1 Division of Pulmonary and Critical Care Medicine, Johns Hopkins University School of Medicine, Baltimore, Maryland 21224; and 2 Department of Pharmacological Sciences, Health Science Center, State University of New York at Stony Brook, Stony Brook, New York 11794
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ABSTRACT |
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We previously demonstrated that diperoxovanadate (DPV), a synthetic peroxovanadium compound and cell-permeable oxidant that acts as a protein tyrosine phosphatase inhibitor and insulinomimetic, increased phospholipase D (PLD) activation in endothelial cells (ECs). In this report, the regulation of DPV-induced PLD activation by mitogen-activated protein kinases (MAPKs) was investigated. DPV activated extracellular signal-regulated kinase, c-Jun NH2-terminal kinase (JNK), and p38 MAPK in a dose- and time-dependent fashion. Treatment of ECs with p38 MAPK inhibitors SB-203580 and SB-202190 or transient transfection with a p38 dominant negative mutant mitigated the PLD activation by DPV but not by phorbol ester. SB-202190 blocked DPV-mediated p38 MAPK activity as determined by activated transcription factor-2 phosphorylation. Immunoprecipitation of PLD from EC lysates with PLD1 and PLD2 antibodies revealed both PLD isoforms associated with p38 MAPK. Similarly, PLD1 and PLD2 were detected in p38 immunoprecipitates from control and DPV-challenged ECs. Binding assays demonstrated interaction of glutathione S-transferase-p38 fusion protein with PLD1 and PLD2. Both PLD1 and PLD2 were phosphorylated by p38 MAPK in vitro, and DPV increased phosphorylation of PLD1 and PLD2 in vivo. However, phosphorylation of PLD by p38 failed to affect PLD activity in vitro. These results provide evidence for p38 MAPK-mediated regulation of PLD in ECs.
reactive oxygen species; p38 mitogen-activated protein kinase; vascular endothelium; phospholipase D; phosphorylation; peroxovanadate
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INTRODUCTION |
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PEROXOVANADIUM COMPOUNDS such as pervanadate, prepared by reacting H2O2 and vanadate, are potent modulators of signal transduction pathways in mammalian cells and are useful agents to the study of protein phosphorylation/dephosphorylation in cell proliferation, differentiation, and apoptosis. For example, pervanadate is an insulin mimetic that modulates glucose transport, glycogen metabolism, and lipolysis and activates the insulin receptor tyrosine kinase (10, 31, 45, 50). Emerging evidence suggests that pervanadate promotes insulin receptor tyrosine phosphorylation by inhibiting phosphotyrosine phosphatase associated with the insulin receptor (49). Furthermore, pervanadate activates signal transducer and activator of transcription (STAT) proteins (15), mitogen-activated protein kinases (MAPKs; see Refs. 5, 36, 55), myosin light chain kinase (14), and focal adhesion kinase (11, 14), elevates intracellular Ca2+ (36), and enhances hydrolysis of phospholipids (3, 20, 53). Recently, we have shown that diperoxovanadate (DPV), a potent inhibitor of protein tyrosine phosphatases and activator of tyrosine kinases (21), stimulates extracellular signal-regulated kinases (ERKs), caveolin, and focal adhesion kinase (14, 36, 51) in endothelial cells (ECs). In addition, DPV activates phospholipases A2, C, and D (PLD) as evidenced by enhanced generation of arachidonic acid, diacylglycerol, and phosphatidic acid (PA), respectively (30, 36). Regulation of PLD activation by a wide variety of stimuli is complex and involves changes in intracellular Ca2+, protein kinase C (PKC), heterotrimeric G proteins, small molecular weight G proteins, intracellular thiols, and tyrosine kinase/protein tyrosine phosphatases (7, 12, 13, 34, 39, 40, 44), suggesting multiple pathways of PLD activation (9, 38) or perhaps PLD isoform-specific regulation (16, 19, 46).
Activation of PLD results in the generation of PA, which is subsequently metabolized to lysoPA by phospholipase A2/A1 (2). Both PA and lysoPA have been implicated in cellular processes such as proliferation, differentiation, protein trafficking, and secretion (9, 16). The physiological relevance of PLD activation and role of PA/lysoPA in vascular function is unclear. We have previously demonstrated that exogenous PA but not lysoPA enhanced endothelial monolayer permeability to albumin and altered levels of intracellular free calcium (8). Furthermore, the effects of ectopic PA on endothelial barrier dysfunction were mimicked by PA confined to neutrophil plasma membranes. In porcine aortic ECs, it was shown that lysoPA-mediated stimulation of actin stress fiber formation was dependent on PLD activation and PA generation (5). Overexpression of PLD1 in ECs potentiated oxidant-induced permeability to albumin, suggesting a potential role for PLD1 in barrier dysfunction (Natarajan, unpublished results). These data indicate that PA generated by the PLD signaling pathway in response to oxidative stress disrupts integrity of the vascular endothelium, an early event in the pathophysiology of lung injury.
In view of our earlier observation that DPV activated MAPKs, e.g., ERK1/2 in ECs (36, 41), and ectopic PA altered barrier function of the endothelium (8), we investigated the role of MAPKs in regulating DPV-induced PLD activation. We found that p38 MAPK activation regulates DPV-induced PLD stimulation in ECs. We also show that PLD1 and PLD2 are phosphorylated by p38 MAPK in vitro and in vivo and that DPV treatment enhances PLD1 and PLD2 activities associated with p38 MAPK immunoprecipitates. Furthermore, PLD1 and PLD2 immunoprecipitates from DPV-treated ECs exhibited p38 MAPK activity and were associated with p38 MAPK. Also, association between PLD1/PLD2 and p38 MAPK occurred in vitro with glutathione S-transferase (GST) fusion protein. These results, for the first time, provide evidence for p38 MAPK-dependent regulation of PLD via direct association in basal and DPV-stimulated ECs.
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METHODS |
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Materials.
Bovine pulmonary artery ECs (CCL-209, passage 16) were
obtained from the American Type Culture Collection (Manassas, VA). MEM,
nonessential amino acids, trypsin, FBS,
12-O-tetradecanoylphorbol 13-acetate (TPA),
penicillin-streptomycin, DMEM-phosphate-free modified medium, and DMSO
were purchased from Sigma Chemical (St. Louis, MO). Phosphatidylbutanol
(PBt), dipalmitoylphosphatidylcholine (DPPC), dioleoyl
phosphatidylethanolamine (DOPE), and phosphatidylinositol 4,5-bisphosphate (PIP2) were from Avanti Polar Lipids
(Alabaster, AL). [32P]orthophosphate
(carrier-free), [3H]myristic acid (sp. act. 64.7 Ci/mmol), and [-32P]ATP (sp. act. 5 Ci/mmol in Tris
buffer) were obtained from New England Nuclear (Wilmington, DE).
Phosphatidylcholine and
L-
-[choline-methyl-3H]dipalmitoyl
(sp. act. 30-60 Ci/mmol) were purchased from American Radiolabeled
Chemicals (St. Louis, MO). SB-202190, SB-203580, SB-202474, and
PD-98059 were obtained from Calbiochem (San Diego, CA). EC growth
factor, affinity-purified anti-phosphotyrosine antibody (4G10), SAPK
2a/p38
-active (recombinant enzyme expressed in Escherichia
coli), and monoclonal p38 MAPK antibodies were from Upstate
Biotechnology (Lake Placid, NY). Phospho-specific p38 and ERK1/ERK2
antibodies, p38 MAPK assay kit, and activated transcription factor
(ATF)-2 were purchased from New England Biolabs (Beverly, MA).
Polyclonal antibodies to ERK1 and ERK2, p38 MAPK, c-Jun
NH2-terminal kinase (JNK), and protein A/G plus Sepharose were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Spodoptera frugiperda (Sf9) cells were purchased from
Invitrogen (Carlsbad, CA). The enhanced chemiluminescence (ECL) kit for
the detection of proteins by Western blots was from Amersham (Arlington Heights, IL).
Culture of ECs. Bovine pulmonary artery ECs were grown to confluence in MEM supplemented with 20% FBS, 100 U/ml penicillin and streptomycin, 5 µg/ml EC growth factor, and 1% nonessential amino acids at 37°C in a 5% CO2-95% air atmosphere as described earlier (42). Cells were grown in 35- or 100-mm dishes or T-75-cm flasks, and cells at passages between 18 and 20 were used in all of the experiments.
Baculovirus overexpression of human PLD1 and PLD2 in Sf9 cells. Sf9 cells in suspension were infected with recombinant baculovirus containing PLD1a and PLD2 as described by Hammond et al. (19). The infected cells were grown at 30°C in complete Sf9 medium (Sf900 II SFM; GIBCO BRL) containing 10% FBS for 72 h. Cells were centrifuged, and the pellet was washed in ice-cold PBS and lysed on ice with lysis buffer containing 50 mM Tris · HCl (pH 7.4), 150 mM NaCl, 1% Nonidet P-40 (NP-40), 0.25% sodium deoxycholate, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM Na3VO4, 1 µg/ml aprotinin, 1 µg/ml leupeptin, and 1 µg/ml pepstatin. Cell lysates were used for immunoprecipitation with PLD1 or PLD2 antibodies.
PLD activation in intact ECs. Bovine pulmonary artery ECs in 35-mm dishes were labeled with either [32P]orthophosphate (5 µCi/ml) or [3H]myristic acid (1 µCi/ml) in DMEM-phosphate-free medium containing 2% FBS for 18-24 h (37, 42). Cells were washed in MEM and incubated with 1 ml of MEM or 1 ml of MEM plus DPV and 0.05% butanol for various time periods as indicated at 37°C. In some experiments, cells were pretreated with various MAPK inhibitors for 1-2 h before addition of DPV (1-5 µM). The incubations were terminated by addition of methanol-concentrated HCl (100:1 vol/vol). Lipids were extracted essentially according to the method of Bligh and Dyer (42) as described previously. [32P]PBt or [3H]PBt formed as a result of PLD activation and transphosphatidylation reaction, an index of in vivo PLD stimulation (32), was separated by TLC in 1% potassium oxalate-impregnated silica gel H plates using the upper phase of ethyl acetate-2,2,4-trimethyl pentane-glacial acetic acid-water (65:10:15:50 vol/vol/vol/vol) as the developing solvent system (13, 37). Unlabeled PBt was added as a carrier during the lipid separation by TLC and was visualized under iodine vapors. Radioactivity associated with the PBt was quantified by liquid scintillation counting, and data are expressed as dpm normalized to 106 counts in total lipid extract or as a percentage of control.
Preparation of cell lysates and Western blotting. After stimulation with DPV, cells were rinsed two times with ice-cold PBS, scraped in 1 ml of lysis buffer containing 20 mM Tris · HCl (pH, 7.4), 150 mM NaCl, 2 mM EGTA, 5 mM glycerophosphate, 1 mM MgCl2, 1% Triton X-100, 1 mM sodium orthovanadate, 10 µg/ml protease inhibitors, 1 µg/ml aprotinin, 1 µg/ml leupeptin, and 1 µg/ml pepstatin, incubated at 4°C for 20 min, and cleared by centrifugation in a microfuge at 10,000 g for 5 min at 4°C. After determination of the total protein in the lysates by the bicinchoninic acid method, 6× Laemmli sample buffer was added to cell lysates and the lysates were boiled for 5 min. Proteins were separated on 7 or 12% gels by SDS-PAGE, transferred to polyvinylidene difluoride membranes, and subjected to immunoblotting with either anti-ERK1/ERK2 (1:1,000 dilution), anti-p38 MAPK (1:2,000 dilution), anti-JNK (1:2,000 dilution), anti-phospho-ERK1/ERK2 (1:2,000 dilution), anti-phospho-p38 MAPK (1:500 dilution), anti-PLD1 (1:500 dilution), anti-PLD2 (1:500 dilution), or anti-phosphotyrosine 4G10 (1:2,000 dilution) overnight at 4°C. The membranes were washed at least three times with Tris-buffered saline containing 0.1% Tween 20 (TBS-T) and were incubated for 2-4 h at room temperature in horseradish peroxidase-conjugated goat anti-rabbit (1:2,000 dilution in TBS-T containing 5% BSA) or goat anti-mouse secondary antibodies (1:5,000 dilution in TBS-T containing 5% nonfat milk). The immunoblots were developed with ECL according to the manufacturer's recommendation.
In vitro PLD assay. Immunoprecipitates of PLD1, PLD2, and p38 MAPK from control and DPV-treated bovine pulmonary artery ECs were assayed for PLD activity by the head group release assay (4). Liposomal substrate (DOPE-PIP2-DPPC in a molar ratio of 16:1.4:1) was prepared by sonicating the lipid mixture with a probe sonicator at a setting of six for 5 × 30 s at room temperature in vesicle buffer containing 50 mM Na-HEPES (pH 7.5), 3 mM EGTA, 80 mM KCl, 1 mM dithiothreitol (DTT), and 3 mM MgCl2. [Choline-methyl-3H]DPPC was added to the liposomal substrate to give 50,000-70,000 dpm/assay. Equal volumes of the immunoprecipitates were added to the reaction mixture containing the liposomal substrate in a final volume of 100 µl in 50 mM Na-HEPES (pH 7.5), 3 mM EGTA, 80 mM KCl, 1 mM DTT, 3 mM MgCl2, and 2 mM CaCl2 and incubated at 37°C for 30 min. To differentiate between PLD1 and PLD2, the incubation mixtures contained ADP-ribosylating factor (Arf)-guanosine 5'-O-(3-thiotriphosphate) (1 µg), wherever required (18). The reactions were terminated by the addition of 200 µl of 10% TCA and 100 µl of 1% BSA. The mixture was briefly vortexed and centrifuged in a microfuge at 10,000 g for 5 min. Radioactivity in the supernatant was measured by liquid scintillation spectroscopy. PLD activity is expressed as picomoles of the substrate hydrolyzed in 30 min in the total immunoprecipitate.
MAPK assays.
Cell lysates (1 mg of protein) were subjected to immunoprecipitation
with anti-ERK1 plus ERK2 (2 µg/ml) at 4°C for 12 h. Protein A/G (20 µl) was added and incubated for an additional 2 h at
4°C (36, 41). The antibody complex was pelleted and
washed two times in ice-cold PBS plus vanadate, and the activity of ERK
in the immunocomplex was determined in a final volume of 50 µl
containing 20 mM Tris · HCl (pH 7.4), 2 mM EGTA, 10 mM
MgCl2, 10 µM [-32P]ATP, and 5 µg of
myelin basic protein (17). For p38 MAPK activity, cell
lysates were subjected to immunoprecipitation with anti-phospho-p38 MAPK antibody, and immunocomplex kinase activity was carried out in a
final volume of 100 µl containing 25 mM Tris · HCl (pH 7.4), 5 mM
-glycerophosphate, 2 mM DTT, 0.1 mM sodium vanadate, 200 µM
ATP, 2 µg of ATF-2 fusion protein, 10 mM [
-32P]ATP
(10 Ci/mmol), and 10 mM MgCl2. Samples were
incubated at 30°C for 30 min, and reactions were terminated by the
addition of 6× Laemmli sample buffer and boiling for 5 min. ATF-2
phosphorylation was detected by autoradiography and quantified by image analysis.
In vitro phosphorylation of PLD1 and
PLD2 with p38 MAPK.
Phosphorylation of PLD1 and PLD2 immunoprecipitates from ECs or
Sf9-overexpressing cells was determined with immunoprecipitates of p38 MAPK from control and DPV-treated cells. Phosphorylation of PLD1
and PLD2 was performed in a final volume of 60 µl containing 20 mM
Tris · HCl (pH, 7.5), 2 mM EGTA, 10 mM MgCl2, 10 µM [-32P]ATP (sp. act. 2,000 dpm/pmol), human (h)
PLD1 or hPLD2 immunocomplexes from 1 mg of total cell lysates, and p38
MAPK immunoprecipitates obtained from control and DPV-treated ECs. In
some experiments, commercial p38 MAPK (activated; 250 ng/incubation)
was used to determine phosphorylation of hPLD1 and hPLD2
immunoprecipitates obtained from either ECs or Sf9 cells
overexpressing hPLD1 or hPLD2 (19). Samples were incubated
for 30 min at 30°C, and reactions were terminated by the addition of
6× Laemmli sample buffer and boiling. Substrate phosphorylations
were detected by autoradiography and quantified by image analysis.
In vivo phosphorylation of PLD1 and PLD2 by DPV. Confluent bovine pulmonary artery ECs in 100-mm dishes were labeled with 10 ml of [32P]orthophosphate (25 µCi/ml) in DMEM-phosphate-free medium containing 2% FBS for 4 h at 37°C. Cells were washed in MEM and pretreated with SB-202190 (25 µM) for 2 h before challenge with DPV (5 µM) for 5 min. Cells were rinsed two times with ice-cold PBS and scraped in 1 ml of lysis buffer containing 20 mM Tris · HCl (pH 7.4), 150 mM NaCl, 2 mM EGTA, 5 mM glycerophosphate, 1 mM MgCl2, 1% Triton X-100, 1 mM sodium orthovanadate, and 10 µg/ml protease inhibitors, incubated at 4°C for 20 min, and cleared by centrifugation at 5,000 g for 5 min at 4°C. Aliquots of equal protein were subjected to immunoprecipitation with anti-PLD1 or anti-PLD2 antibodies (5 µg/ml each of internal plus NH2-terminal antibodies) overnight at 4°C. Protein A/G plus agarose (20 µl) was added, incubated for an additional 2-4 h at 4°C, and centrifuged at 5,000 g for 5 min. The immunocomplexes were washed three times with ice-cold PBS and dissociated by boiling in 1× SDS sample buffer. The samples were subjected to SDS-PAGE on 10% gels, dried in a gel dryer, analyzed by autoradiography, and quantified by image analysis.
Transient expression of dominant negative mutant of p38
MAPK.
Bovine pulmonary artery ECs were cultured in 35-mm dishes until they
reached 80% confluence. Each well was transfected with 1.5 µg/ml of
vector or a plasmid DNA of the dominant negative mutant of p38
cloned in pCMV-5 cDNA in 5 µl of LipofectAMINE reagent in MEM without
serum according to the manufacturer's recommendation. After
5 h of transfection, the medium was replaced by complete medium.
After 24 h of transfection, the cells were labeled with [3H]myristic acid (1 µCi/ml) in complete MEM with 20%
serum for 18-24 h, and the effect of DPV or TPA on PLD activation
was determined. Transiently transfected cells were analyzed for
expression of p38 MAPK by Western blotting, and p38 MAPK activity was
determined by ATF-2 phosphorylation using a p38 kinase assay kit.
p38 MAPK/PLD binding assays in vitro.
GST alone or GST-p38 fusion protein was incubated with 20 µl of
glutathione-Sepharose in 250 µl of PBS containing 0.1% NP-40, 1 mM
PMSF, 5 mM DTT, 1 µg/ml leupeptin, 1 µg/ml aprotinin, and 1 µg/ml
pepstatin for 3 h at 4°C (52). The beads were
washed three times with 200 µl of binding assay buffer [20 mM HEPES
(pH 7.6), 500 mM NaCl, 0.5 mM EDTA, and 0.1% NP-40] and incubated with Sf9 lysates from control or PLD1a/PLD2-overexpressed
cells. The beads were washed with PBS, eluted with 20 µl of SDS
sample buffer [75 mM Tris · HCl (pH 6.8), 0.5% glycerol, 1%
SDS, 4% -mercaptoethanol, and 0.01% bromphenol blue], and boiled
for 5 min before separation on 8% SDS-PAGE. PLD1/PLD2 were
immunodetected by Western blotting with anti-PLD1 or anti-PLD2 antibodies.
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RESULTS |
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DPV activates ERK, p38, and
JNK MAPKs in ECs.
To investigate the role of MAPKs in DPV-induced PLD activation, bovine
pulmonary artery ECs were treated with DPV (5 µM) for various time
periods, and total cell lysates were analyzed for ERK1/2, p38 MAPK, and
JNK activation by Western blotting with phosphospecific antibodies.
As shown in Fig. 1, A-C,
DPV stimulated ERK1/2, p38, and JNK activities in a time-dependent
manner as evidenced by enhanced phosphorylation of threonine/tyrosine
residues. The activation of ERK1/2 and p38 by DPV was dose dependent,
with increased phosphorylation being detected with concentrations as low as 1 µM (Table 1). Figure
2 shows that DPV also increased tyrosine phosphorylation of the three MAPKs detected by immunoblotting with anti-phosphotyrosine antibody. Also, it was observed that DPV
treatment of ECs resulted in increased phosphorylation of myelin basic
protein or ATF-2 in in vitro kinase assays (Fig. 2).
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DPV-induced PLD activation in
ECs requires p38 MAPK activation.
To further elucidate the role of MAPKs in mediating DPV-induced PLD
stimulation, we used selective and known chemical inhibitors for
ERK1/ERK2 and p38 MAPKs. As shown in Table
2, pretreatment of ECs for 2 h with
PD-98059 (25 µM), a specific inhibitor of MAPK kinase (MEK) 1/MEK2
(1), did not affect the DPV-induced [32P]PBt
formation. In contrast, pretreatment of ECs with selective p38
MAPK inhibitor SB-203580 (25 µM), SB-202190 (25 µM), or PD-169316 (25 µM; see Ref. 54) for 2 h before DPV challenge
partially mitigated DPV-induced PLD activation without altering basal
PLD activity (Table 2). In addition, SB-202474 (25 µM), a negative control for SB-203580 and SB-202190, did not significantly alter the
DPV-induced [32P]PBt formation. The effect of SB-202190
on DPV-induced [32P]PBt formation was dose and time
dependent (Fig. 3,
A and B). In parallel experiments, we also
examined the influence of SB-202190 on DPV-induced p38 MAPK activation.
As shown in Fig. 5C, SB-202190 in a dose-dependent manner
attenuated p38 activity as determined by phosphorylation of ATF-2.
SB-202190 had no effect on DPV-induced phosphorylation of p38 MAPK
(Fig. 3C) as determined with phospho-specific p38
antibodies, since SB-202190 directly binds to p38 MAPK and alters its
kinase activity. Furthermore, the effect of SB-202190 in inhibiting p38
MAPK-dependent phosphorylation of ATF-2 was specific, since no effect
on phosphorylation of ERK1/2 was observed (data not shown). These data
suggest that p38 MAPK activation is part of the signaling cascade
involved in DPV-induced PLD stimulation.
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Effect of SB-202190 on TPA-,
bradykinin-, and ATP-induced PLD
activation in ECs.
To further characterize the specificity of the p38 MAPK inhibitor
SB-202190 on agonist-mediated PLD activation, ECs were preincubated with SB-202190 (25 µM) for 2 h before challenge with TPA (25 nM), bradykinin (1 µM), ATP (100 µM), or DPV (5 µM), and
[32P]PBt accumulation was measured. As shown in Table
3, 25 µM SB-202190 attenuated
DPV-induced PLD activation by ~50% but failed to mitigate either
TPA- or bradykinin- or ATP-mediated PLD stimulation. In addition to
SB-202190, we also examined the effect of the MEK1/MEK2 inhibitor
PD-98059 on TPA- or DPV-induced PLD stimulation, which, unlike the p38
MAPK inhibitor, failed to alter either TPA- or DPV-induced
[32P]PBt formation (data not shown). Under similar
incubation conditions, PD-98059 partially blocked activation of ERK1/2
by TPA (25 nM; data not shown). These results further confirm the
specificity of SB-202190 in attenuating DPV-induced p38 MAPK and PLD
activation in ECs.
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Expressing a dominant negative mutant of p38 MAPK
attenuates DPV-induced PLD activation.
To further confirm a role for p38 MAPK in DPV-induced PLD activation,
we transiently transfected ECs with a dominant negative mutant of p38
MAPK or a vector control. Cell lysates from vector or dominant negative
p38 MAPK-transfected ECs were subjected to immunoprecipitation with
anti-p38 MAPK antibody, and the immunoprecipitates were analyzed for
the expression of p38 MAPK protein and assayed for kinase activity in
vehicle- or DPV-treated cells by measuring phosphorylation of ATF-2.
Western blotting of the lysates from vector or dominant negative p38
MAPK-transfected cells revealed overexpression of the p38 MAPK protein
(Fig. 4A). However, p38 MAPK
immunoprecipitates from cells transfected with the dominant negative
mutant of p38 MAPK exhibited lower kinase activity as determined by
phosphorylation of ATF-2 with and without DPV treatment. Transfection
of ECs with the dominant negative mutant of p38 MAPK had no effect on
DPV-induced activation of ERK1/2 or JNK (data not shown). Under similar
experimental conditions, transfection with vector plasmid alone had no
effect on DPV-induced activation of p38 MAPK (Fig. 4A).
Expression of the dominant negative mutant of p38 attenuated
DPV-induced [3H]PBt formation by 40% (Fig.
4B) without altering TPA-mediated PLD activation. These
results further confirm participation of p38 MAPK in DPV-induced PLD
stimulation in ECs.
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Association of PLD1 and PLD2 with p38
MAPK immunoprecipitates.
Because the above data suggested the importance of p38 MAPK in the
activation of PLD by DPV, we investigated whether direct interaction
exists between p38 MAPK and PLD. ECs were exposed to medium alone or
medium containing DPV (5 µM) for 5 min, and cell lysates were
subjected to immunoprecipitation under native conditions with anti-p38
MAPK antibody plus protein A/G Sepharose. Western blot analysis of the
p38 MAPK immunoprecipitates against hPLD1 or hPLD2 antibodies revealed
the presence of PLD1 and PLD2 in the p38 MAPK immunocomplex with and
without treatment with DPV (Fig.
5A). To further confirm the
presence of PLD1 or PLD2 in the p38 MAPK immunocomplex, we evaluated
the p38 MAPK immunoprecipitates from control or DPV-treated cells for
PLD activity using an in vitro assay employing PIP2 and
phospholipid liposomes as described by Brown et al. (4).
As shown in Fig. 5B, a transient increase in
PIP2-dependent PLD activity was observed 5 min after DPV
treatment but returned to near basal values at 30 min. Inclusion of Arf (1 µg/incubation), a known cofactor for PLD1 activity, to the in
vitro assay system further increased the hydrolysis of
phosphatidylcholine in the presence of PIP2. As expected,
DPV treatment enhanced p38 MAPK activity in the immunoprecipitates as
determined by phosphorylation of ATF-2 using [-32P]ATP
followed by SDS-PAGE (data not shown). These data strongly suggest that
PLD1 and PLD2 are associated with p38 MAPK under basal and stimulated
conditions.
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Association of p38 MAPK with hPLD1
and hPLD2 immunoprecipitates.
Based on the above data, we next investigated the possible association
of p38 MAPK in PLD1 and PLD2 immunoprecipitates. Western blot analysis
of hPLD1 and hPLD2 immunoprecipitates from control and DPV-treated ECs
with anti-p38 MAPK antibodies showed an immunodetectable protein band
on SDS-PAGE in the range of 38 kDa, suggesting the possible presence of
p38 MAPK in PLD immunoprecipitates (Fig. 6, A and B). Also,
the hPLD1 and hPLD2 immunoprecipitates from control cells
phosphorylated ATF-2, which was enhanced after DPV treatment (Fig. 6,
A and B). These results further confirm an association between p38 MAPK and hPLD1/hPLD2 under basal and stimulated conditions.
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Binding of p38 MAPK to PLD in vitro.
To further study the interaction between p38 MAPK and PLD, we tested
the binding of an active p38-GST fusion protein to PLD1 and PLD2 in
vitro. Lysates from control Sf9 cells or Sf9 cells overexpressing hPLD1 or hPLD2 were incubated with GST beads or p38
MAPK-GST fusion protein with physiological salt concentrations. Samples
were subjected to SDS-PAGE, and PLD1 and PLD2 were immunodetected by
Western blotting. As shown in Fig. 7,
A and B, PLD1 and PLD2 were detected bound to p38
MAPK-GST fusion protein in lysates from Sf9 cells overexpressing
hPLD1 or hPLD2. No binding of PLD1 to GST was detected when cell
lysates were incubated with GST beads in the absence of p38 MAPK-GST
fusion protein. Compared with PLD1, low binding of PLD2 to GST was
detected (Fig. 7). In addition to Western blotting, the GST and p38-GST
plus PLD pull-down assays were evaluated for PLD1 and PLD2 activities.
As shown in Fig. 7C, incubation of Sf9 cell lysates
overexpressing PLD1 and PLD2 with GST beads alone exhibited activities
of 20.8 and 112.0 pmol of substrate hydrolyzed in 30 min, respectively,
compared with 4.7 pmol of substrate hydrolyzed by cell lysates from
control Sf9 cells. However, in the presence of p38-GST fusion
protein, the beads in the pull-down assay expressed PLD1 and PLD2
activities of 50.4 and 310.2, respectively. These in vitro
protein-protein binding assays further suggest interaction between p38
MAPK and PLD1/PLD2.
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PLD1 and PLD2 are phosphorylated in
vitro by p38 MAPK.
The above data suggest that p38 MAPK activation is involved in
DPV-induced PLD stimulation in ECs. At present, it is not clear whether
PLD is a substrate for phosphorylation by p38 MAPK. To test this, we
incubated p38 MAPK immunoprecipitates prepared from control and
DPV-treated ECs with hPLD1 or hPLD2 immunoprecipitates from control
cells that served as substrates for phosphorylation. As shown in Fig.
8A, incubation of hPLD1 or
hPLD2 immunoprecipitates with [-32P]ATP in the absence
of exogenously added p38 MAPK resulted in slight phosphorylation of
both substrates, suggesting association of p38 MAPK with PLD.
Interestingly, addition of p38 MAPK immunoprecipitates obtained
from DPV-treated cells (as determined by increased
phosphorylation of p38 MAPK and ATF-2 phosphorylation) significantly
enhanced the phosphorylation of hPLD1 and hPLD2. Maximum
phosphorylation of hPLD1 or hPLD2 was observed at 5 min and declined at
15 and 30 min of DPV treatment (Fig. 8A). A commercial
preparation of activated p38 MAPK also phosphorylated hPLD1 and hPLD2
obtained from Sf9-overexpressed cells, thus confirming the
ability of p38 MAPK to use PLD1 or PLD2 as substrates (Fig.
8B). These data suggest that both PLD1 and PLD2 are
phosphorylated by activated p38 MAPK in vitro and that p38 MAPK is
constitutively associated with PLD1 and PLD2.
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SB-202190 attenuates DPV-induced in
vivo phosphorylation of PLD.
To examine whether PLD is a physiological substrate of p38 MAPK, ECs
prelabeled with [32P]orthophosphate were incubated with
25 µM SB-202190 for 2 h before DPV challenge. In vivo PLD1 and
PLD2 phosphorylation increased by 3.5 (Fig.
9A)- and 4.8-fold (Fig.
9B), respectively, after 5 min of DPV treatment.
Interestingly, some basal phosphorylation of PLD1 and PLD2 was observed
in control cells without DPV stimulation. The DPV-induced PLD1 and PLD2
phosphorylation was partially attenuated by the p38 inhibitor
SB-202190. These results demonstrate a p38 MAPK-dependent in vivo
phosphorylation of PLD1 and PLD2 after stimulation by DPV but do not
exclude involvement of other kinases, such as Src, receptor tyrosine
kinases, or protein kinase C (PKC).
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Effect of p38 phosphorylation on PLD activity in
vitro.
The physiological relevance of p38 MAPK-dependent phosphorylation of
PLD was examined by determining PLD activity in the presence of
PIP2. With the use of a commercial preparation of active
p38 MAPK, phosphorylation of PLD1 from Sf9 cells overexpressing
hPLD1 using [32-P]ATP and PLD1 activity at various
time periods of phosphorylation was followed. As shown in Fig.
10A, p38 MAPK phosphorylated
PLD1 in a time-dependent fashion, and no phosphorylation of PLD1 was observed in the absence of p38 or PLD1. Under similar experimental conditions, PLD1 was phosphorylated by p38 for 30 min, and the ability
of the phosphorylated form to hydrolyze
[3H]phosphatidylcholine to choline and PA in the presence
of PIP2 was tested. As shown in Fig. 10B,
addition of p38 MAPK plus ATP in the presence of 5 or 7 µM
PIP2 had no effect on PLD1-catalyzed hydrolysis of
[3H]phosphatidylcholine. In this in vitro PLD assay, the
addition of PIP2 is absolutely essential for PLD1-catalyzed
hydrolysis of the substrate, and attempts to measure the activity of
the phosphorylated PLD1 in the absence of PIP2 were
unsuccessful (data not shown). Taken together, the above results
suggest that phosphorylation of PLD1 by p38 MAPK may not be directly
involved in modulating PLD activation.
|
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DISCUSSION |
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---|
We have demonstrated for the first time the involvement of p38 MAPK in DPV-induced PLD activation in vascular ECs. This conclusion is based on the following experimental evidence: 1) the p38 MAPK inhibitor SB-202190 but not the MEK1/MEK2 inhibitor PD-98059 mitigated PLD activation by DPV without altering TPA, bradykinin, and ATP responses; 2) a rapid and early onset (2 min) of p38 MAPK activation by DPV correlated with the in vitro phosphorylation of PLD1 and PLD2 by p38 MAPK immunoprecipitation from DPV-treated cells, suggesting that PLD is a target of p38 MAPK; 3) there was a stable association between PLD1, PLD2, and p38 MAPK in control and DPV-treated cells; 4) expression of a dominant negative mutant of p38 MAPK attenuated DPV- but not the TPA-induced PLD activation; 5) in vitro binding of PLD1 and PLD2 to p38 MAPK was demonstrated using p38-GST fusion protein; 6) DPV enhanced phosphorylation of PLD1 and PLD2 in vivo, which was attenuated by SB-202190; and 7) in vitro phosphorylation of PLD1 by p38 MAPK had no effect on the PLD activity.
In a variety of mammalian cells and tissues, PLD is activated by hormones, growth factors, neurotransmitters, cytokines, and reactive oxygen species (ROS; see Refs. 9, 12, 34). Depending upon the stimulus, activation of PLD is dependent on PKC, changes in intracellular calcium, heterotrimeric G proteins, modulation of protein tyrosine kinases/phosphatases, and low molecular weight G proteins of the Rho family of GTPases (9, 12). Earlier studies in ECs have shown that ROS- and DPV-induced PLD stimulation was not attenuated by PKC inhibitors or downregulation of PKC by prolonged exposure to TPA (40-42), whereas thrombin-, bradykinin-, or ATP-mediated PLD stimulation was highly dependent on PKC activation (13, 34). However, the ROS- or DPV-induced PLD stimulation in ECs was partially blocked by the Src kinase inhibitor PP-1, indicating a role for protein tyrosine phosphorylation in regulating PLD activation either directly or indirectly (9, 12). Although PLD is phosphorylated at serine/threonine residues in intact cells in response to TPA (24) or at tyrosine residues by pervanadate (28, 47), it is still unclear whether this phosphorylation of PLD in response to a stimulus is of physiological relevance to enhanced hydrolysis of phosphatidylcholine or other phospholipid substrates.
p38 MAPK belongs to subfamilies of mammalian MAPKs that also include ERKs, JNK/stress-activated protein kinases (SAPKs), and ERK5. p38 MAPKs are generally activated via phosphorylation of both threonine and tyrosine residues by distinct and dual specific serine/threonine MAPK kinases (MKK3, MKK4, and MKK6), which in turn are phosphorylated and activated by upstream MKK kinases (43). In bovine pulmonary artery ECs, DPV activated all three MAPK families, as determined by enhanced phosphorylation and activity against exogenous substrates. It was apparent that there were some differences between activity measurements and Western blots with anti-phosphotyrosine antibodies. Although MBP activity or ATF-2 phosphorylation reflects enhanced phosphorylation at threonine/tyrosine residues, the Western blots with anti-phosphotyrosine antibody represent increased phosphorylation at tyrosine residues of ERK or p38 MAPK. Also, dephosphorylation of threonine/tyrosine residues by specific phosphatases may explain the apparent differences between Western blots and activity measurements. These data demonstrate that DPV enhances the phosphorylation and activity of MAPKs in ECs.
In contrast to DPV, treatment with TPA resulted only in the activation of ERK1 and ERK2. Our studies with PD-98059 showed that this MEK1/MEK2 inhibitor specifically attenuated TPA- or DPV-induced ERK phosphorylation but had no effect on TPA- or DPV-induced PLD activation. Similarly, SB-202190, which inhibits by binding to phosphorylated p38 MAPK (and not upstream of MKK3, MKK4, or MKK6), specifically blocked DPV-induced phosphorylation of ATF-2 (54). Furthermore, a 40% reduction in DPV-induced PLD stimulation was observed after transient transfection of ECs with a dominant negative mutant of p38 MAPK, confirming a role for p38 MAPK in the regulation of PLD activation. Under similar experimental conditions of transfection, the expressed p38 MAPK mutant protein exhibited ~10% phosphorylation of ATF-2 in vitro compared with vector controls. This suggests that in addition to p38 MAPK, other pathways involving Src or receptor tyrosine kinases may be involved in DPV-induced PLD activation in ECs.
Although ERK activation regulates PLD activity in PC-12 cells (22), neutrophils (6), and smooth muscle cells (33), our results do not suggest the participation of ERK1/2 in DPV-induced PLD activation. In rat pheochromocytoma PC-12 cells, the H2O2-induced PLD activation and MAPK phosphorylation were attenuated dose dependently by PD-98059 (22). Similarly, PD-98059 blocked formyl-Met-Leu-Phe-mediated activation of ERK and PLD, with a concomitant reduction in respiratory burst in human neutrophils (6). Recent studies in rabbit aortic smooth muscle cells demonstrated that norepinephrine-mediated PLD activation was attenuated by farnesyltransferase inhibitors and by PD-98059, suggesting the Ras/MAPK pathway in regulating PLD activity via a phosphorylation-dependent mechanism (33). In contrast to ERK-dependent activation of PLD in rabbit smooth muscle cells, data reported here confirm the earlier studies showing that vasopressin-induced PLD activation was not dependent on ERK in A7r5 rat vascular smooth muscle cells (23). Thus, depending on the cell type and stimulus, activation of ERK or p38 MAPK regulates PLD either directly or indirectly. Although JNK is also activated by DPV, the present study did not address the role of JNK in DPV-induced PLD stimulation.
We have performed preliminary studies to identify possible mechanism(s)
of PLD regulation by p38 MAPK. Interestingly, our data show a physical
association between PLD1/PLD2 and p38 MAPK in control and DPV-treated
ECs. Furthermore, both PLD1 and PLD2 were phosphorylated in vitro by
p38 MAPK immunoprecipitates from control and DPV-treated cells and a
commercial preparation of activated p38 MAPK, suggesting PLD as a
substrate for the kinase. Also, it was observed that DPV treatment of
ECs enhanced in vivo phosphorylation of PLD1 and PLD2, which was
partially attenuated by the p38 MAPK inhibitor SB-202190. Earlier
studies have identified several other downstream substrates of p38
MAPK. These include ATF-1, ATF-2, cAMP response element binding
protein, MAPK activated protein kinase 2/3, myocyte enhancer factor 2C,
CCAAT/enhancer binding protein, p38-regulated/activated protein kinase,
p53 tumor suppressor protein, and cytosolic phospholipase
A2 (cPLA2; see Ref. 43).
Recently, it has been demonstrated that cPLA2 is a physiological target of p38 MAPK in thrombin-stimulated platelets. However, the p38 MAPK-dependent phosphorylation of cPLA2
appears not to be involved in thrombin-stimulated release of
arachidonic acid (26). We have shown that both PLD1 and
PLD2 are substrates for p38 MAPK-mediated phosphorylation in vivo and
in vitro in ECs; however, in vitro phosphorylation of PLD1 did not
affect the PLD1 activity measured in the presence of PIP2
and Arf. Further studies are necessary to understand the physiological
role of PLD phosphorylation relative to secretion, interaction with
other proteins, or translocation in response to external stimuli. DPV treatment increased tyrosine phosphorylation of PLD1 in HL-60 granulocytes (28) and ECs (47).
Although a variety of agonists, including TPA, activate PLD via a
PKC-dependent pathway, the mechanism of PLD stimulation by PKC is still
controversial. Inhibitors of PKC that block its catalytic activity can
also alter agonist-induced PLD activation, suggesting a
phosphorylation-dependent regulation (27). However, in
vitro studies examining the mechanism of PKC-dependent activation of
PLD have been varied. PKC- and -
isoenzymes in a phorbol ester-dependent manner enhance PLD activity in membrane preparations, partially purified PLD, and baculovirus-expressed PLD isoforms (9). This activation occurs in the absence of ATP,
indicating a non-phosphorylation-dependent mechanism of PLD stimulation
such as protein-specific interaction. In additional studies, it has been shown that PLD1 is associated with PKC-
immunoprecipitates after TPA challenge of fibroblasts (29), and it was
suggested that possible protein-protein interaction between PKC and
PLD1 may be involved in the activation (12). In Rat1
fibroblasts, coimmunoprecipitates of PKC-
and recombinant PLD1 were
associated with an unidentified 220-kDa protein in response to TPA
(12), which may represent a scaffolding protein (9,
29). However, other studies point out that PKC-mediated
phosphorylation of PLD1 in vitro results in an inhibition of PLD
activity (12). In the present study, DPV treatment of ECs
for 30 min compared with 5 min resulted in an inhibition of PLD1
activity measured in the p38 MAPK immunoprecipitates (Fig.
5B). At this time, it is not known if this phosphorylation
also occurs in vivo or if this phosphorylation is important for
activation of the enzyme. Only recently, direct evidence for in vivo
phosphorylation of PLD1 by PKC after TPA stimulation in 3Y1 cells and
in COS-7 cells transiently expressing PLD1 was reported
(24). Inhibitors of PKC or downregulation of PKC by
phorbol esters attenuated TPA-induced phosphorylation and activation of
PLD1. In vitro, in the presence of TPA, purified PLD1 was
phosphorylated by PKC-
at serine-2, threonine-147, and serine-561
residues. Mutation of serine-2, threonine-147, or serine-157 reduced
TPA-induced PLD1 activity in vitro, suggesting involvement of
phosphorylation of PLD1 in regulation of its activation
(24).
The physiological significance of p38 MAPK-mediated phosphorylation of PLD1 or PLD2 is unclear. Although in vitro phosphorylation of PLD1 and PLD2 by a commercial preparation of p38 MAPK had no effect on the catalytic activity, further studies on the role of in vivo phosphorylation of PLD are essential to determine a link between phosphorylation and activation of the enzyme. Because PLD is known to regulate protein trafficking and secretion, it is possible that serine/threonine and/or tyrosine phosphorylation of PLD1 or PLD2 may be critical for its interaction with other Src homology (SH) 2- or SH3- containing proteins and not for activity. Further studies on mapping the p38 MAPK-dependent phosphorylation sites combined with mutation of specific amino acids involved in phosphorylation should provide information to further define the physiological relevance between phosphorylation and function.
In conclusion, our data indicate that p38 MAPK is involved in DPV-induced PLD activation in bovine pulmonary artery ECs. In ECs, both PLD1 and PLD2 were constitutively associated with p38 MAPK, and a dominant negative mutant of p38 MAPK partially mitigated DPV-mediated PLD activation without affecting the TPA response. DPV treatment of ECs enhanced phosphorylation of PLD1 and PLD2 in vivo, which was partially attenuated by SB-202190. However, in vitro phosphorylation of PLD1 by p38 MAPK did not enhance PLD1 activity, suggesting that either phosphorylation may not be directly involved in the activation of the enzyme or that other factors in addition to p38 MAPK-mediated phosphorylation are required for the enhanced enzymatic activity. Further studies are in progress to characterize the binding sites between p38 MAPK and PLD, sites of phosphorylation by p38 MAPK, and a possible physiological role of phosphorylation of PLD activation and generation of PA in EC function.
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ACKNOWLEDGEMENTS |
---|
We thank Patricia Lyon, Wanda Moran, and Dawn Walcott for secretarial assistance and Dr. Rhett Cummings for critical reading of the manuscript.
![]() |
FOOTNOTES |
---|
This study was supported by National Heart, Lung, and Blood Institute Grants HL-47671, HL-57260, and HL-58064 to V. Natarajan.
Address for reprint requests and other correspondence: V. Natarajan, Johns Hopkins Asthma and Allergy Bldg., Division of Pulmonary and Critical Care Medicine, 5501 Hopkins Bayview Circle, Baltimore, MD 21224 (E-mail: vnataraj{at}welch.jhu.edu).
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.
Received 16 January 2001; accepted in final form 7 March 2001.
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