho
accumulates as a result of a block in its conversion to PtdEtn. Cells
maintain a specific profile of phospholipids when PtdCho synthesis is
accelerated (data not shown) by degrading PtdEtn to GPE in addition to
degrading PtdCho to GPC. Cells also maintain a specific phospholipid
profile when PtdCho synthesis is inhibited by antineoplastic ether
lipids such as ET-18-OCH3 (5). The data reveal the general
nature of the cellular response to excess phospholipid and suggest that
the maintenance of the PtdCho pool is synonymous with the preservation
of total membrane phospholipid content and composition in cultured
cells. The rapid and virtually complete uptake of exogenous
phospholipid (Fig. 9A) suggests that cells must have a
physiological mechanism to adjust to variations in serum lipid
composition while maintaining their typical membrane composition and function.
Our finding that loading cells with non-hydrolyzable alkyl-phospholipid
analogs inhibits proliferation suggests that the regulated deacylation
of membrane phospholipids is critical to normal cell proliferation.
These data also demonstrate that alternative pathways of degradation
will not substitute. The doubling of membrane phospholipid mass is a
periodic, cell cycle-regulated event that occurs during S phase (19).
Phospholipid turnover is high during the G1 phase, and the
accumulation of phospholipid in S phase correlates with the abrupt
cessation of phospholipid turnover at the G1/S boundary (19). Thus, the down-regulation of the excess phospholipid response is
likely instrumental in permitting the accumulation of phospholipid mass
in preparation for cell division.
The inhibition of the excess phospholipid response by BEL implicates a
calcium-independent (group VI) phospholipase A2 as an
initiator of the cellular response to excess phospholipid. The
identification of GPC and GPE as the primary products generated in
response to phospholipid overproduction points to a phospholipase A2 as the initiating event. There are several classes of
phospholipase A2 enzymes, and understanding their
physiological roles and regulatory properties is an area of active
investigation (29, 33, 34). BEL is an irreversible mechanism-based
inhibitor of iPLA2 and is known not to effect the
activities of other phospholipase A2 enzymes (29, 35, 36).
Thus, BEL inhibition of PtdCho deacylation points directly to
iPLA2 as a phospholipase that is critical to the response
of cells to excess phospholipid. BEL is not a completely selective
iPLA2 inhibitor since it also blocks the activity of Mg2+-dependent phosphatidic acid phosphatase
(37), some proteases (38), and perhaps other enzymes as well. However,
the degradation of PtdCho to GPC in response to the acylation of
exogenous LPC does not involve phosphatidic acid phosphatase. There are
a number of lysophospholipases in cells (39), and one or more of these enzymes may participate in the removal of the second fatty acid to form
GPC. BEL also decreased the formation of GPC in cells that were not
stimulated to overproduce phospholipids, indicating that
iPLA2-dependent phospholipid breakdown via GPC
formation occurs under normal physiological circumstances. However, BEL did not eliminate GPC formation in cells, indicating that there is
another mechanism or process that impacts on the steady-state levels of
this metabolite. Cellular GPC is an organic osmolyte that is modulated
in response to changes in medium osmolarity and the activity of a
zinc-dependent phosphodiesterase (40-42). Thus, the
portion of GPC metabolism that is not affected by BEL may be part of an
osmotic regulatory system that operates independently of phospholipid homeostasis.
The ubiquitous occurrence of iPLA2 and the absence of
regulation by phosphorylation or calcium are consistent with a
fundamental, housekeeping role for this class of enzymes in cell
physiology as opposed to an involvement in cell signaling and
eicosanoid release (29). The iPLA2 enzymes are universally
expressed, and iPLA2s are being purified and characterized
from an increasing number of sources (43-52). The participation of
iPLA2 in the remodeling of membrane phospholipids is the
most clearly defined function for this enzyme (29). This process is an
ongoing cycle of phospholipid deacylation followed by the reacylation
of the resulting lysophospholipid that generates a different
phospholipid molecular species with new biophysical properties (17,
53). This remodeling cycle is a major route for the incorporation of
arachidonic acid into phospholipids, and the inhibition of this process
by BEL (17) and iPLA2 antisense RNA (54) support a role for
iPLA2 in the deacylation component of this cycle.
Experiments with BEL have also implicated iPLA2 as the
phospholipase responsible for the stimulated release of fatty acids
from glycerophospholipids during Fas-mediated apoptosis (55). Our
results point to a determinant role for iPLA2 in the
deacylation reactions involved in phospholipid homeostasis and reveal
another manifestation of the housekeeping functions of
iPLA2. One of the major challenges for the future is
understanding how iPLA2 activity is modulated. The
potential role of oligomerization of iPLA2 (43, 48) and the
regulated expression of splice variants that act as dominant negative
inhibitors (50) are two possible mechanisms that warrant evaluation.
The second step in GPC formation is the action of a lysophospholipase
on the LPC generated by iPLA2. The lysophospholipases are
grouped into two categories based on their molecular size, and
representatives of both classes have been cloned. The low molecular
mass (24.7 kDa) lysophospholipase is similar to bacterial esterases in
that it has a GXSXG motif associated with the
active sites of serine esterases (39, 56). The high molecular mass (60 kDa) lysophospholipases are also widely distributed in tissues. Like the iPLA2, the 60-kDa lysophospholipase cloned
from rat liver possesses ankyrin repeats (57). The presence of these
protein-protein interaction domains on these two phospholipases
suggests that their similar biochemical properties may
collaborate in membrane phospholipid homeostasis.