From the Department of Molecular Physiology and
Biophysics, the
Howard Hughes Medical Institute, and the
Departments of § Biochemistry and ¶ Medicine,
Vanderbilt University School of Medicine, Nashville,
Tennessee 37232-0295
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ABSTRACT |
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Platelet-derived growth factor (PDGF) activates
phospholipase D (PLD) in mouse embryo fibroblasts (MEFs). In order to
investigate a role for phospholipase C-1 (PLC-
1), we used
targeted disruption of the Plcg1 gene in the mouse to
develop Plcg1+/+ and
Plcg1
/
cell lines.
Plcg1+/+ MEFs treated with PDGF showed a time-
and dose-dependent increase in the production of total
inositol phosphates that was substantially reduced in
Plcg1
/
cells.
Plcg1+/+ cells also showed a PDGF-induced
increase in PLD activity that had a similar dose dependence to the PLC
response but was down-regulated after 15 min. Phospholipase D activity,
however, was markedly reduced in Plcg1
/
cells. The PDGF-induced inositol phosphate formation and the PLD
activity that remained in the Plcg1
/
cells
could be attributed to the presence of phospholipase C-
2 (PLC-
2)
in the Plcg1
/
cells. The PLC-
2 expressed
in the Plcg1
/
cells was phosphorylated on
tyrosine in response to PDGF treatment, and a small but significant
fraction of the Plcg1
/
cells showed
Ca2+ mobilization in response to PDGF, suggesting that the
PLC-
2 expressed in the Plcg1
/
cells was
activated in response to PDGF. The inhibition of PDGF-induced phospholipid hydrolysis in Plcg1
/
cells was
not due to differences in the level of PDGF receptor or in the ability
of PDGF to cause autophosphorylation of the receptor. Upon treatment of
the Plcg1
/
cells with oleoylacetylglycerol
and the Ca2+ ionophore ionomycin to mimic the effect of
PLC-
1, PLD activity was restored. The targeted disruption of
Plcg1 did not result in universal changes in the cell
signaling pathways of Plcg1
/
cells, because
the phosphorylation of mitogen-activated protein kinase was similar in
Plcg1+/+ and Plcg1
/
cells. Because increased plasma membrane ruffles occurred in both
Plcg1+/+ and Plcg1
/
cells following PDGF treatment, it is possible neither PLC nor PLD are
necessary for this growth factor response. In summary, these data
indicate that PLC-
is required for growth factor-induced activation
of PLD in MEFs.
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INTRODUCTION |
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Platelet-derived growth factor
(PDGF)1 binds receptors
(PDGFRs) located on the cell surface. Upon ligand binding, these
receptors undergo dimerization and activation of the intrinsic tyrosine kinase, which results in autophosphorylation of the receptor (1-3). The phosphorylated tyrosine residues on the PDGFR act as docking sites
for the SH2 domain of cytosolic signaling molecules, including phosphoinositide phospholipase C- (PLC-
), the phosphotyrosine phosphatase syp, the regulatory subunit of
phosphatidylinositol 3-kinase, the Ras GTPase activating protein, the
cytosolic tyrosine kinase Src, and adapter proteins, such as Shc, Grb2,
and Nck (4, 5). Upon binding to phosphorylated Tyr-1021, PLC-
is
phosphorylated and activated, resulting in the hydrolysis of
phosphatidylinositol 4,5-bisphosphate (PIP2) to
diacylglycerol (DAG) and inositol-1,4,5-trisphosphate (IP3). Diacylglycerol activates protein kinase C (PKC),
whereas IP3 liberates Ca2+ from stores in the
endoplasmic reticulum (6, 7).
Phospholipase D hydrolyzes phosphatidylcholine, generating
choline and phosphatidic acid (PA) (8, 9). Phosphatidic acid exerts
many effects in vitro, including the stimulation of PLC-, phosphatidylinositol-4-phosphate kinase, and protein kinases (10). In
addition, through the actions of PA phosphohydrolase and a specific
phospholipase A2, PA can be converted to DAG and the signaling molecule lysophosphatidic acid, respectively (11).
There is conflicting evidence about whether the activation of
PLC- and the subsequent activation of PKC are necessary for agonist
stimulation of PLD. Although there are many studies reporting the
involvement of PKC in the activation of PLD by agonists (12-15), there
also are reports that PKC is not involved (16-19). Furthermore, some
studies have indicated that PLD can be activated by certain agonists in
the absence of detectable PIP2 hydrolysis. For example, in
Madin-Darby canine kidney cells, studies with neomycin indicate that
activation of PLD by purinergic agonists is independent of PLC-
activity (20). In certain fibroblasts, PLD activation by epidermal
growth factor (EGF) has been reported to occur in the absence of
measurable PIP2 breakdown (21). However, in Swiss 3T3
fibroblasts and TRMP cells, activation of PLC-
1 is necessary for
stimulation of PLD activity by PDGF (22). Previously, we used
homologous recombination to selectively disrupt the Plcg1 gene encoding PLC-
1 in mice (23). Although this mutation was lethal,
immortal mouse embryo fibroblast (MEF) cell lines were produced from
Plcg1+/+ and Plcg1
/
embryos. We have now used these cells to study PDGF-induced PLD activity. The results indicate that PLC-
1 activity is required for
PDGF-induced PLD activation.
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EXPERIMENTAL PROCEDURES |
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Materials--
Dulbecco's modified Eagle's medium (DMEM) with
L-glutamine and high glucose and Earle's modified Eagle's
medium were purchased from Life Technologies, Inc. EGF was from G. Carpenter, PDGF B/B (human recombinant) was from Boehringer Mannheim,
and ionomycin was from Sigma. 1-Oleoyl-2-acetyl-sn-glycerol
(OAG) and phosphatidylbutanol (PtdBut) standards were from Avanti Polar
Lipids. [9,10-3H]Myristate and
myo-[2-3H]inositol were purchased from NEN
Life Science Products. Antibodies to the PDGF type B receptor (rabbit
anti-human) and phosphotyrosine (P-Tyr) (4G10) were from Upstate
Biotechnology, and antibodies to p44/42 mitogen-activated protein (MAP)
kinase and phospho-specific p44/42 MAP kinase (Thr-202/Tyr-204) were
purchased from New England Biolabs. The antibody to PLC-2 was
purchased from Santa Cruz. SDS-polyacrylamide gels were from NOVEX.
BODIPY 558/568 phalloidin and fluo-3 AM were from Molecular Probes.
Cell Culture--
MEFs were prepared from embryonic day 9.5 embryos with (Plcg1/
) and without
(Plcg1+/+) targeted disruption of the
Plcg1 gene, using the targeting vectors TVI and TVII (23).
Wild type and null cells from the same litter were established in
culture according to standard methods and maintained as immortalized
non-transformed cell lines. Plcg1 gene disruption by
targeting vector TVI disrupted exons encoding the X domain and the two
SH2 domains of PLC-
1, whereas targeting vector TVII replaces at the
genomic level exons encoding the X domain and two SH2 domains of
PLC-
1 with a LacZ sequence. Cells targeted with TVII produce a
fusion protein of the PLC-
1 N terminus and
-galactosidase (23).
The fibroblasts were passaged in DMEM containing 10% fetal bovine
serum at 37 °C in a humidified atmosphere with 5%
CO2.
Phospholipase D Assay-- Cells were plated in 60-mm tissue-culture plates. The cells were serum-starved in DMEM (Life Technologies, Inc.) containing 0.5% bovine serum albumin for 16 h prior to the start of the assay. During this time, they were labeled with 1 µCi/ml [9,10-3H]myristic acid. At the start of the experiment, the cells were washed three times with 5 ml of phosphate-buffered saline (PBS) and pre-equilibrated at 37 °C in serum-free DMEM for 1 h. For the final 10 min of preincubation, 0.3% butan-1-ol was included. At the end of the preincubation, cells were treated with the indicated agonist for 10 min or the times indicated. Incubations were terminated by removing the medium, washing on ice with 5 ml of ice-cold PBS, and adding 1.5 ml of ice-cold methanol. Cells were scraped off the plates, and the lipids were extracted and separated with methanol/chloroform/0.1 N HCl (1:1:1) according to the method of Bligh and Dyer (24). The lower phase was dried under N2, resuspended in 30 µl of chloroform/methanol (2:1), and spotted onto silica gel 60A thin layer chromatography plates (Whatman). The plates were developed in the upper phase of the solvent system of ethyl acetate/iso-octane/H2O/acetic acid (55:25:50:10) and stained with iodine. A PtdBut standard (Avanti Polar Lipids) was used to locate the bands, which were scraped into scintillation vials containing 500 µl of methanol and 7.5 ml of Ready Organic scintillation mixture (Beckman). Radioactivity incorporated into total phospholipids was measured, and the results were presented as percentage of total lipid cpm incorporated into PtdBut.
Measurement of Total Inositol Phosphates-- Inositol phosphates were measured as described by Yeo and Exton (12). Briefly, cells were plated on 6-well tissue culture plates and labeled for 16 h with 1 µCi/ml myo-[2-3H]inositol in inositol-free, serum-free DMEM supplemented with 0.5% bovine serum albumin. At the start of the experiment, the cells were preincubated for 1 h at 37 °C. During the last 10 min of preincubation, 20 mM LiCl was included. The cells were treated with the indicated growth factors for 15 min. The experiment was terminated by removing the medium, washing with 5 ml of ice-cold PBS, and adding 750 µl of ice-cold 20 mM formic acid. After 30 min of incubation on ice, the cells were neutralized with 250 µl of 50 mM NH4OH. The cells were scraped on ice, transferred to a 1.5-ml Eppendorf tube, and spun at maximum speed for 10 min at 4 °C. The supernatant (900 µl) was loaded onto Bio-Rad 10-ml columns containing 1 ml of AG1-8X resin. Inositol (Ins) and inositol phosphates (InsPx) were eluted according to Simpson et al. (25). Five hundred µl of each eluted fraction was counted in 15 ml of Ready Safe (Beckman). The results are expressed as ((InsPx)/(InsPx + Ins)) × 1000.
Immunoprecipitation of PLC-2--
Subconfluent cells on
100-mm dishes were lysed in radioimmune precipitation assay buffer (50 mM Tris, pH 7.4, 1% Nonidet P-40, 0.25% sodium
deoxycholate, 150 mM NaCl, 1 mM EGTA, 1 mM NaF, 1 mM phenylmethylsulfonyl fluoride
(PMSF), 1 mM Na3VO4, 1 mg/ml aprotinin, 1 mg/ml leupeptin, 1 mg/ml pepstatin) and precleared for
1 h with 1 µg of anti-rabbit IgG and 20 µl of protein A/G PLUS-agarose (Santa Cruz) at 4 °C with rocking. A protein assay was
performed on the precleared cell lysate, which was then diluted to 1 mg/ml in 500 µl and immunoprecipitated with 3 µg of rabbit polyclonal PLC-
2 antibody for 1 h at 4 °C. Twenty µl of
protein A/G PLUS-agarose was added overnight at 4 °C with rocking.
In the morning, the protein A/G PLUS-agarose beads were collected with
centrifugation, washed three times with 1 ml of radioimmune precipitation buffer, and resuspended in 20 µl of 2× SDS sample buffer. The samples were boiled, and the immunoprecipitated proteins were separated on a 6% SDS-PAGE gel. The proteins were transferred to
Immobilon-P, blocked in 3% dry milk in PBS, and probed with an
antibody to P-Tyr at a concentration of 1 µg/ml.
Western Blotting-- Subconfluent cells in 100-mm dishes that were serum-starved in serum-free DMEM containing 0.5% bovine serum albumin for 24 h were washed once with 5 ml of PBS and scraped directly in 300 µl of PBS containing 1 mM EDTA, 1 mM EGTA, 1 mM NaF, 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3VO4, 1 µg/ml aprotinin, 1 µg/ml leupeptin, and 1 µg/ml pepstatin. A protein assay using the BCA method (Pierce) was performed on the samples, and cells on a duplicate 100-mm dish were counted using a hemocytometer. Proteins were separated by SDS-PAGE on a 6% (PDGFR and phosphotyrosine) or a 4-20% gradient gel (MAP kinase and phospho-MAP kinase), transferred to Immobilon-P, blocked in 5% dry milk (MAP kinase) or 3% dry milk/PBS (P-Tyr), and probed with the antibody to PDGFR, P-Tyr, p44/42 MAP kinase, or phospho p44/42 MAP kinase.
Mobilization of Intracellular
Ca2+--
Plcg1+/+ and
Plcg1/
cells were plated on coverslips and
grown to 80-90% confluence before the addition of Earle's modified
Eagle's medium plus 0.5% fetal calf serum overnight. The coverslips
were then washed twice with wash buffer (10 mM Hepes, pH
7.4, 140 mM NaCl, 5 mM KCl, 1 mM
MgCl2, 0.55 mM glucose), and the cells were loaded with 1 mM fluo-3 AM for 45 min at room temperature.
The coverslips were washed and placed into the microscope chamber, followed by the addition of 1 ml of wash buffer containing an additional 1 mM CaCl2. This was followed by the
addition of 25 ng/ml PDGF or 1% fresh fetal calf serum. The number of
cells emitting fluorescence at a wavelength of 488 nm were counted
using a Zeiss Axiovert 135 confocal microscope.
PDGF-induced Membrane
Ruffling--
Plcg1+/+ and
Plcg1/
cells were plated on glass coverslips
in 6-well plates. Subconfluent cells were serum-starved for 24 h,
at which time they were treated with 5 ng/ml PDGF or serum-free DMEM containing 0.5% fatty acid-free bovine serum albumin for 10 min at
37 °C. The medium containing agonist was removed, and the cells were
washed once with 1 ml of PBS at room temperature and fixed by adding 1 ml 3.7% formaldehyde in PBS to each well for 10 min with rocking. The
fixed cells were washed twice with 5 ml of PBS at room temperature and
permeabilized with 1 ml 0.2% Triton X-100 in PBS/well for 5 min with
rocking. The cells were washed twice with 1 ml of PBS, and the actin
stained with 3.75 units/ml BODIPY 558/568 phalloidin for 20 min in the
dark with rocking. Continuing in the dark, the cells were washed three
times with 1 ml of PBS, and the coverslips removed from the dish and
allowed to dry in air overnight. The coverslips were sealed on a slide
using clear nail polish, and fluorescence was observed with a ×40
objective using a Leica DMRB microscope.
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RESULTS |
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Growth Factor-induced PLD Activity--
In order to test for a
role for PLC-1 in growth factor-induced PLD activity, we treated
TVI- and TVII-targeted Plcg1+/+ and
Plcg1
/
MEFs with EGF or PDGF. TVI-targeted
Plcg1+/+ MEFs have approximately 2.8 × 104 EGF receptors/cell, whereas the TVI
Plcg1
/
MEFs have approximately 5.2 × 104 EGF receptors/cell (26). In TVI-targeted
Plcg1+/+ MEFs, 3 nM EGF caused a
2-fold increase in PLD activity but no increase in TVI-targeted
Plcg1
/
cells. In TVII-targeted
Plcg1+/+ and Plcg1
/
MEFs, EGF did not induce significant PLD activity (Fig.
1A). In TVI-targeted
Plcg1+/+ MEFs, 50 ng/ml PDGF elicited a 3-fold
increase in PLD activity that was inhibited in TVI-targeted
Plcg1
/
MEFs. Most strikingly, PDGF produced
a robust PLD response in TVII-targeted Plcg1+/+
MEFs that was greatly inhibited in TVII-targeted
Plcg1
/
MEFs (Fig. 1B). Thus, both
the TVI- and TVII-targeted Plcg1+/+ MEFs showed
PDGF-induced PLD activity that was inhibited in
Plcg1
/
MEFs. Because the response to PDGF
was so much greater in the TVII-targeted
Plcg1+/+ MEFs, we selected this cell line to
further investigate the role of PLC-
1 in the PDGF-induced PLD
response.
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InsPx Formation and PtdBut Formation in
Plcg1+/+ and Plcg1/
MEFs--
In order to
investigate the extent to which disruption of PLC-
1 eliminated
PDGF-induced inositol phosphate formation, we measured total
InsPx in the Plcg1+/+ and
Plcg1
/
MEFs in response to PDGF.
Platelet-derived growth factor increased the production of
InsPx in Plcg1+/+ cells in a dose-
and time-dependent fashion (Fig.
2). In Plcg1
/
MEFs, PDGF caused a smaller increase in InsPx formation. As
shown in Fig. 3, PDGF treatment of
Plcg1+/+ MEFs resulted in a dose- and
time-dependent increase in PtdBut, an unambiguous marker of
PLD activity when cells are treated with agonist in the presence of
butan-1-ol (27). However, PDGF-induced PLD activity was inhibited in
Plcg1
/
MEFs. Whereas the dose-response
curves for InsPx were similar to those for PtdBut (Fig.
3A; cf. Fig. 2A), the time course for PtdBut indicated no further production after 15 min despite further increases in InsPx (Fig. 3B; cf. Fig.
2B) These results are consistent with PLC-
1 acting
upstream of PLD in growth factor-induced activation, but they do not
explain the cessation of PLD activation at 15 min.
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Expression of PLC-2 and Ca2+ Mobilization in
Plcg1
/
Cells--
In Plcg1
/
cells, PDGF caused a small increase in both inositol phosphate
production (Fig. 2) and phospholipase D activity (Fig. 3). Furthermore,
a small fraction (13.2%) of the Plcg1
/
cells showed intracellular Ca2+ mobilization upon treatment
with 25 ng/ml PDGF, whereas a large majority (94%) of the
Plcg1
/
cells showed intracellular
Ca2+ mobilization upon treatment with 1% fetal calf serum.
It has been reported that cells of hematopoietic origin express
PLC-
2, a PLC isoform that is closely related to PLC-
1 (28).
Treatment of Rat 2 cells overexpressing PLC-
2 with PDGF causes an
increase in the tyrosine phosphorylation and activation of PLC-
2
(29). Thus, we investigated whether Plcg1
/
cells expressed PLC-
2. Western blot analysis of
Plcg1+/+ and Plcg1
/
cells with a PLC-
2 antibody showed a large expression of PLC-
2 in
Plcg1
/
cells when compared with
Plcg1+/+ cells (Fig.
4A). Furthermore, upon
treatment of Plcg1
/
cells with 25 ng/ml
PDGF, PLC-
2 was phosphorylated on tyrosine and co-immunoprecipitated
with the PDGF receptor. In Plcg1+/+ cells
treated with PDGF, there was no apparent tyrosine phosphorylation of
PLC-
2 and very little co-immunoprecipitation with the PDGF receptor
(Fig. 4B). These data suggest that the
Plcg1
/
cells may compensate for the
disruption of Plcg1 by up-regulating PLC-
2 and that the
activation of PLC-
2 by PDGF accounts for the small increase in
inositol phosphate formation, PLD activation, and the Ca2+
mobilization seen in the Plcg1
/
cells.
|
PDGF Receptor Level and Autophosphorylation--
To establish that
the decrease in PDGF-induced production of InsPx and PtdBut
in Plcg1/
cells was not due to a decrease in
the number of PDGF receptors, Western blotting of the receptors was
performed. This showed no difference in the level of PDGF receptors in
Plcg1
/
and Plcg1+/+
cells (data not shown). We also investigated the ability of the PDGFR
to be autophosphorylated upon treatment of the
Plcg1+/+ and Plcg1
/
cells with 25 ng/ml PDGF. As shown in Fig.
5, phosphorylation of the PDGFR on
tyrosine residues was rapid and sustained for up to 1 h in both
cell types, although the effect occurred more rapidly and declined
faster in the Plcg1
/
cells. Thus,
differences in the level and autophosphorylation of the PDGFR do not
account for the decrease in PDGF-induced phospholipase responses in the
Plcg1
/
MEFs.
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Reconstitution of PDGF Induced PLD Activation in
Plcg1/
Cells--
In cells treated with PDGF, PLC-
1
is recruited via its SH2 domain to the receptor, where it is activated
(6). The activated PLC-
1 hydrolyzes PIP2, resulting in
the formation of DAG and IP3. Diacylglycerol activates most
isozymes of PKC, whereas IP3 promotes the release of
intracellular Ca2+. It has been proposed that these two
second messengers, acting alone or in combination, mediate growth
factor activation of PLD (30). If this is true, then replacing these
PLC-
1 products should reconstitute PDGF-induced PLD activation in
Plcg1
/
MEFs. As shown in Fig.
6, the addition of the calcium ionophore ionomycin (5 µM) plus the cell-permeable DAG analog OAG
(40 µM) resulted in an equal PLD response in the
Plcg1+/+ and Plcg1
/
cells. The response in the Plcg1
/
cells to
OAG plus ionomycin was similar to that induced by 50 ng/ml PDGF in
Plcg1+/+ cells. These data show that the
deficient PDGF-induced PLD response in
Plcg1
/
cells can be entirely restored by
addition of agents that mimic the activation of PLC-
1.
|
PDGF-Induced Activation of MAP Kinase--
Treatment of many cell
types with growth factors results in activation of the MAP kinase
pathway. This activation occurs through the activation of Ras, followed
by the activation of MEK kinase, MEK, and finally, MAP kinase (31).
There is evidence that phospholipase C-1 is not involved in the
growth factor-induced activation of MAP kinase (26, 32). In order to
confirm that the deletion of Plcg1 by targeted gene
disruption did not result in secondary changes to other PDGF signaling
pathways in the MEFs derived from the targeted embryos, we assessed the
activation of MAP kinase in Plcg1+/+ and
Plcg1
/
cells using an antibody that
recognizes the phosphorylated form of MAP kinase. We showed that PDGF
treatment of Plcg1+/+ and
Plcg1
/
cells resulted in a rapid and
transient increase in MAP kinase phosphorylation that was similar
between the two cells (Fig. 7). This
result suggests that only pathways that require the activation of
PLC-
1 are inhibited in Plcg1
/
MEFs.
|
Requirement of PLD for PDGF-induced Membrane Ruffling--
Two
recent reports suggest that the stimulation of actin stress fiber
formation by lysophosphatidic acid or -thrombin is mediated by the
activation of PLD (33, 34). Platelet-derived growth factor has been
shown to induce the formation of polymerized actin at the plasma
membrane in Swiss 3T3 cells, forming membrane ruffles (35). Because PLD
activation by PDGF is inhibited in Plcg
/
cells, we used these cells to test whether PLD activation was necessary
for PDGF-induced membrane ruffles. Fig. 8
shows that PDGF induces membrane ruffles in
Plcg1+/+ cells, and this is not inhibited in
Plcg1
/
MEFs. These data suggest that PLD
activation may not be required for the increase of polymerized actin
localized in ruffles at the plasma membrane induced by PDGF.
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DISCUSSION |
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Growth factors stimulate the PLD-induced hydrolysis of
phosphatidylcholine (PC) to PA and choline in a variety of cell types (30). The exact pathway by which this occurs is not clear. A number of
mechanisms of activation have been proposed involving protein tyrosine
kinases, PKC, Ca2+, and GTP-binding proteins (36). In an
elegant study seeking to define a role for PLC-1 in PDGF-induced PLD
activation, Yeo et al. (22) measured PLD activity in TRMP
cells (a kidney epithelial cell line) expressing wild type PDGF
receptors or various tyrosine mutated PDGF receptors. They reported
that PDGF had no effect on PLD activity in PDGFR kinase-deficient TRMP
cells, but the PDGF-induced PLD activity was restored in cells
containing a mutant PDGFR that was able to bind PLC-
1 but not other
signaling proteins. Furthermore, they showed that a mutant PDGFR that
could not activate PLC-
1 was unable to activate PLD. These data
suggest that PLC-
1 is necessary and sufficient for PDGF-induced PLD
activity. However, the experiments were conducted with a cell line that
normally lacked the PDGFR (37). Further evidence for a role of PLC-
in the activation of PLD came from a study in which PLC-
1 was overexpressed in NIH3T3 cells. Lee et al. (38) found that
PDGF-induced PLD activity was directly related to the level of PLC-
1
expressed in the cells, and that down-regulation of PKC by PMA
pretreatment completely blocked PLD activation. These data again
suggest PLD lies downstream of PLC-
1 and PKC.
On the other hand, there are data that show agonist-induced PC hydrolysis or PLD activation in the absence of detectable PIP2 breakdown (16, 18, 20). Cook and Wakelam (21) showed EGF stimulation of PLD activity in Swiss 3T3 cells in the absence of measurable PIP2 hydrolysis and in the presence of a PKC inhibitor, although it was later found that EGF induced a small increase in InsPx in these cells and that a PKC inhibitor did decrease the PLD response (12). When these reports are coupled with the frequent finding that PKC inhibitors produce only partial inhibition of the actions of growth factors and other agonists on PLD (10, 30), questions remain about the extent of the contribution of PIP2 hydrolysis and PKC to the regulation of PLD.
The data presented strongly suggest that PLC-1 is required for PLD
activation by PDGF. The PLD response of TVI-targeted
Plcg1+/+ cells to EGF and PDGF is small, and the
response is inhibited in TVI-targeted Plcg1
/
cells (Fig. 1). Furthermore, the PDGF-induced PLD response of TVII-targeted Plcg1+/+ cells is robust and is
greatly inhibited in TVII-targeted Plcg1
/
cells (Fig. 1). A small but reproducible, increase in PLD activity with
EGF and a contrasting robust PLD response to PDGF have been seen in a
variety of cell types, including Rat1 and Swiss 3T3 fibroblasts (12,
39). The present study thus adds to earlier data reporting differences
in the signal transduction pathways for the two growth factors
(40-42).
Phospholipase C is responsible for the hydrolysis of PIP2
to IP3 and DAG (6). In Plcg1+/+
cells, PDGF elicits an increase in the production of total inositol phosphates in a dose- and time-dependent manner (Fig. 2).
In Plcg1/
cells, PDGF caused a small
increase in inositol phosphate production (Fig. 2). Our data suggest
that the small increase in inositol phosphate production in
Plcg1
/
cells is due to the expression of
PLC-
2 in the Plcg1
/
cells (Fig.
4A). Treatment of Plcg1
/
cells
with PDGF resulted in tyrosine phosphorylation of PLC-
2 (Fig.
4B). PDGF treatment of rat-2 cells overexpressing PLC-
2 increases the tyrosine phosphorylation and the in vivo
activity of PLC-
2 (29). Moreover, treatment of
Plcg1
/
cells with PDGF resulted in the
mobilization of intracellular Ca2+ in a small population of
the cells. Over-expression of PLC-
2 in NIH3T3 cells also enhances
PDGF-induced mobilization of intracellular Ca2+ (43). Thus,
our data suggest that the disruption of Plcg1 resulted in a
compensatory up-regulation of PLC-
2 and that this isoform, which is
closely related to PLC-
1, is responsible for the increase in
InsPx formation (Fig. 2), PLD activity (Fig. 3), and
intracellular Ca2+ mobilization seen in
Plcg1
/
cells upon treatment with PDGF.
If PLC-1 acts upstream of PLD in the PDGF-induced PLD-activation
pathway, then PLD activity should be inhibited in
Plcg1
/
cells. Furthermore, there should be a
correlation between PLC and PLD activities. Treatment of
Plcg1+/+ cells with PDGF results in a
dose-dependent increase in PtdBut formation that mirrors
the dose-response curve for PDGF-induced InsPx production
in Plcg1+/+ (Fig. 3A; cf.
Fig. 2A). The PLD response to PDGF treatment in Plcg1
/
cells is inhibited in parallel with
the decrease in the PLC response (Fig. 3A; cf.
Fig. 2A). However, the PDGF-induced PLD response reached a
maximum at 15 min in the Plcg1+/+ or the
Plcg1
/
cells, at which time the production
of PtdBut ceased (Fig. 3B). This is in contrast to the
PDGF-induced InsPx production, which was still increasing
at 60 min. This same pattern of phosphatidylalcohol and inositol
phosphate production was reported in NIH3T3
-1 cells, which
overexpress PLC-
1 and in which phosphatidylethanol production reached a maximum at 10 min in response to PDGF, whereas
InsPx production was still increasing at 30 min (44).
Exploration of the reasons for the cessation of PtdBut formation is
outside the scope of the present study, but it is possible that
activation of PLC and the consequent activation of PKC and mobilization
of Ca2+ could have an initial stimulatory effect on PLD
followed by an inhibitory action, due perhaps to phosphorylation of PLD
or an inhibitory protein. Phosphorylation of PLD by PKC has recently been reported to inhibit its activity (45).
The inhibition of PDGF-induced PLD activity in
Plcg1/
cannot be attributed to a decreased
level of PDGF receptors in the Plcg1
/
cells
(data not shown) or to a defect in PDGF-induced autophosphorylation (Fig. 5). In fact, the autophosphorylation of the receptor occurred more rapidly in the Plcg1
/
cells as compared
with the Plcg1+/+ cells, reaching a maximum
level at 5 min and decreasing toward basal level by 60 min, but this
difference in autophosphorylation cannot account for the decreased
PDGF-induced PLD activity seen in the Plcg1
/
cells. Another possible explanation is that PKC is deficient in the
Plcg1
/
cells. However, this does not seem to
be the case because treatment of Plcg1+/+ and
Plcg1
/
cells with phorbol ester, an
activator of PKC, results in a similar dose-dependent
activation of PLD (data not shown).
In cells treated with growth factors, activated PLC-1 hydrolyzes
PIP2 to form IP3 and DAG, resulting in an
increase in intracellular Ca2+ and the activation of PKC.
Treatment of Plcg1
/
cells with the
Ca2+ ionophore ionomycin and the cell-permeable DAG
analogue OAG resulted in a PLD response that was similar to that in
Plcg1+/+ cells and slightly greater than the PLD
response induced by PDGF (Fig. 6). Thus, the addition of PLC-
1
activation products to the Plcg1
/
cells
reconstituted the PDGF-induced PLD response in the
Plcg1
/
cells to the level seen in the wild
type cells. These data suggest that the PLC-
1 activation products
are sufficient to completely restore the PDGF-induced PLD response lost
in the Plcg1
/
upon disruption of PLC-
1 in
these cells. Furthermore, these results and those with phorbol ester
prove that PLD is not deficient in the
Plcg1
/
cells.
Growth factor treatment of cells results in a mitogenic response that
is mediated by the MAP kinase. The growth factor-induced activation of
MAP kinase involves the sequential activation of Ras, MEK kinase, and
MEK (31). In a 3T3 cell line derivative, NR6 cells, EGF-stimulated MAP
kinase activity was not affected by the inhibition of PLC with U73122
(32), and data from Ji et al. (26) showed that EGF-induced
activation of MAP kinase in the TVI-targeted
Plcg1+/+ and Plcg1/
cells was similar. Thus, these findings indicate that PLC-
1 is not
involved in the phosphorylation and activation of MAP kinase by growth
factors. We also observed PDGF-induced phosphorylation of MAP kinase in
Plcg1+/+ and Plcg1
/
cells (Fig. 7), indicating that the targeted gene disruption of
PLC-
1 did not result in global changes to PDGF-signaling
pathways.
Data from two recent reports suggest a role for PA in the polymerization of actin stress fibers (33, 34). Actin stress fibers are a major component of the cytoskeleton in fibroblasts, where actin filaments can exist in three types of structures, including actin stress fibers, the cortical actin network, and cell surface protrusions, such as membrane ruffles and filopodia (35). Ha and Exton (34) reported that treatment of IIC9 fibroblasts with thrombin, PLD from Streptomyces chromofuscus, or exogenous PA resulted in actin stress fiber formation. In porcine aortic endothelial cells, lysophosphatidic acid treatment activated PLD, resulting in the formation of PA, in the apparent absence of the formation of other lipid second messengers (33). Lysophosphatidic acid, like exogenously added PA, also stimulated the formation of actin stress fibers (33). Although these observations generally support a role for PLD in stress fiber formation, it is possible that signals evoked by the exogenous PLD and PA are different from those elicited by activation of endogenous PLD (46). For example, they could generate lysophosphatidic acid, which could induce actin polymerization by a different mechanism.
In Swiss 3T3 cells, PDGF has been shown to induce the formation of
membrane ruffles (35), and we therefore utilized the Plcg1/
cells to examine the role of PLD in
this effect. In Plcg1
/
cells, 5 ng/ml PDGF
induced membrane ruffles similar to those induced in
Plcg1+/+ cells (Fig. 8), even though activation
of PLD was significantly inhibited (Fig. 2B). Thus, it
appears that PLD and PLC activity may not be necessary for the
PDGF-induced formation of membrane ruffles. It seems unlikely that the
small level of PLC-
2, inositol phosphate formation, and
Ca2+ mobilization would be sufficient to provoke maximal
ruffling response.
In summary, the present data suggest that in mouse embryo fibroblasts,
PLC-1 activation is necessary for the PDGF-induced activation of
PLD. However, caution should be exercised in extrapolating the findings
to other agonists or cell types. We are currently investigating a role
for PLC-
1 in the activation of PLD by various other agonists,
including those that activate heterotrimeric G-proteins.
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FOOTNOTES |
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* Supported in part by Grants T32KD07061-24 (to J. A. H.) and R01CA75195 (to G. C.) from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
** Investigator of the Howard Hughes Medical Institute. To whom correspondence should be addressed. Tel.: 615-322-6494; Fax: 615-322-4381; E-mail: john.exton{at}mcmail.vanderbilt.edu.
The abbreviations used are:
PDGF, platelet-derived growth factor; PDGFR, PDGF receptor; PLD, phospholipase D; MEF, mouse embryo fibroblast; PLC-1, phospholipase
C; PIP2, phosphatidylinositol 4,5-bisphosphateDAG, diacylglycerolIP3, inositol-1,4,5-trisphosphatePKC, protein kinase CPA, phosphatidic acidDMEM, Dulbecco's modified
Eagle's mediumOAG, 1-oleoyl-2-acetyl-sn-glycerolPBS, phosphate-buffered salineIns, inositolInsPx, inositol
phosphatesP-Tyr, phosphotyrosineEGF, epidermal growth factorMAP
kinase, mitogen-activated protein kinasePtdBut, phosphatidylbutanol.
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