Inhibition of PDGF-Mediated Proliferation of Vascular Smooth Muscle Cells by Calcium Antagonists
Background and Purpose The mechanism by which calcium antagonists (CAs) inhibit proliferation in vascular smooth muscle cells (VSMCs) is not yet fully understood. We investigated the effects of four CAs (clentiazem, verapamil, diltiazem, and nifedipine) on signal transduction pathways activated by platelet-derived growth factor (PDGF). To determine these effects, the levels of inositol phosphates (IPs), protein kinase C (PKC), and the induction of the transcription factor activator protein-1 (AP-1) were measured.
Methods The mitogenic effect of PDGF on VSMCs was measured by [3H]thymidine incorporated into DNA. IP production was monitored by [3H]myo-inositol incorporation. PKC activation was determined by measurement of myristoylated, alanine-rich C kinase substrate (MARCKS) phosphorylation in digitonin-permeabilized VSMCs. The induction of AP-1 complex was detected by electrophoretic mobility shift assays.
Results Each CA significantly inhibited the [3H]thymidine incorporation into DNA in unstimulated cells. Similar significant decreases in [3H]thymidine incorporation by CAs were observed when cells were stimulated by rPDGF-BB. The phosphorylation of MARCKS mediated by rPDGF-BB was significantly reduced by each CA. Clentiazem and verapamil significantly reduced the expression of AP-1 induced by rPDGF-BB (P<.01, P<.05). Clentiazem also significantly reduced the expression of AP-1 induced by rPDGF-AB (P<.05).
Conclusions PDGF-mediated proliferation of VSMCs correlates with activation of PKC but not with induction of the AP-1 complexes. In addition, our results suggest that CAs block proliferation of VSMCs by inhibiting DNA synthesis, possibly via PKC.
The proliferation of VSMCs is a key event in the pathogenesis of atherosclerosis.1 PDGF is thought to play an important role in the proliferation of VSMCs as well as the formation of atherosclerotic lesions in vivo.1 2 3 These effects are believed to be transmitted via the PDGF receptor by modulation of mitogenesis, phosphoinositide turnover, and the induction of proto-oncogenes. PDGF has two known isoforms (A and B types), which are able to form AA and BB homodimers and AB heterodimers. Similarly, the two forms of PDGF receptor (α and β) are αα and ββ homodimers and αβ heterodimers. It has been reported that PDGF-B transcription is increased in human atherosclerotic plaques compared with normal arteries in vivo and has also been seen during proliferation of VSMCs in vitro.3 4 5 6 The homodimer PDGF-BB stimulates VSMC migration and intimal thickening in a rat carotid injury model and induces intimal hyperplasia in porcine arteries in vitro.7 8 Furthermore, rPDGF-BB mediates a variety of cell signals, including phosphorylation of PDGF receptors, activation of PLC-γ, phosphatidylinositide turnover, DAG production, and increases in intracellular Ca2+ ([Ca2+]i).9 10 In addition, PDGF has been shown to induce c-fos, which is a component of AP-1 complexes.11 12 The induction of AP-1 complex is thought to play an essential role in cell proliferation.13 14 Previous reports suggest that the CAs possess antiatherogenic properties.15 16 17 It has been shown that CAs inhibit the effects of PDGF-induced signal transduction, particularly that involving phosphatidylinositol turnover and PKC activation.18 19 However, the effect of CAs on PDGF-mediated proliferation of VSMCs is not well understood. In view of the potential ability of CAs to retard atherogenesis, the elucidation of antimitogenic mechanisms of CAs at the cellular level is of major interest. In this study, we investigated the inhibitory effects of CAs (including a new benzothiazepine derivative, clentiazem) on proliferation of VSMCs induced by PDGF. Specifically, we examined DNA synthesis, the activation of PKC, and the induction of AP-1 complexes.
Materials and Methods
Natural PDGF and rPDGF-AA, -BB, and -AB were purchased from Upstate Biotechnology Inc. Methyl [3H]thymidine, [3H]myo-inositol, [γ-32P]ATP, and [α-32P]dCTP were obtained from Amersham International. DMEM, FBS, staurosporine, verapamil, diltiazem, and nifedipine were purchased from Sigma. Poly(dI-dC) and Moloney murine leukemia virus reverse transcriptase were purchased from Pharmacia. Clentiazem (TA-3090) was obtained from Marion Laboratories Inc.
VSMCs from the rat thoracic aorta (A-10) were obtained from the American Type Culture Collection and cultured in DMEM supplemented with 10% FBS, penicillins, and streptomycin (each 100 mg/mL). Cells between the 15th and 18th passages were used in these experiments.
Measurement of DNA Synthesis
Mitogenesis was assayed by measurement of [3H]thymidine incorporation into DNA. Subconfluent (70% to 80%) VSMCs were starved with FBS-free DMEM containing 0.1% BSA and 0.1% glucose at 37°C for 24 hours. Test cultures were treated with rPDGF-AA (40 ng/mL), -BB (40 ng/mL), or -AB (40 ng/mL) and/or clentiazem (10−5 mol/L), verapamil (10−6 mol/L), diltiazem (10−6 mol/L), and nifedipine (10−6 mol/L). Control cultures received only FBS-free DMEM containing 0.1% BSA and glucose. Quiescent VSMCs were supplemented with 1 mCi/mL of [3H]thymidine, incubated for 24 hours at 37°C, washed with PBS and 5% trichloroacetic acid, then dissolved in 0.25N NaOH containing 0.1% SDS. The radioactivity in the cell lysates was then determined in an LS 3133 Beckman scintillation counter.
Confluent VSMCs were cultured for 48 hours in a medium free of myo-inositol that contained 10% dialyzed serum and 1 mCi/mL [3H]myo-inositol. The cells were incubated in PBS containing 10 mmol/L LiCl for 10 minutes. Cells were then treated with rPDGF-AA (10 ng/mL), rPDGF-BB (10 ng/mL), and/or clentiazem (10−5 mol/L) for 20 minutes. At the end of the incubation, cells were washed twice with PBS and treated with 5% trichloroacetic acid. The cells were scraped and pelletized by centrifugation. Cell were then dissolved in 0.1 mol/L NaOH, and cellular radioactivity and protein concentrations were determined. The supernatant solution was extracted with ether, and the aqueous phase was applied to an ion exchange column (Bio-Rad AG 1X8) to separate the IPs. The column was washed with 5 mmol/L myo-inositol until no radioactivity was detected in the eluent. This was followed by elution with 0.2, 0.5, 1.0, and 1.5 mol/L ammonium formate in 0.1 mol/L formic acid. The radioactivity of each inositol phosphate fraction (IP1, IP2, IP3, and IP4) was determined from the eluted solutions.
Phosphorylation of PKC Substrate
MARCKS was identified by a previously described method.20 Confluent and quiescent VSMCs were starved with FBS-free DMEM for 48 hours and then treated with a CA (10−5 mol/L) for 30 minutes before the addition of rPDGF-BB. Before the phosphorylation experiments were initiated, cells were washed twice with FBS-free DMEM followed by two washes with isotonic KCl salt solution (in mmol/L: KCl 120, NaCl 30, MgCl2 1, K2HPO4 1, sodium PIPES 10 [pH 7.0], EGTA 1, and CaCl2 0.037) at 37°C. Phosphorylation was initiated by replacement of the salt solution from the last wash with permeabilization medium containing 40 mmol/L digitonin and 10 mmol/L [γ-32P]ATP (0.2 to 1 Ci/mmol) in isotonic KCl solutions. The cells were then incubated at 37°C for 5 minutes with 200 nmol/L PDBu, 10 ng/mL rPDGF-BB, and/or CAs. The cells were immediately scraped off the dishes and heated to 80°C for 5 minutes in SDS-PAGE sample buffer. Samples were resolved by one-dimensional 10% SDS-PAGE. After electrophoresis, the gels were dried and exposed to Kodak XAR5 film at −70°C for autoradiography. The intensity of the 80-kD band corresponding to the migration of MARCKS was quantified by a Bio-Rad imaging densitometer (model GS-670).
Induction of AP-1 Complexes
Subconfluent VSMCs (90%) were deprived of serum for 24 hours. The cells were treated with a CA (10−5 mol/L) for 15 minutes before the addition of growth factors. rPDGF-AA, -BB, and -AB (10 ng/mL) were added, followed by incubation at 37°C for 45 minutes. The reaction was terminated by removal of the incubation medium followed by a washing with 5 mL ice-cold PBS.
Preparation of Nuclear Extracts
Nuclear extracts were prepared by a modification of the procedure described by Dignam et al.21 All steps were performed at 4°C. VSMCs were washed with ice-cold PBS and scraped into 5 mL PBS. Cells were sedimented by centrifugation (500g for 5 minutes), then resuspended in 5 mL hypotonic solution (in mmol/L: Tris-HCl 10 [pH 7.9], MgCl2 12.5, KCl 10, DTT 0.5, and PMSF 1) and allowed to swell on ice for 10 minutes. The cells were then homogenized by 20 strokes of a glass Dounce homogenizer using the tight pestle. The nuclei were sedimented by centrifugation at 1000g for 5 minutes and then resuspended in the nucleic resuspension buffer (in mmol/L: Tris-HCl 20 [pH 7.9], MgCl2 1.5, DTT 0.5, and PMSF 1, and 20% glycerol) followed by the addition of 4 mol/L KCl to a final concentration of 0.4 mol/L. The suspension was rocked gently for 30 minutes, then centrifuged at 13 000g for 15 minutes. The supernatant solution containing the nuclear proteins was stored at −70°C until assayed.
Electrophoretic Mobility Shift Assay
An AP-1 binding site containing oligonucleotide (5′-GATCTGTGACTCAGCGC GA-3′) and its complement (5′-GATCTCGCGCTGAGTCACA-3′) were hybridized and used for EMSAs. The hybridized oligonucleotides were radiolabeled with [α-32P]dCTP with Moloney murine leukemia virus reverse transcriptase. The assays were carried out essentially as described by Dash and Moore,22 with minor modifications. Nuclear extract (≈5 μg protein) was incubated in 20 μL binding buffer (10 mmol/L Tris-HCl [pH 7.9], 5 mmol/L MgCl2, 0.5 mmol/L DTT, 0.5% glycerol) containing 2 μg poly(dI-dC). The 32P-labeled AP-1 probe (0.5 ng) was added, and the mixture was incubated for 30 minutes at room temperature. The reaction mixture was loaded directly onto a preelectrophoresed 5% polyacrylamide gel in 0.25×TBE (25 mmol/L Trizma base, 25 mmol/L boric acid, 1 mmol/L EDTA). The protein-probe complexes were separated from the free probe by electrophoresis at 150 V for 1.5 hours. The gel was dried and analyzed by autoradiography. The intensities of the AP-1 bands were quantified as the integrated area of optical density value by a Bio-Rad imaging densitometer (model GS-650). EMSAs were repeated in three independent experiments. The optical density value of each band was converted into a percentage of that obtained for the band of the naive nuclear extract from VSMCs on the same film, and the resulting percent values, presented in the “Results,” were analyzed statistically. Oligonucleotide specificity was examined by competition assay, in which 50 or 100 ng unlabeled AP-1 oligomer or 50 or 100 ng of unrelated oligomer (5′-TGTCGAATGCAAATCAGAA-3′, 3′-ACAGCTTACGTTTAGTGATCTT-5′) containing the consensus sequence of octamer-1 (Promega) was added to the reaction mixture 20 minutes before the labeled probe was added.
Data are presented as mean±SD. Comparisons between groups were done with Student's t test for unpaired variables. A value of P<.05 was used to assign statistical significance.
Antimitogenic Effect of CAs on VSMCs
Under unstimulated conditions, the CAs clentiazem (10−5 mol/L), diltiazem (10−5 mol/L), verapamil (10−5 mol/L), and nifedipine (10−5 mol/L) each significantly inhibited [3H]thymidine incorporation, to 75.7±4.3%, 59.1±4.6%, 40.1±0.3%, and 57.1±3.2% of control in VSMCs, respectively (P<.005, P<.001, P<.0001, and P<.001, n=6). These inhibitory effects of the CAs were dose dependent (Fig 1⇓). Addition of the growth factor rPDGF-BB (40 ng/mL) increased thymidine incorporation to 143.8±28% of control. The rPDGF-BB–induced thymidine incorporation was significantly decreased by clentiazem (10−5 mol/L) to 106.6±13.0%, diltiazem (10−6 mol/L) to 82.0±7.6%, verapamil (10−6 mol/L) to 93.3±7.5%, and nifedipine (10−6 mol/L) to 95.6±1.6% (P<.05, P<.01, P<.01, and P<.01, respectively, n=6) (Fig 2⇓). In contrast to rPDGF-BB, rPDGF-AA (40 ng/mL) and -AB (40 ng/mL) failed to stimulate thymidine incorporation.
Effect of Clentiazem on Production of IPs
Natural PDGF (at 40 ng/mL) significantly stimulated the production of total IPs (IP1, IP2, IP3, and IP4) to 28 190±4200 cpm/mg protein (227% of control level). Specifically, natural PDGF significantly stimulated the production of IP3 to 2120±780 cpm/mg protein (177%). Clentiazem had no significant effect on the PDGF-induced production of IPs (24 270±6960 cpm/mg protein) and PGDF-induced production of IP3 (1990±470 cpm/mg protein) in VSMCs. rPDGF-BB (10 ng/mL) also significantly stimulated the production of total IPs to 7240±1540 cpm/mg protein (176% of control) (P<.01, n=3). To specifically investigate the production of the intracellular IP3, it was found that rPDGF-BB stimulated the production of IP3 significantly, to 480±110 cpm/mg protein (133% of control) (P<.05). Clentiazem (10−5 mol/L) failed to inhibit the production of IP3 induced by rPDGF-BB (430±40 cpm/mg protein). Moreover, rPDGF-AA (10 ng/mL) failed to stimulate the production of IPs.
Effect of CAs on Phosphorylation of MARCKS
PDBu (200 nmol/L) markedly stimulated the phosphorylation of the 80-kD PKC-specific substrate protein (MARCKS) in digitonin-permeabilized VSMCs to 240±13.3% of control. Moreover, rPDGF-BB (40 ng/mL) increased the phosphorylation of MARCKS to 275±12.5% of control. Each CA at 10−5 mol/L significantly reduced rPDGF-BB–mediated phosphorylation of MARCKS (191.6±12.5% for clentiazem, 162.5±12.8% for diltiazem, 154.2±12.5% for verapamil, and 129.2±8.3% for nifedipine; P<.05, P<.02, P<.02, and P<.01, respectively; n=3). The PKC inhibitor staurosporine (20 μmol/L) completely inhibited the phosphorylation of MARCKS mediated by PDBu and rPDGF-BB.
Effect of CAs on AP-1 Expression
EMSAs were carried out to determine the amount of AP-1 complexes induced by PDGF. Low levels of AP-1 binding were detected in unstimulated cells (Fig 3⇓). Densitometry of the EMSAs showed that rPDGF-AA (10 ng/mL) significantly increased AP-1 levels to 186.1±36.7% (P<.05, n=3), rPDGF-BB (10 ng/mL) to 181.8±22.3% (P<.001, n=3), and rPDGF-AB (10 ng/mL) to 168.2±16.7% (P<.05, n=3) of control (100±28.4%, n=3). The induction of AP-1 complex by rPDGF-BB and rPDGF-AB were reduced by clentiazem (10−5 mol/L). On average, clentiazem significantly reduced rPDGF-BB–induced AP-1 levels (rPDGF-BB, 181.8±22.3% versus rPDGF-BB+clentiazem, 91.8±13.6%, P<.01; n=3) and rPDGF-AB–induced AP-1 levels (rPDGF-AB, 168.2±16.7% versus rPDGF-AB+clentiazem, 118.2±12.7%, P<.05; n=3). Clentiazem did not reduce rPDGF-AA–induced AP-1 levels (Fig 3⇓). Clentiazem significantly reduced the induction of AP-1 complexes by rPDGF-BB (10 ng/mL) in a dose-dependent manner (rPDGF-BB, 187±15.4% versus rPDGF-BB+clentiazem 10−5 mol/L, 128±15.0%, P<.05; n=3; rPDGF-BB versus rPDGF-BB+clentiazem 10−6 mol/L, 141±17.6%, P<.05; n=3; rPDGF-BB versus rPDGF-BB+clentiazem 10−7 mol/L, 177±24.6%, P=NS; n=3) (data not shown). The induction of AP-1 complex by rPDGF-BB was also significantly reduced by verapamil 10−6 mol/L (rPDGF-BB, 182±18.4% versus rPDGF-BB+verapamil, 126±16.5%, P<.05; n=3). Diltiazem 10−6 mol/L and nifedipine 10−6 mol/L each had no significant inhibitory effect on the rPDGF-BB–induced AP-1 expression (Fig 4A⇓). Fig 4B⇓ shows the gel retardation with the nuclear extract prepared from VSMCs treated by rPDGF-BB. Excess octamer-1, an unrelated oligonucleotide, had no effect on the AP-1 binding. The unlabeled AP-1 (100 ng) abolished the binding activity in the nuclear extract from VSMCs, indicating the specific binding for the AP-1 sequence.
In this report, we present data to help elucidate the underlying biochemical mechanisms by which PDGF may induce proliferation of VSMCs. Moreover, we also examined the effects of CAs on these biochemical changes activated by PDGF. Using thymidine incorporation into DNA as a marker for mitogenesis, we investigated the effects of CAs on both basal and PDGF-stimulated VSMC proliferation. All four CAs tested significantly inhibited thymidine incorporation in unstimulated cells in a dose-dependent manner. These findings are in agreement with previous reports that entry of calcium via VOCCs is necessary for cell division.22 23 24 25 Exposure of VSMCs to rPDGF-BB but not rPDGF-AA or rPDGF-AB significantly increased thymidine incorporation. It has been postulated that rPDGF-BB binds two β receptors.26 Therefore, the increase in thymidine incorporation we observe is consistent with previous reports that PDGF-ββ receptors mediate mitogenesis.9 10 27 The mechanism by which PDGF-ββ receptors induce mitogenesis is thought be linked to its ability to mobilize calcium. Consistent with this idea, when CAs were added in the presence of rPDGF-BB, the incorporation of thymidine was significantly reduced.
The increase in intracellular calcium induced by rPDGF-BB involves the influx of calcium ions through VOCCs as well as the release of internal calcium from IP3-sensitive endoplasmic reticulum stores.9 10 27 PLC-γ cleaves phosphatidylinositol to release DAG and IP3. When rPDGF-BB was added to VSMCs, IP3 production was significantly enhanced. The new calcium channel antagonist clentiazem was unable to decrease this enhanced IP production. This suggests that in VSMCs, the production of IPs is most likely a result of direct activation of PLC-γ by PDGF-ββ receptors, rather than activation of PLC-γ via an influx of calcium through VOCCs. As a consequence of PDGF-β receptor activation, in addition to IP3 production, the generation of DAG is increased. Both DAG production and the influx of calcium via VOCCs or release of calcium from IP3-sensitive internal stores could result in PKC activation. Exposure of VSMCs to rPDGF-BB resulted in increased phosphorylation of MARCKS. The degree of phosphorylation of MARCKS was significantly reduced by CAs. CAs are thought to bind the α-subunits of VOCC, thus inhibiting the transmembrane flux of Ca2+.28 29 30 31 This suggests that the influx of calcium through VOCCs is a major factor in PKC activation and MARCKS phosphorylation after rPDGF-BB application.
The apparent paradox of the inability of rPDGF-AA to provoke significant incorporation of labeled thymidine into VSMCs while nullifying the suppression of thymidine incorporation by CAs cannot be answered without further experimental investigations. However, the studies by Cadena and Gill32 and Hughes et al33 on the differential activation of the three PDGF dimeric receptors may provide a possible explanation. Each of the PDGF isoforms activates the several receptors, αα, αβ, or ββ; with autophosphorylation on tyrosine residues, activation of second messenger systems occurs, such as PLC-γ, phosphatidylinositol-3 kinase, and ras p21, to provoke proliferation, as measured by labeled thymidine incorporation, although thymidine incorporation with rPDGF-AA is substantially less than that of both AB and BB. In our study, the inability of rPDGF-AA to reduce a significant increase in DNA synthesis might be explained by this less effective mobilization of calcium through VOCCs. Since a parallel signal pathway exists, however, the absence of the suppressive activity of CAs on DNA synthesis may reflect the unopposed activity of the parallel system. This interpretation does not adequately explain why no significant increase in thymidine incorporation occurred with rPDGF-AA; further experiments are required to clarify the exact mechanisms of this apparent paradox.
PKC activation can cause increased expression of TRE containing the AP-1 binding site via the AP-1 complex. Activation of TRE-containing genes is thought to be involved in cellular differentiation and proliferation.14 Clentiazem and verapamil inhibited AP-1 induction by rPDGF-BB, with no significant effect on AP-1 induction by rPDGF-AA. This binding appears to be specific, since molar excess of unlabeled AP-1 binding site–containing oligonucleotide has been reported.34 35 In addition, this indicates that AP-1 induction by rPDGF-AA is via a pathway other than calcium influx through VOCCs. Increased thymidine incorporation, IP production, and MARCKS phosphorylation was enhanced by exposure of VSMCs to rPDGF-BB but not by exposure to rPDGF-AA or rPDGF-AB. However, all three growth factors were able to induce AP-1 binding. This suggests that IP3 production and PKC activation are linked to VSMC proliferation, but although AP-1 induction may be necessary, it is not sufficient to cause cellular proliferation.
rPDGF-BB is a potent mitogen for cell proliferation and may have an important role in myointimal hyperplasia.1 2 3 7 8 Clentiazem produced consistent inhibitory effects on rPDGF-BB–induced MARCKS phosphorylation, AP-1 expression, and DNA synthesis. Therefore, this benzothiazepine derivative may inhibit cellular proliferation via its effects on PKC-mediated transcription. Interestingly, we find that several isoenzymes of PKC (α, δ, ε, and ζ) are highly expressed in rat VSMCs and may be the gene product candidate responsible for PDGF-induced proliferation. Of the various CAs tested, the stimulatory effect of the ζ enzyme is primarily inhibited by clentiazem.36 Because CAs, including clentiazem, could be beneficial in preventing the formation of the atherosclerotic lesion, especially with respect to myointimal hyperplasia of VSMCs, their inhibitory effects on the rPDGF-BB–induced signal transduction pathway, including the expression of transcription factor AP-1, may help in preventing the formation of atheromatous plaques.
Selected Abbreviations and Acronyms
|EMSA||=||electrophoretic mobility shift assay|
|MARCKS||=||myristoylated, alanine-rich C kinase substrate|
|PDGF||=||platelet-derived growth factor|
|PKC||=||protein kinase C|
|rPDGF||=||recombinant platelet-derived growth factor|
|TRE||=||tetradecanoyl phorbol acetate response element|
|VOCC||=||voltage-operated calcium channel|
|VSMC||=||vascular smooth muscle cell|
This research was supported by the Clayton Foundation Research Fund, Houston, Tex; the Kanazawa Medical University Overseas Research Fund; and the Klingenstein Foundation (MH 49962). We thank Marion Laboratories Inc (Kansas City) for providing clentiazem (TA-3090). The authors wish to thank Maureen Casey and Elaine Marsh for their excellent secretarial assistance. Also, we thank Anthony Moore and Georgina Bui for their excellent technical assistance.
- Received November 8, 1995.
- Revision received October 29, 1996.
- Accepted October 30, 1996.
- Copyright © 1997 by American Heart Association
Williams LT. Signal transduction by the platelet-derived growth factor receptor. Science.. 1989;243:1564-1570.
Barrett TB, Benditt EP. sis (platelet-derived growth factor B chain) gene transcript levels are elevated in human atherosclerotic lesions compared to normal artery. Proc Natl Acad Sci U S A.. 1987;84:1099-1103.
Barrett TB, Benditt EP. Platelet-derived growth factor gene expression in human atherosclerotic plaques and normal artery wall. Proc Natl Acad Sci U S A.. 1988;85:2810-2814.
Majesky MW, Benditt EP, Schwartz SM. Expression and developmental control of platelet-derived growth factor A-chain and B-chain/Sis genes in rat aortic smooth muscle cells. Proc Natl Acad Sci U S A.. 1988;85:1524-1528.
Jawien A, Bowen-Pope DF, Lindner V, Schwartz SM, Clowes AW. Platelet-derived growth factor promotes smooth muscle migration and intimal thickening in a rat model of balloon angioplasty. J Clin Invest.. 1992;89:507-511.
Nabel EG, Yang Z, Liptay S, San H, Gordon D, Haudenschild CC, Nabel GJ. Recombinant platelet-derived growth factor B gene expression in porcine arteries induces intimal hyperplasia in vivo. J Clin Invest.. 1993;91:1822-1829.
Sachinidis A, Locher R, Vetter W, Tatje D, Hoppe J. Different effects of platelet-derived growth factor isoforms on rat vascular smooth muscle cells. J Biol Chem.. 1990;265:10238-10243.
Kondo T, Konishi F, Inui H, Inagami T. Differing signal transductions elicited by three isoforms of platelet-derived growth factor in vascular smooth muscle cells. J Biol Chem.. 1993;268:4458-4464.
Hall DJ, Stiles CD. Platelet-derived growth factor-inducible genes respond differentially to at least two distinct intracellular second messengers. J Biol Chem.. 1987;262:15302-15308.
Jackson LJ, Bush RC, Bowyer DE. Inhibitory effect of calcium antagonists on balloon catheter-induced arterial smooth muscle cell proliferation and lesion size. Atherosclerosis.. 1988;8:115-122.
Block LH, Emmons LR, Vogt E, Sachinidis A, Vetter W, Hoppe J. Ca2+-channel blockers inhibit the action of recombinant platelet-derived growth factor in vascular smooth muscle cells. Proc Natl Acad Sci U S A.. 1989;86:2388-2392.
Block LH, Keul R, Crabos M, Ziesche R, Roth M. Transcription activation of low density lipoprotein receptor gene by angiotensin-converting enzyme inhibitors and Ca2+-channel blockers involves protein kinase C isoforms. Proc Natl Acad Sci U S A.. 1993;90:4097-4101.
Erusalimsky JD, Friedberg I, Rozengurt E. Bombesin, diacylglycerols, and phorbol esters rapidly stimulate the phosphorylation of an Mr=80,000 protein kinase C substrate in permeabilized 3T3 cells. J Biol Chem.. 1988;263:19188-19194.
Dignam JD, Lebowitz RM, Roeder RG. Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res.. 1983;11:1475-1489.
Kuga T, Sadoshima J, Tomoike H, Kanaide H, Akaike N, Nakamura M. Action of Ca2+ antagonists on two types of Ca2+ channels in rat aorta smooth muscle cells in primary culture. Circ Res.. 1990;67:469-480.
Wang Z, Estacion M, Mordan LJ. Ca2+ influx via T-type channels modulates PDGF-induced replication of mouse fibroblasts. Am J Physiol. 1993;265(Cell Physiol 34):C1239-C1246.
Cirillo M, Quinn SJ, Romero JR, Canessa ML. Regulation of Ca2+ transport by platelet-derived growth factor-BB in rat vascular smooth muscle cells. Circ Res.. 1993;72:847-856.
Hart CE, Forstrom JW, Kelly JD, Seifert RA, Smith RA, Ross R, Murray MJ, Bowen-Pope DF. Two classes of PDGF receptor recognize different isoforms of PDGF. Science.. 1988;240:1529-1531.
Inui H, Kitami Y, Tani M, Kondo T, Inagami T. Differences in signal transduction between platelet-derived growth factor (PDGF) α and β receptors in vascular smooth muscle cells. J Biol Chem.. 1994;269:30546-30552.
Marks AR. Calcium channels expressed in vascular smooth muscle. Circulation. 1992;86(suppl III):III-61-III-67.
Mori T, Takai Y, Minakuchi R, Yu B, Nishizuka Y. Inhibitory action of chlorpromazine, dibucaine, and other phospholipid-interacting drugs on calcium-activated, phospholipid-dependent protein kinase. J Biol Chem.. 1980;255:8378-8380.
Hosey MM, Borsotto M, Lazdunski M. Phosphorylation and dephosphorylation and dihydropyridine-sensitive voltage-dependent Ca2+ channel in skeletal muscle membranes by cAMP- and Ca2+-dependent processes. Proc Natl Acad Sci U S A.. 1986;83:3733-3737.
Takahashi M, Seagar MJ, Jones JF, Reber BFX, Catterall WA. Subunit structure of dihydropyridine-sensitive calcium channels from skeletal muscle. Proc Natl Acad Sci U S A.. 1987;84:5478-5482.
Cadena DL, Gill GN. Receptor tyrosine kinase. FASEB J.. 1992;6:2332-2337.
Dash PK, Moore AN, Dixon CE. Spatial memory deficit, increased phosphorylation of the transcription factor CREB, and induction of the AP-1 complex following experimental brain injury. J Neurosci.. 1995;15:2030-2039.