©1996 by The American Society for Biochemistry and Molecular Biology, Inc.
Platelet-derived Growth Factor Stimulates Protein Kinase A through a Mitogen-activated Protein Kinase-dependent Pathway in Human Arterial Smooth Muscle Cells (*)

(Received for publication, August 1, 1995; and in revised form, October 16, 1995)

Lee M. Graves (1)(§) Karin E. Bornfeldt (2) Jaspreet S. Sidhu (3) Gretchen M. Argast (1) Elaine W. Raines (2) Russell Ross (2) Christina C. Leslie (4) Edwin G. Krebs (1)(¶)

From the  (1)Departments of Pharmacology, (2)Pathology, and (3)Environmental Health, University of Washington, Seattle, Washington 98195 and the (4)Department of Pediatrics, The National Jewish Center, Denver, Colorado 80206

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The abilities of platelet-derived growth factor (PDGF) and insulin-like growth factor (IGF-I) to regulate cAMP metabolism and mitogen-activated protein kinase (MAP kinase) activity were compared in human arterial smooth muscle cells (hSMC). PDGF-BB stimulated cAMP accumulation up to 150-fold in a concentration-dependent manner (EC approx 0.7 nM). The peak of cAMP formation and cAMP-dependent protein kinase (PKA) activity occurred approximately 5 min after the addition of PDGF and rapidly declined thereafter. Incubating cells with PDGF and 3-isobutyl-1-methylxanthine (IBMX, a phosphodiesterase inhibitor) enhanced the accumulation of cAMP and PKA activity by an additional 2.5-3-fold, whereas IBMX alone was essentially without effect. The PDGF-stimulated increase in cAMP was prevented by addition of the cyclooxygenase inhibitor indomethacin, consistent with release of prostaglandins stimulating cAMP. PDGF, but not IGF-I, stimulated MAPK activity, cytosolic phospholipase A(2) (cPLA(2)) phosphorylation, and cAMP synthesis which indicated a key role for MAP kinase in the activation of cPLA(2). Further, PDGF stimulated the rapid release of arachidonic acid and synthesis of prostaglandin E(2) (PGE(2)) which could be inhibited by a cPLA(2) inhibitor (AACOCF(3)). Calcium mobilization was required for PDGF-induced arachidonic acid release and PGE(2) synthesis but not for MAPK activation, whereas PKC was required for PGE(2)-mediated activation of PKA. In summary, these results demonstrate that PDGF increases cAMP formation and PKA activity through a MAP kinase-mediated activation of cPLA(2), arachidonic acid release, and PGE(2) synthesis in human arterial smooth muscle cells.


INTRODUCTION

The proliferation of arterial smooth muscle cells is a key event in the formation and progression of lesions of atherosclerosis and in restenosis following angioplasty. This proliferation is most likely initiated and regulated by growth factors, such as the platelet-derived growth factors, PDGF-AA, (^1)PDGF-BB, and PDGF-AB. In atherosclerotic lesions, a major source of PDGF-BB is activated macrophages, although smooth muscle, endothelial cells, and other cells can also express and secrete PDGF dimers (reviewed in (1) ).

Many growth factor receptors, including PDGF receptors, activate a signal transduction pathway that includes conversion of inactive RasbulletGDP to active RasbulletGTP, activation of Raf, MAP kinase kinase (MAPKK or MEK(2) ), and MAP kinase (MAPK) (for review, see (3) ). Activation of the MAPK cascade results in the stimulation of DNA synthesis and cell proliferation, which can be inhibited by expression of a dominant-negative MAPKK(4, 5) . Conversely, expression of a constitutively active form of MAPKK can stimulate cell proliferation and transformation(4, 6) . A number of nuclear and non-nuclear proteins have been identified as substrates for MAPK. Among the latter is phospholipase A(2), thereby providing a potential link to arachidonic acid metabolism (reviewed in (7) ).

Activation and translocation of the cytosolic 85-kDa phospholipase (cPLA(2)), which catalyzes the release of arachidonic acid from the sn-2 position of phospholipids in the plasma membrane, is an important signal leading to prostaglandin synthesis (reviewed in (8) and (9) ). This enzyme can be distinguished from the low molecular weight forms of PLA(2) by insensitivity to disulfide-reducing agents or inhibition by arachidonic acid analogues (for reviews, see (10) and (11) ). Regulation of cPLA(2) appears to be critically dependent on the integration of multiple signals, including intracellular calcium, protein kinase C (PKC), and phosphorylation by MAP kinase(12, 13, 14, 15) . However, in some cells, calcium-independent forms of cPLA(2)(16) and PKC-independent mechanisms of cPLA(2) activation have also been observed (15) .

Although the coupling of hormonal and neurotransmitter receptors to cAMP synthesis is well established (reviewed in (17) ), the means by which growth factor receptor tyrosine kinases (e.g. the PDGF receptor) regulate cAMP metabolism is less well understood. At present, there are few examples of growth factors stimulating cAMP accumulation. Before elucidation of the MAP kinase cascade, it was reported that PDGF (in the presence of phosphodiesterase inhibitors) increased cAMP synthesis 6-8-fold in Swiss 3T3 cells(18) . In perfused rat hearts, epidermal growth factor (EGF) was found to stimulate cAMP accumulation (19) . In epithelial cells overexpressing the EGF receptor (A431 cells), EGF alone did not affect cAMP accumulation, but did potentiate an increase in cAMP in response to cAMP-elevating agents(20) .

Sustained elevation of cAMP inhibits the proliferation of many cell types, including smooth muscle cells(21) . This phenomena may in part be explained by the fact that in many cell types, including human arterial SMC (hSMC), MAP kinase activation is inhibited by the cyclic AMP-dependent protein kinase (PKA)(22, 23, 24, 25, 26) . The target for inhibition by PKA in the MAP kinase pathway may be Raf-1(23, 27) , although other unidentified targets are also likely to play an important role(28) .

To further evaluate how growth regulatory molecules may regulate cAMP metabolism in hSMC, we examined the effect of two major factors known to influence hSMC (i.e. PDGF and IGF-I). We report here that PDGF rapidly stimulates cAMP synthesis through a mechanism requiring intracellular calcium, PKC activity, MAPK-dependent phosphorylation of cPLA(2), and activation of prostaglandin synthesis, an effect which culminates in increased PKA activity.


EXPERIMENTAL PROCEDURES

Materials

Recombinant human PDGF-BB and PDGF-AA were gifts from Hoffman-La Roche Inc. (Basel, Switzerland). Human recombinant IGF-I was obtained from Upstate Biotechnology Inc. (UBI, Lake Placid, NY). PKI peptide (TTYADFIASGRTGRRNAIHD), a specific peptide inhibitor of PKA(29) , and Leu-Arg-Arg-Ala-Ser-Leu-Gly (Kemptide) (30) were synthesized at the Peptide Synthesis Facility, Howard Hughes Medical Institute (Seattle, WA). Recombinant rat Erk2 was a gift from Dr. M. Cobb (University of Texas, Southwestern, Dallas, TX), and anti-peptide antibodies to this protein were developed previously in this laboratory by Dr. R. Seger. Goat anti-rabbit alkaline phosphatase conjugate was obtained from Promega. Indomethacin, PMA, IBMX, and thapsigargin were obtained from Sigma, and forskolin was obtained from Calbiochem. These compounds were dissolved in dimethyl sulfoxide prior to use. AACOCF(3), AACOCH(3), and recombinant cPLA(2) were gifts from Dr. M. Gelb (University of Washington, Seattle, WA). In some instances, AACOCF(3) was obtained from Biomol (Plymouth Meeting, PA) as well as was the PKC inhibitor GF109203X. The arachidonic acid analogues were dissolved in ethanol or dimethyl sulfoxide prior to use. Polyclonal antibodies to cPLA(2) were prepared in rabbits immunized with human recombinant cPLA(2) as described previously (31) .

Cell Cultures

Human newborn arterial smooth muscle cells were obtained from the thoracic aorta of infants following death due to congenital heart defects or sudden infant death. The cells were isolated by the explant method and cultured as described previously (32) . Cells were used at passages 5 to 10, and were characterized as smooth muscle cells by morphologic criteria and by expression of smooth muscle alpha-actin. The cells were negative in mycoplasma assays and had a normal chromosome number. Subconfluent cell cultures were kept in Dulbecco's modified Eagle's medium (DMEM)/1% plasma-derived serum (PDS) for 2 days prior to experiments.

Measurement of Cyclic AMP

Smooth muscle cells in 10-cm dishes were stimulated with growth factors, tumor promoters, or inhibitors for the indicated periods of time. A typical experiment contained 3 times 10^6 cells per 10-cm plate. The plates were washed 3 times with cold phosphate-buffered saline, and the cAMP was rapidly extracted with the addition of 1 ml of ice-cold ethanol (70%). After scraping the plates, the suspension was centrifuged at 13,000 times g for 20 min at 4 °C, the supernatant was collected, and 0.5 ml was evaporated in a Speed Vac centrifuge. Levels of cAMP were determined using a cAMP enzyme-linked immunosorbent assay kit in prototypic stage; this was generously provided by Life Technologies, Inc.

Measurement of PKA Activity

PKA was assayed by measuring phosphorylation of the peptide substrate Kemptide(30) , (0.17 mM) in the presence or absence of a 10 µM concentration of the PKA inhibitor peptide (PKI) (29) as described(22) . PKA activity was calculated as the amount of Kemptide phosphorylated in the absence of PKI peptide minus that phosphorylated in the presence of PKI peptide. The assay as described is influenced by the amount of cAMP carried over from the lysates into the kinase assay. In some instances, the activity ratio was determined for PKA as described by Corbin(33) .

Measurement of MAPK and MAPKK Activity

MAPK and MAPKK activity was measured as described earlier(32) . Cells in 10-cm dishes (approximately 3 times 10^6 cells) were incubated with PDGF, IGF-I, or PMA for the indicated times. Immediately after stimulation, the cells were scraped and sonicated in buffer H (50 mM beta-glycerophosphate, pH 7.4, 1.5 mM EGTA, 0.1 mM Na(3)VO(4), 1 mM dithiothreitol, 25 µg/ml aprotinin, 25 µg/ml leupeptin, and 0.5 mM phenylmethylsulfonyl fluoride. For measurement of MAPKK activity, cell lysates were precleared on a DE52 mini column, and the phosphorylation of myelin basic protein by activated recombinant Erk2 was measured. MAPK activity was determined by the incorporation of radioactivity from [-P]ATP into myelin basic protein after a 15-min incubation at 30 °C.

Immunoblotting of MAP Kinase

Samples were immunoblotted for MAP (Erk1 and Erk2) after separation of samples on SDS-PAGE and transfer to Immobilon-P (Millipore). The antibody(7884) was raised against a carboxyl-terminal peptide of Erk2 and previously shown to detect both Erk1 and Erk2. The detection was completed by the use of goat-anti rabbit antibodies (alkaline phosphatase conjugate), and the color was developed as per the manufacturer's instruction (Promega).

Measurement of Arachidonic Acid Release and Prostaglandin E(2) (PGE(2)) Release

For measurement of arachidonic acid release, cells in 6-well plates were labeled with [5,6,8,11,12,14,15-^3H]arachidonic acid (Dupont NEN) at 1 µCi/ml in DMEM, 1% human PDS for 24 h at 37 °C according to Domin and Rozengurt(34) . The cells were washed three times in new DMEM without [^3H]arachidonic acid and subsequently stimulated as indicated. The medium (1 ml) was collected, and the amount of released [^3H]arachidonic acid was determined by measuring the radioactivity in the medium. The radioactivity released prior to stimulation was subtracted from the amount released at the end of the stimulation. For measurement of PGE(2) synthesis, cells in 6-well plates were washed three times in DMEM (without PDS or bovine serum albumin), and then stimulated as indicated in 1 ml of new medium. PGE(2) released to the medium was measured after a 1:10-1:30-fold dilution using a PGE(2) enzyme immunoassay (Amersham). The cross-reactivity of this assay is less than 7% with PGE(1) and less than 5% with other related prostaglandins.

Phosphorylation of cPLA(2)

Cells in two maxi-plates (approximately 15 times 10^6 cells/sample) were stimulated with vehicle (10 mM acetic acid, 0.25% bovine serum albumin), 1 nM PDGF-BB, or 10 nM IGF-I for 5 min. Cell extracts were prepared in Buffer H as described above and applied to a Mono Q (Pharmacia) column, which was developed with a linear gradient of 0-400 mM NaCl in the same buffer. Fractions of 1 ml were collected and were assayed (5 µl) for cPLA(2) phosphorylation by adding 1 µg of recombinant cPLA(2) to a reaction mixture as described above for the MAP kinase assay. The recombinant cPLA(2) was confirmed by immunoblotting with antibodies to cPLA(2). To control for nonspecific phosphorylation, in some instances cPLA(2) was omitted from the assay. The reaction was terminated by the addition of SDS-PAGE sample buffer, the samples were heated, separated on SDS-PAGE (10% acrylamide), and the gels were dried. Radioactive cPLA(2) was visualized by autoradiography. Mono Q fractions were also assayed for MAPK activity and immunoblotted for MAPK (Erk1, -2) protein as described above.

Immunoblotting of cPLA(2)

Lysates from cells incubated with PDGF-BB or IGF-I for 5 min were obtained by harvesting the cells in a buffer containing 50 mM Hepes, pH 7.4, 10% glycerol, 150 mM NaCl, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 200 µM sodium orthovanadate, 10 mM tetrasodium pyrophosphate, 100 mM sodium fluoride, 3 µMp-nitrophenyl phosphate, 10 µg/ml aprotinin, 10 µg/ml leupeptin, and 1 mM phenylmethylsulfonyl fluoride. The samples were added to an equal volume of SDS-PAGE sample buffer, boiled for 5 min, and were applied to a 10% SDS-polyacrylamide gel containing 1% bisacrylamide. Electrophoresis was carried out until the 80-kDa prestained standard migrated to 1 cm above the end of the gel. After being transferred to nitrocellulose, the membranes were incubated with polyclonal antibody to cPLA2 (1:1000) and the immunodetection was accomplished by the use of goat anti-rabbit horse radish peroxidase and enhanced chemiluminescence (ECL).


RESULTS

PDGF Stimulates a Transient Increase in cAMP and PKA Activity

Incubation of hSMC with PDGF-BB (1 nM) stimulated a rapid increase in cAMP accumulation (approximately 50-fold) that was detected as early as 2.5 min after the addition of the growth factor and reached a maximal level by 5 min (Fig. 1A). The PDGF-stimulated increase in cAMP did not require the addition of phosphodiesterase inhibitors. In parallel with cAMP, PDGF-BB increased PKA activity up to 7-8-fold as determined by Kemptide phosphorylation (Fig. 1A). The PDGF-stimulated increase in cAMP and PKA activity was transient and disappeared within 15 min with no further increase in cAMP detected up to 1 h after PDGF addition. These results were confirmed in different isolates of human newborn arterial smooth muscle cells (data not shown). Incubation of hSMC with the phosphodiesterase inhibitor, IBMX (500 µM), increased the PDGF-stimulated PKA activity by an additional 2-fold, whereas incubation with IBMX alone only slightly elevated the PKA activity (data not shown). Similar results were found with IBMX on cAMP accumulation and suggested that PDGF was stimulating cAMP synthesis, rather than inhibiting cAMP breakdown.


Figure 1: PDGF-BB and PDGF-AA, but not IGF-I, increase cAMP and activate PKA. Human SMC were incubated in 10-cm plates containing DMEM plus 1% PDS for 48 h. A shows the time course of cAMP formation by 1 nM PDGF-BB (box) and 1 nM IGF-I (circle) and PKA activation by 1 nM PDGF-BB (). Samples were rapidly harvested, and the cAMP concentration was measured in triplicate by radioimmunoassay whereas PKA activity was measured as phosphorylation of Kemptide in the presence or absence of the PKA inhibitor PKI. B shows dose-response curves for PDGF-BB (box), PDGF-AA (), and IGF-I (circle) on cAMP formation. The results are shown as mean ± S.E. of triplicate or duplicate samples. These experiments were repeated 3 times with similar results.



The stimulation of cAMP accumulation was dose-dependent, with up to a 150-fold increase over basal levels (10-50 pmol/ml) seen at the highest concentration (10 nM) of PDGF-BB (Fig. 1B). The PDGF isoform, PDGF-AA, also increased cAMP levels in hSMC, although not to the same extent as PDGF-BB. This is likely a reflection of the 10-fold lower number of PDGF receptor alpha subunits compared to PDGF receptor beta-subunits in these cells(32) . Interestingly, despite the fact that hSMC also contain functional IGF-I receptors (similar in number to PDGF-alpha receptors), IGF-I, at concentrations as high as 10 nM, did not increase cAMP synthesis, even in the presence of the phosphodiesterase inhibitor IBMX (Fig. 1, A and B, and data not shown).

Connection between cAMP Synthesis, MAP Kinase Activation, Intracellular Calcium, and PKC Activity

PDGF-BB stimulated MAPK activity approximately 2.5-fold, whereas IGF-I was without effect (Fig. 2). The PDGF-induced MAPK activation was transient, with a peak of activity occurring at 5 min after stimulation in hSMC and levels returning to basal 15 min later(32) . Thus, activation of MAPK correlated with activation of PKA.


Figure 2: PDGF-BB-induced cAMP formation, but not MAPK activation is dependent on calcium mobilization and PKC. Human SMC in 10-cm plates were incubated for 5 min with 1 nM PDGF-BB, 1 nM IGF-I or for 20 min with 100 nM PMA. In some cells, PKC was down-regulated by a 20-h preincubation with 1 µM PMA, or intracellular calcium stores were depleted by a 1-h preincubation with 300 nM thapsigargin. Levels of cAMP (A) were measured by the cAMP immunoassay as in Fig. 1, and MAPK activity (B) was measured as phosphorylation of MBP for 15 min at 30 °C. These results are shown as mean ± S.E. of duplicate samples. These experiments were repeated twice with similar results.



Since calcium is a key second messenger in the regulation of cAMP metabolism in many cell types (reviewed in (35) and (36) ), we investigated whether calcium was required for the stimulation of cAMP synthesis by exposing cells to the tumor promoter, thapsigargin. Thapsigargin stimulates the release of calcium from intracellular stores, resulting in an initial increase of intracellular calcium and later (at approximately 1 h), as the calcium is transferred to the extracellular space, a depletion of calcium from intracellular stores (37) . Addition of PDGF-BB to hSMC rapidly increases intracellular calcium, an event that can be inhibited by prior incubation with 300 nM thapsigargin for 1 h. (^2)Preincubation with thapsigargin (300 nM, 1 h) eliminated the synthesis of cAMP stimulated by PDGF-BB, demonstrating a requirement for intracellular calcium in this process (Fig. 2A). The effect of this compound on the activation of MAPK by PDGF was examined. Neither an extended (1-h) (Fig. 2B) nor a brief (5-min) (data not shown) exposure of cells to thapsigargin affected the basal or PDGF-stimulated levels of MAP kinase activity in these cells (Fig. 2B). However, a brief incubation with thapsigargin (5 min) did increase PKA activity above basal levels (thapsigargin, 404 ± 16 pmol/min/ml; basal, 80 ± 2 pmol/min/ml) and cAMP accumulation (data not shown).

Protein kinase C (PKC) has also been implicated in the regulation of cAMP formation in many cell types (reviewed in (36) ). Therefore, the requirement for protein kinase C (PKC) activity in PDGF-stimulated cAMP synthesis was investigated. Incubating hSMC with PMA (1 µM, 20 h) resulted in the complete loss of PMA-stimulated MAPK and MAPKK activity and in keeping with a known effect of this procedure for down-regulation of PKC activity (data not shown). Down-regulation of PMA-sensitive PKC activity inhibited the PDGF-stimulated increase in cAMP by >90% (Fig. 2A), without a significant influence on the activation of MAPK by the growth factor (Fig. 2B). Similarly, addition of the PKC inhibitor (1 µM), bisindoylmaleimide (GF109203X) prevented the formation of cAMP by PDGF without inhibiting that stimulated by forskolin (data not shown). Interestingly, PMA alone did not increase cAMP accumulation, although it did activate MAP kinase as expected (Fig. 2B).

Prostaglandin E(2) Synthesis Is Required for cAMP Formation

Previously, Rozengurt reported that PDGF stimulates cAMP synthesis through a prostaglandin-dependent pathway in Swiss 3T3 cells(18) . Since the rate-limiting step in prostaglandin synthesis is believed to be the cPLA(2)-catalyzed release of arachidonic acid(8, 9) , the ability of PDGF and IGF-I to stimulate arachidonic acid release in hSMC was examined. PDGF-BB stimulated the release of [^3H]arachidonic acid within 2.5-5 min of addition, after which the accumulation leveled off (between 10 and 20 min) (data not shown). As shown in Fig. 3A, both PDGF-BB and PDGF-AA (less potently) stimulated arachidonic acid release, whereas IGF-I was without effect. Incubation of hSMC with thapsigargin for 1 h prior to PDGF abolished the PDGF-stimulated increase of arachidonic acid release (Fig. 3A). Previously, it was shown that cPLA(2) activity could be inhibited by a class of arachidonic acid analogues(39, 40) . To confirm that the release of [^3H]arachidonic acid was due to cPLA(2) activation, we incubated hSMC with the analogue AACOCF(3) or the noninhibitory analogue AACOCH(3) prior to the addition of PDGF. The stimulation of [^3H]arachidonic acid release by PDGF was significantly inhibited by AACOCF(3) (>50%), whereas the analogue AACOCH(3) was without effect (Fig. 3A).


Figure 3: PDGF stimulates arachidonic acid- and PGE(2) release via a cPLA(2) and calcium-dependent mechanism. Human SMC in 6-well plates were incubated in DMEM with 1% PDS for 48 h. In A, the cells were labeled with 1 µCi/ml [^3H]arachidonic acid for the last 24 h. The cells were washed three times with fresh DMEM without [^3H]arachidonic acid, and release of [^3H]arachidonic acid to the medium during a 5-min stimulation with 1 nM PDGF-BB, 1 nM PDGF-AA, 1 nM IGF-I was measured. In some instances, intracellular calcium stores were depleted by preincubation with 300 nM thapsigargin, or cPLA2 activity was inhibited by a 30-min preincubation with 30 µM AACOCF(3). The AACOCF(3) analogue AACOCH(3) (30 µM) was used as a control. In B, the cells were washed 3 times with DMEM and then stimulated with 1 nM PDGF-BB (box) or vehicle (circle, 10 mM acetic acid, 0.25% bovine serum albumin) for the indicated periods of time. PGE(2) release into the medium was measured using a PGE(2) enzyme immunoassay (Amersham). The results are expressed as mean ± S.D. of triplicate samples. The experiment was repeated twice with similar results.



One of the major products of arachidonic acid metabolism in hSMC is prostaglandin E2 (PGE(2))(38) , which is formed by the action of cyclooxygenases on arachidonyl precursors. To investigate the potential role of prostaglandin release in cAMP formation, hSMC were incubated with the cyclooxygenase inhibitor, indomethacin, prior to the addition of PDGF-BB. As seen in Table 1, indomethacin (10 µM, 30 min) completely inhibited the PDGF-stimulated increase in cAMP and PKA activity and slightly inhibited the basal levels of cAMP. Incubation with indomethacin did not affect the activation of MAPK, suggesting that the effects of this compound were specific to inhibition of cyclooxygenase activity. These results demonstrated that prostaglandin synthesis was required for PDGF-stimulated cAMP synthesis.



To specifically investigate whether PGE(2) was involved in stimulating the increase in cAMP, PGE(2) formation was measured in response to PDGF. Within 1 min of addition, PDGF increased PGE(2) synthesis in hSMC, which continued to accumulate with extended exposure to PDGF (Fig. 3B). PGE(2) increased PKA activity by 330% as early as 1 min after addition (187.6 ± 0.6 to 622.6 ± 12.0 pmol/min/ml). The peak of PKA activity occurred within 5 min of PGE(2) addition, in good agreement with the ability of PDGF to rapidly increase PKA activity through PGE(2) release (Table 2). PDGF-induced PGE(2) release was dependent on intracellular calcium. Thus, both arachidonic acid release and PGE(2) release stimulated by PDGF were inhibited by depletion of intracellular calcium stores using thapsigargin (Fig. 3A and data not shown).



Down-regulation of PKC inhibited the ability of PGE(2) to stimulate PKA activity from 6.5-fold in vehicle-treated cells to 1.4-fold in cells subjected to PKC down-regulation (Table 2). In contrast, neither down-regulation of PKC nor addition of the PKC inhibitor, bisindolylmaleimide, inhibited the formation of cAMP stimulated directly by forskolin (data not shown). PKC down-regulation did not inhibit PDGF-induced PGE(2) release during a 5-min stimulation (11.1 ± 0.6 to 30.6 ± 0.7 ng of PGE(2) released/10^6 cells subjected to PKC down-regulation compared to 3.3 ± 0.4 to 25.7 ± 1.4 ng of PGE(2) released/10^6 vehicle-treated cells).

PDGF Stimulates MAPK Activation and cPLA(2) Phosphorylation

Since the peak of PDGF-stimulated cAMP and PKA correlated with the maximal activation of MAPK in these cells ( (32) and Fig. 1), we investigated whether MAPK was directly involved in the mechanism of PKA activation. To investigate the phosphorylation of cPLA(2) by MAPK, cell extracts were subjected to Mono Q chromatography and examined for their MBP phosphorylating activity. The fractions testing positive for MAPK activity were then characterized with respect to the presence of this enzyme as shown immunochemically and by cPLA(2) phosphorylating ability. As seen in Fig. 4A, PDGF-BB potently stimulated MAP kinase activity (MBP phosphorylation), whereas IGF-I did not increase this activity. Immunoblotting of the peak fractions from PDGF-treated cells confirmed that these samples contained phosphorylated and active MAPK (Erk1 and Erk2) as judged by band shift and further demonstrated that fractions from unstimulated or IGF-I-stimulated cells did not contain detectable MAPK activity (Fig. 4B). As shown in Fig. 4, fractions from PDGF-stimulated cells contained one major peak of MBP phosphorylating activity (Fig. 4A) that co-eluted with the peak of cPLA(2) phosphorylation (Fig. 4C). Fractions from IGF-I-stimulated cells did not contain cPLA(2) phosphorylating activity above basal levels (Fig. 4C). These results suggested that the deficiency in cAMP synthesis observed with IGF-I was due to the inability of this growth factor to significantly activate MAPK.


Figure 4: PDGF-induced MAPK activity phosphorylates cPLA(2)in vitro. Human SMC in 10-cm plates were incubated without addition (up triangle), with 1 nM PDGF-BB (box) or 1 nM IGF-I (circle) for 5 min. The samples were separated on a Mono Q column, and the fractions were measured for MAP kinase activity (A). Fractions 22, 30, 32, 34, 36, and 44 from each sample were immunoblotted for the presence of MAPK by using an anti-ERK antibody (B). The same fractions were incubated with 1 µg of recombinant cPLA(2), and the phosphorylation of cPLA(2) (C) was measured as described under ``Experimental Procedures.'' Lane C contains active Erk2 (B) or cPLA(2) phosphorylated by recombinant Erk2 (C). This experiment was repeated twice with similar results.



We further investigated the phosphorylation of endogenous cPLA(2) under conditions that led to the activation of cAMP synthesis in hSMC. Phosphorylation of cPLA(2) by MAPK results in a mobility shift on SDS-PAGE that correlates with the activation of this enzyme(13) . Extracts of hSMC stimulated with PDGF or IGF-I were examined by immunoblotting for cPLA(2). As seen in Fig. 5, the cPLA(2) from untreated hSMC was a doublet similar to the baculovirus-expressed human recombinant cPLA(2) standard which is partially phosphorylated in the SF9 cells(31) . Samples from PDGF-treated cells showed that cPLA(2) mobility was shifted completely relative to the untreated or IGF-I-treated samples. The lack of effect of IGF-I is consistent with the finding that IGF-I did not activate MAPK in these cells and that cPLA(2)-dependent release of arachidonic acid was not stimulated by IGF-I.


Figure 5: PDGF-induced phosphorylation of cPLA(2) in hSMC. Lysates (50 µg) from hSMC treated with vehicle (lane 2), 1 nM PDGF-BB (lane 3), or 10 nM IGF-I (lane 4) for 5 min were immunoblotted using an antibody to cPLA(2) as described under ``Experimental Procedures.'' The standard human cPLA(2) (5 ng) was purified from baculovirus-infected Sf9 cells (lane 1). The upper band represents the phosphorylated form of cPLA(2). The experiment was repeated twice with similar results.



Influence of PDGF-stimulated PKA on MAPK Activation

We previously demonstrated that PKA could inhibit PDGF-stimulated MAPKK and MAPK activity in hSMC, an effect that was observed with forskolin or PGE(2)(22) . We therefore investigated whether the stimulation of PKA activity by PDGF could affect the activation of the MAP kinase cascade. Addition of indomethacin (10 µM, 30 min) prior to PDGF abolished the growth factor-stimulated increase in cAMP and PKA activity without affecting the magnitude of MAPKK or MAPK activation by PDGF ( Table 1and data not shown). Furthermore, addition of indomethacin did not alter the time course of MAPKK or MAPK activation in response to PDGF in hSMC (data not shown).


DISCUSSION

PDGF initiates a multitude of biological effects through the activation of intracellular signal transduction pathways such as the MAP kinase cascade, phosphatidylinositol turnover, and calcium mobilization (reviewed in (41) ), and these effects are believed to contribute to smooth muscle cell proliferation and directed migration (32) . Further, changes in eicosanoid metabolism can regulate smooth muscle cell growth and contraction through alterations in cAMP metabolism and calcium homeostasis (reviewed in (38) ). How these key signal transduction pathways are integrated is not presently well understood. Because of our interest in the cross-talk between the MAPK cascade and PKA, we investigated the effects of growth factors on cAMP metabolism in hSMC. We found that PDGF induces a strong and rapid formation of cAMP through a mechanism that includes MAP kinase-mediated activation of cPLA(2), release of arachidonic acid, prostaglandin PGE(2), and the subsequent activation of adenylyl cyclase. Although parts of these signaling pathways have been described previously in other cell types, we report here the complete conversion of a growth factor signal (i.e. PDGF) to cAMP in primary cultures of normal diploid cells, specifically human arterial SMC. In addition, our studies demonstrate that at least three independent signals, i.e. calcium, PKC, and MAP kinase activity are necessary for this event to occur. Several observations support these concepts.

Prostaglandins, such as PGE(2) are produced from arachidonyl precursors and potently stimulate cAMP formation and PKA activity in human and other smooth muscle cells(22, 38, 42) . In the experiments described here, the PDGF-dependent formation of cAMP required prostaglandin synthesis, as it was completely inhibited by the cyclooxygenase inhibitor indomethacin. Our results demonstrate that both arachidonic acid release and PGE(2) formation were stimulated by PDGF and that the rapid increase in PGE(2) synthesis could account for the formation of cAMP. Addition of PGE(2) was sufficient to trigger PKA activation in hSMC, and the time course of PGE(2) formation was consistent with the activation of PKA elicited by PDGF. In support of this model, Rozengurt et al. (18) reported that addition of PDGF to Swiss 3T3 cells stimulated a slow, sustained formation of cAMP that was prevented by the addition of indomethacin. However, in Swiss 3T3 cells, addition of phosphodiesterase inhibitors was required to observe this effect. In contrast, in hSMC, PDGF stimulates a potent increase in cAMP and PKA activity in the absence of phosphodiesterase inhibitors. Interestingly, cyclic AMP is a mitogen for Swiss 3T3 cells(43) , whereas this nucleotide inhibits the proliferation of smooth muscle cells(21) .

We examined the possibility that, in addition to effects of PDGF on prostaglandin metabolism, alternative mechanisms could facilitate the coupling of growth factor signals to cAMP synthesis in hSMC. We were unable to find a direct effect of PDGF on adenylyl cyclase activity in membranes obtained from hSMC or on the phosphorylation of the alpha-subunit of the G-protein G(s), (^3)in contrast to results obtained in epidermal growth factor (EGF)-stimulated cells(44, 45, 46) . Instead, our results suggest that increased prostaglandin metabolism through the activation of cPLA(2) can account for the majority of PDGF-stimulated cAMP synthesis observed in hSMC.

Both PDGF-BB and PDGF-AA potently stimulated MAPK activity, cPLA(2) phosphorylation, arachidonic acid release, and cAMP synthesis in this study. IGF-I did not influence any of these events, despite the fact that in hSMC the number of IGF-I receptors is equivalent to the number of PDGF-alpha receptors, and that the IGF-I receptor is coupled to phosphatidylinositol turnover, calcium mobilization, and chemotaxis(32) . In the present study, the inability of IGF-I to elevate cAMP correlates with an absence of effect of this growth factor on MAPK activity and cPLA(2) phosphorylation in hSMC. These experiments confirm the findings of others(13, 14, 15) , demonstrating that MAPK phosphorylation is an essential signal in the activation of cPLA(2). Recently, Sa et al. (47) reported that, in endothelial cells, basic fibroblast growth factor stimulates a MAPK-dependent activation of cPLA(2), supporting the role for MAPK in cPLA(2) regulation in other cell types.

In addition to MAP kinase activation, PKC activity and intracellular calcium mobilization were critical for the activation of PKA by PDGF in SMC. Since both cPLA(2)(13, 14, 15) and adenylate cyclase (48, 49, 50) can be regulated by calcium and PKC in other cell types, we investigated some of the mechanisms responsible for this calcium and PKC dependence. PMA did not significantly stimulate arachidonic acid release, PGE(2) release, or cAMP synthesis, although PMA increased both MAPK and PKC activities as expected. Thus, activation of MAPK or PKC alone is insufficient for stimulation of cAMP synthesis in hSMC. PKC down-regulation blocked both PDGF-induced and PGE(2)-induced PKA activation, without affecting the ability of forskolin to activate the adenylate cyclase or the ability of PDGF to stimulate PGE(2) release. Together, these results suggest that PKC is required for PGE(2) receptor signaling to PKA activation. Depletion of calcium from intracellular stores blocked PDGF-induced arachidonic acid release, PGE(2) release, and the subsequent PKA activation in hSMC without inhibiting MAPK activation. Further, no effect on PGE(2)-stimulated PKA activation was seen, suggesting that inhibition of PDGF-induced PKA activation by intracellular calcium depletion is due to inhibition of cPLA(2) activation. In addition, transient increases in cAMP synthesis were observed in hSMC when intracellular calcium levels were increased dramatically by a short stimulation with thapsigargin or by sphingosine-1-phosphate(51) , for example. This effect was independent of MAPK activation, and the concentrations of intracellular calcium required for direct cAMP stimulation were higher than those obtained following stimulation with PDGF or IGF-I (51) . (^4)Possibly, high intracellular concentrations of calcium may directly stimulate the adenylate cyclase types I and III (reviewed in (35) ).

Previously, we reported that PKA can inhibit PDGF-stimulated MAPK signaling in hSMC(22) . Therefore, we examined whether the increase in cAMP and PKA activity in response to PDGF could inhibit the MAP kinase cascade in a ``feedback'' manner. Such a negative feedback mechanism on MAPK (Erk2) activity has recently been demonstrated by Pyne and co-workers following bradykinin-induced cAMP accumulation(52) . We were unable to find an effect of the PDGF-stimulated PKA activity on the time course of MAPKK or MAPK activation in response to PDGF, nor was the PDGF-stimulated increase in PKA activity sufficient to limit the magnitude of MAPK activation by this growth factor. At this time, we can only speculate that the PDGF-stimulated increase in PKA activity occurs too transiently to sufficiently impede the activation of MAPK in hSMC. Alternatively, the PDGF-stimulated increase in cAMP and PKA activity may be involved in PKA-mediated transcription or cytoskeletal remodeling events such as actin filament reorganization which is known to occur in response to PDGF(51) .


FOOTNOTES

*
The study was supported in part by National Institutes of Health Grant DK 42528, the International Human Frontier Science Program, a Pilot Project grant from the W. M. Keck Center for Advanced Studies of Neural Signaling at the University of Washington, a grant from the Muscular Dystrophy Association (to E. G. K.), NHLBI Grants HL-18645 and HL-03174 from the National Institutes of Health (to R. R.), Grant HL34303 (to C. C. L.), and a grant from the American Heart Association (Washington Affiliate) (to L. M. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Current address: University of North Carolina at Chapel Hill, Chapel Hill, NC 27599-7365.

To whom correspondence should be addressed. Tel.: 206-543-8500; Fax: 206-543-0858.

(^1)
The abbreviations used are: PDGF, platelet-derived growth factor; EGF, epidermal growth factor; IBMX, 3-isobutyl-1-methylxanthine; MAPK, mitogen-activated protein kinase; MAPKK, MAPK kinase; IGF-I, insulin-like growth factor I; MBP, myelin basic protein; hSMC, human arterial smooth muscle cells; cPLA(2), cytosolic phospholipase A(2); PMA, phorbol 12-myristate 13-acetate; DMEM, Dulbecco's modified Eagle's medium; PDS, plasma-derived serum; PAGE, polyacrylamide gel electrophoresis; PGE(2), prostaglandin E(2); PKA, cAMP-dependent protein kinase; PKC, protein kinase C.

(^2)
K. E. Bornfeldt, unpublished observations.

(^3)
L. M. Graves, unpublished observations.

(^4)
L. M. Graves, K. E. Bornfeldt, J. S. Sidhu, G. M. Argast, E. W. Raines, R. Ross, C. C. Leslie, and E. G. Krebs, unpublished observations.


ACKNOWLEDGEMENTS

We are grateful to Dr. Michael Gelb and the members of his laboratory for providing recombinant cPLA(2), AACOCF(3), and AACOCH(3) and technical advice. We also acknowledge Dr. Daniel Storm and Dr. Joseph Beavo for critical evaluation of this work. We thank Melanie Cobb (University of Texas) for providing recombinant Erk2. We thank Kris Carroll and Alan Jennings for the preparation of the figures and for graphic assistance. The excellent technical assistance of Li-Chuan Huang and Karen Engel is gratefully acknowledged.


REFERENCES

  1. Ross, R. (1993) Nature 362, 801-809 [CrossRef][Medline] [Order article via Infotrieve]
  2. Crews, C. M., Alessandrini, A., and Erikson, R. L. (1992) Science 258, 478-480 [Medline] [Order article via Infotrieve]
  3. Feig, L. A., and Schaffhausen, B. (1994) Nature 370, 508-509 [CrossRef][Medline] [Order article via Infotrieve]
  4. Cowley, S., Paterson, H., Kemp, P., and Marshall, C. J. (1994) Cell 77, 841-852 [Medline] [Order article via Infotrieve]
  5. Seger, R., Seger, D., Reszka, A. A., Munar, E. S., Eldar-Finkelman, H., Dobrowolska, G., Jensen, A. M., Campbell, J. S., Fischer, E. H., and Krebs, E. G. (1994) J. Biol. Chem. 269, 25699-25709 [Abstract/Free Full Text]
  6. Mansour, S. J., Matten, W. T., Hermann, A. S., Candia, J. M., Rong, S., Fukasawa, K., Vande Woude, G. F., and Ahn, N. G. (1994) Science 265, 966-970 [Medline] [Order article via Infotrieve]
  7. Davis, R. J. (1993) J. Biol. Chem. 268, 14553-14556 [Free Full Text]
  8. Dennis, E. A. (1994) J. Biol. Chem. 269, 13057-13060 [Free Full Text]
  9. Kramer, R. M. (1995) in Signal-activated Phospholipases (Liscovitch, M., ed) pp. 13-30, R. G. Landes Co., Austin, TX
  10. Glaser, K. B., Mobilio, D., Chang, J. Y., and Senko, N. (1993) Trends Pharmacol. Sci. 14, 92-98 [CrossRef][Medline] [Order article via Infotrieve]
  11. Gelb, M. H., Jain, M. K., and Berg, O. G. (1994) FASEB J. 8, 916-924 [Abstract/Free Full Text]
  12. Channon, J. Y., and Leslie, C. C. (1990) J. Biol. Chem. 265, 5409-5413 [Abstract/Free Full Text]
  13. Lin, L. L., Wartmann, M., Lin, A. Y., Knopf, J. L., Seth, A., and Davis, R. J. (1993) Cell 72, 269-278 [Medline] [Order article via Infotrieve]
  14. Nemenoff, R. A., Winitz, S., Qian, N.-X., Van Putten, V., Johnson, G. L., and Heasly, L. E. (1993) J. Biol. Chem. 268, 1960-1964 [Abstract/Free Full Text]
  15. Qui, Z. H., and Leslie, C. C. (1994) J. Biol. Chem. 269, 19480-19487 [Abstract/Free Full Text]
  16. Miyake, R., and Gross, R. W. (1992) Biochim. Biophys. Acta 1165, 167-176 [Medline] [Order article via Infotrieve]
  17. Taussig, R., and Gilman, A. G. (1995) J. Biol. Chem. 270, 1-4 [Free Full Text]
  18. Rozengurt, E., Stroobant, P., Waterfield, M. D., Deuel, T. F., and Keehan, M. (1983) Cell 34, 265-272 [Medline] [Order article via Infotrieve]
  19. Nair, B. G., Rashed, H. M., and Patel, T. B. (1989) Biochem. J. 264, 563-571 [Medline] [Order article via Infotrieve]
  20. Ball, R. L., Tanner, K. D., and Carpenter, G. (1990) J. Biol. Chem. 265, 12836-12845 [Abstract/Free Full Text]
  21. Loesberg, C., Van Wijk, R., Zandenbergen, J., Van Aken, W. G., Van Mourik, J. A., and De Groot, P. G. (1985) Exp. Cell. Res. 160, 117-125 [Medline] [Order article via Infotrieve]
  22. Graves, L. M., Bornfeldt, K. E., Raines, E. W., Potts, B. C., Macdonald, S. G., Ross, R., and Krebs, E. G. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10300-10304 [Abstract]
  23. Wu, J., Dent, P., Jelinek, T., Wolfman, A., Weber, M. J., and Sturgill, T. W. (1993) Science 262, 1065-1069 [Medline] [Order article via Infotrieve]
  24. Cook, S. J., and McCormick, F. (1993) Science 262, 1069-1072 [Medline] [Order article via Infotrieve]
  25. Sevetson, B. R., Kong, X., and Lawrence, J. C. J. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10305-10309 [Abstract]
  26. Burgering, B. M., Pronk, G. J., van Weeren, P. C., Chardin, P., and Bos, J. L. (1993) EMBO J. 12, 4211-4220 [Abstract]
  27. Hafner, S. A, Adler, H. S., Mischak, H., Janosch, P., Heidecker, G., Wolfman, A., Pippig, S., Lohse, M., Ueffing, M., and Kolch, W. (1994) Mol. Cell. Biol. 14, 6696-6703 [Abstract]
  28. Burgering, B. M., and Bos, J. L. (1995) Trends Biochem. Sci. 20, 18-22 [CrossRef][Medline] [Order article via Infotrieve]
  29. Walsh, D. A., and Glass, D. B. (1991) Methods Enzymol. 201, 304-316 [Medline] [Order article via Infotrieve]
  30. Kemp, B. E., Graves, D. J., Benjamini, E., and Krebs, E. G. (1977) J. Biol. Chem. 252, 4888-4894 [Medline] [Order article via Infotrieve]
  31. deCarvalho, M. S., McCormack, F. X., and Leslie, C. C. (1993) Arch. Biochem. Biophys. 306, 534-540 [CrossRef][Medline] [Order article via Infotrieve]
  32. Bornfeldt, K. E., Raines, E. W., Nakano, T., Graves, L. M., Krebs, E. G., and Ross, R. (1994) J. Clin. Invest. 93, 1266-1274 [Medline] [Order article via Infotrieve]
  33. Corbin, J. D. (1983) Methods Enzymol. 99, 227-228 [Medline] [Order article via Infotrieve]
  34. Domin, J., and Rozengurt, E. (1993) J. Biol. Chem. 268, 8927-8934 [Abstract/Free Full Text]
  35. Choi, E. J., Xia, Z., Villacres, E. C., and Storm, D. R. (1993) Curr. Opin. Cell Biol. 5, 269-273 [Medline] [Order article via Infotrieve]
  36. Iyengar, R. (1993) FASEB J. 7, 768-775 [Abstract/Free Full Text]
  37. Thastrup, O., Cullen, P. J., Drobak, B. K., Hanley, M. R., and Dawson, A. P. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 2466-2470 [Abstract]
  38. Schror, K. (1993) Eur. Heart J. 14, Suppl. I, 34-41 [Abstract]
  39. Street, I. P., Lin, H. K., Lalibert'e, F., Ghomashchi, F., Wang, Z., Perrier, H., Tremblay, N. M., Huang, Z., Weech, P. K., and Gelb, M. H. (1993) Biochemistry 32, 5935-5940 [Medline] [Order article via Infotrieve]
  40. Bartoli, F., Lin, H. K., Ghomashchi, F., Gelb, M. H., Jain, M. K., and Apitz-Castro, R. (1994) J. Biol. Chem. 269, 15625-15630 [Abstract/Free Full Text]
  41. Claesson-Welsh, L. (1994) J. Biol. Chem. 269, 32023-32026 [Free Full Text]
  42. Vegesna, R. V. K., and Diamond, J. (1986) Eur. J. Pharmacol. 128, 25-31 [Medline] [Order article via Infotrieve]
  43. Rozengurt, E. (1981) Adv. Cyclic Nucleotide Res. 14, 429-442 [Medline] [Order article via Infotrieve]
  44. Nair, B. G., Parikh, B., Milligan, G., and Patel, T. B. (1990) J. Biol. Chem. 265, 21317-21322 [Abstract/Free Full Text]
  45. Budnik, L. T., and Mukhopadhyay, A. K. (1991) J. Biol. Chem. 266, 13908-13913 [Abstract/Free Full Text]
  46. Hart, M. J., Polakis, P. G., Evans, T., and Cerione, R. A. (1990) J. Biol. Chem. 265, 5990-6001 [Abstract/Free Full Text]
  47. Sa, G., Murugesan, G., Jaye, M., Ivashchenko, Y., and Fox, P. L. (1995) J. Biol. Chem. 270, 2360-2366 [Abstract/Free Full Text]
  48. Kawabe, J., Iwami, G., Ebina, T., Ohno, S., Katada, T., Ueda, Y., Homcy, C. J., and Ishikawa, Y. (1994) J. Biol. Chem. 269, 16554-16558 [Abstract/Free Full Text]
  49. Choi, E. J., Wong, S. T., Dittman, A. H., and Storm, D. R. (1993) Biochemistry 32, 1891-1894 [Medline] [Order article via Infotrieve]
  50. Jacobowitz, O., Chen, J., Premont, R. T., and Iyengar, R. (1993) J. Biol. Chem. 268, 3829-3832 [Abstract/Free Full Text]
  51. Bornfeldt, K. E., Graves, L. M., Raines, E. W., Igarishi, Y., Wayman, G., Yamamura, S., Yatomi, Y., Sidhu, J. S., Krebs, E. G., Hakomori, S., and Ross, R. (1995) J. Cell Biol. 130, 193-206 [Abstract]
  52. Pyne, N. J., Moughal, N., Stevens, P. A., Tolan, D., and Pyne, S. (1994) Biochem. J. 304, 611-616 [Medline] [Order article via Infotrieve]

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