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Am J Physiol Heart Circ Physiol 286: H2042-H2051, 2004. First published January 29, 2004; doi:10.1152/ajpheart.00625.2003
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Platelet-derived growth factor-BB and Ets-1 transcription factor negatively regulate transcription of multiple smooth muscle cell differentiation marker genes

Frédéric Dandré and Gary K. Owens

Department of Molecular Physiology and Biological Physics, University of Virginia, Charlottesville, Virginia 22903

Submitted 2 July 2003 ; accepted in final form 23 January 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Platelet-derived growth factor (PDGF)-BB, a potent mitogen for mesenchymal cells, also downregulates expression of multiple smooth muscle (SM) cell (SMC)-specific markers. However, there is conflicting evidence whether PDGF-BB represses SMC marker expression at a transcriptional or posttranscriptional level, and little is known regarding the mechanisms responsible for these effects. Results of the present studies provide clear evidence that PDGF-BB treatment strongly repressed SM {alpha}-actin, SM myosin heavy chain (MHC), and SM22{alpha} promoters in SMCs. Of major significance for resolving previous controversies in the field, we found PDGF-BB-induced repression of SMC marker gene promoters in subconfluent, but not postconfluent, cultures. Treatment of postconfluent SMCs with a tyrosine phosphatase inhibitor restored PDGF-BB-induced repression, whereas treatment of subconfluent SMCs with a tyrosine kinase blocker abolished PDGF-BB-induced repression, suggesting that a tyrosine phosphorylation event mediates cell density-dependent effects. On the basis of previous observations that Ets-1 transcription factor is upregulated within phenotypically modulated neointimal SMCs, we tested whether Ets-1 would repress SMC marker expression. Consistent with this hypothesis, results of cotransfection experiments indicated that Ets-1 overexpression reduced transcriptional activity of SMC marker promoter constructs in SMCs, whereas it increased activity of SM {alpha}-actin promoter in endothelial cells. PDGF-BB treatment increased expression of Ets-1 in cultured SMCs, and SM {alpha}-actin mRNA expression was reduced in multiple independent clones of SMCs stably transfected with an Ets-1-overexpressing construct. Taken together, results of these experiments provide novel insights regarding possible mechanisms whereby PDGF-BB and Ets-1 may contribute to SMC phenotypic switching associated with vascular injury.

vascular injury; promoter; cell density; signaling

SMOOTH MUSCLE (SM) cells (SMCs) are highly specialized cells expressing a unique repertoire of contractile proteins, ion channels, and signaling molecules necessary for their contractile function (38). The differentiation state of SMCs is controlled by a complex combination of local environmental cues such as cell-cell and cell-matrix interactions, neuronal influences, and hemodynamic and mechanical forces (38), as well as intrinsic factors such as clonal origins (6). In contrast to cardiac and skeletal muscle cells, which are terminally differentiated, SMCs can undergo major changes in phenotype, depending on physiological or pathological conditions. For example, in tissue culture or in vascular neointimal lesions, SMCs display diminished expression of a number of proteins that are characteristic of fully differentiated SMCs, including smooth muscle (SM) {alpha}-actin, SM myosin heavy chain (MHC), and SM22{alpha} (20, 42). The precise role of phenotypic modulation of SMCs in the etiology of atherosclerosis is unclear, although it is well established that dedifferentiated SMCs exhibit a number of properties, including enhanced migration, proliferation, increased production of metalloproteinases, and extracellular matrix synthesis, which are likely to play a key role in progression of vascular lesions (46). Nevertheless, the mechanisms and factors that control the process of phenotypic modulation of SMCs are very poorly understood.

One factor that has been shown to be highly efficacious in suppressing expression of SMC markers is PDGF-BB. Members of the PDGF family have been implicated in many different pathophysiological conditions, including vascular development, wound healing, neoplasia, and atherosclerosis (14). These molecules have major mitogenic and chemoattractant properties for SMCs and other mesenchymal cells such as fibroblasts and mesangial cells (14). Moreover, previous studies in our laboratory have indicated that PDGF-BB causes marked decreases in expression of multiple SMC differentiation markers, including SM {alpha}-actin, SM MHC, and {alpha}-tropomyosin in cultured SMCs (4, 8, 15). Decreased expression of these markers was not a direct function of the mitogenic effects of PDGF-BB, because the mitogenic response to 10% serum did not elicit a comparable response. In addition, chronic treatment of cultured SMCs with PDGF-BB resulted in nearly complete suppression of expression of SMC differentiation markers in the absence of a sustained mitogenic response (4). These changes were completely reversible, in that withdrawal of PDGF-BB resulted in reexpression of SM {alpha}-actin and SM MHC (4). Consistent with these findings, Hayashi et al. (13) subsequently showed that PDGF-BB treatment downregulated expression of h-caldesmon and calponin in cultured gizzard SMCs.

There is evidence, albeit controversial, that PDGF-BB-induced repression of SMC marker genes is mediated through transcriptional and posttranscriptional effects. For example, results of previous studies in our laboratory showed that PDGF-BB-induced decreases in expression of SM {alpha}-actin and SM MHC mRNAs were extremely rapid relative to the normal half-life of these transcripts (8). In contrast, PDGF-BB increased expression of nonmuscle {beta}-actin transcripts (7). These results therefore suggested that at least part of the effect of PDGF-BB was posttranscriptional through selective mRNA destabilization. In contrast, Van Putten et al. and Somasundaram et al. presented evidence based on transient transfection studies that PDGF-BB treatment of SMCs repressed the activity of the SM {alpha}-actin promoter (58) and the SM MHC promoter (54). However, these studies did not identify the specific cis-regulatory elements or trans-acting factors that mediated this effect. Moreover, a limitation of these studies was that they involved the analysis of a single SMC promoter and employed truncated SMC promoter constructs known to be insufficient for conferring appropriate expression of these genes in SMCs in vivo (29, 31).

Our laboratory and others previously defined the minimal regions of the SM {alpha}-actin, SM MHC, and SM22{alpha} genes in transgenic reporter systems that recapitulated expression of the endogenous genes throughout development and in adult mice (1, 17, 29, 31). Moreover, our studies of the effect of endothelial denudation on expression of reporter genes in these transgenic mice indicated that expression of these promoter constructs was repressed in response to vascular injury in vivo (42). Thus results indicate that the SMC promoter constructs that we previously described contain all the cis-regulatory elements necessary to transduce signals generated from physiological and pathological stimuli, such as vascular injury, into modulation of SMC marker expression.

Inasmuch as it is not clear whether PDGF-BB decreases expression of SMC markers at the transcriptional or posttranscriptional level, the initial goal of the present studies was to test the effect of PDGF-BB on activity of the SMC marker promoter constructs that have been characterized in transgenic mice. We found that expression of SM {alpha}-actin, SM MHC, and SM22{alpha} promoter-reporter constructs was markedly repressed after PDGF-BB stimulation in subconfluent SMCs but not in postconfluent SMC cultures. We subsequently initiated a series of experiments to determine the potential mechanisms whereby these promoters were inhibited by PDGF-BB and specifically tested the possible role of Ets-1, a transcription factor previously shown to be upregulated after PDGF-BB stimulation and during vascular injury (11, 16). We provide novel evidence that Ets-1 is a very potent and selective inhibitor of SMC-specific gene expression.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents. Recombinant PDGF-BB was purchased from Upstate (Waltham, MA) and resuspended at 10 µg/ml in vehicle [fatty acid-free bovine serum albumin (2 µg/ml) and 10 mM acetic acid]. AG-1295 was purchased from Calbiochem (San Diego, CA) and resuspended at 10 mM in DMSO. Sodium orthovanadate was purchased from Sigma and resuspended at 100 mM in water, boiled, and adjusted to pH 10. Plasmid DNA was prepared using endotoxin-free Qiafilter Plus plasmid kit (Operon, Qiagen).

Cell culture. SMCs were isolated from thoracic aortas of 6-wk-old Sprague-Dawley rats by collagenase-elastase digestion (1). Cells were cultured in DMEM (Invitrogen, Carlsbad, CA) supplemented with 10 mM HEPES buffer (Invitrogen) and 10% fetal bovine serum (Hyclone, Logan, UT). For starvation, SMCs were fed insulin serum-free medium [1:1 DMEM-Ham's F-12 (Invitrogen) supplemented with 10 mM HEPES, 6.25 ng/ml sodium selenite, 5 mg/ml transferrin, and 0.2 mM l-ascorbic acid] for 16–20 h. Bovine aortic endothelial cells (a gift from Dr. B. P. Helmke, University of Virginia) were cultured in DMEM supplemented with 10 mM HEPES buffer and 10% heat-inactivated calf serum (Invitrogen).

DNA constructs. The SM {alpha}-actin-luciferase plasmid was constructed by subcloning a SpeI/SalI rat SM {alpha}-actin fragment (–2555 to +2813) into NheI/XhoI-digested pGL3 basic vector (Promega, Madison, WI). The SM MHC-luciferase plasmid was constructed in two steps: a 40-bp fragment (5'-TAC TAA CTC AGC TGC TCC CTC TGT CTC TTC AGG CG-3') encoding an SM {alpha}-actin intron 1 splice acceptor was ligated with BglII/HindIII-digested pGL3-BV, and then a SalI/SalI rat SM MHC fragment (–4200 to +11600) was subcloned into XhoI-digested pGL3-BV. The SM22{alpha}-luciferase plasmid was constructed by subcloning a HindIII/SacI mouse SM22{alpha} fragment into HindIII/SacI-digested pGL3-BV. The aortic carboxypeptidase-like protein (ACLP)-luciferase plasmid was constructed by subcloning a HindIII/HindIII mouse ACLP fragment (–2500 to +186; provided by Dr. M. D. Layne, Harvard Medical School, Boston, MA) into HindIII-digested pGL3-BV. The c-fos-luciferase plasmid was constructed by subcloning a SalI/BamHI mouse c-fos fragment (–356 to +109) into XhoI/BglII-digested pGL3-BV. The –153 to +2813 SM {alpha}-actin-luciferase plasmid was constructed by deleting the –2555 to –153 region in the full-length SM {alpha}-actin-luciferase construct by KpnI/MscI digestion and religation. The –153 to +20 SM {alpha}-actin-luciferase plasmid was made by cloning the –153 to +20 fragment into KpnI/SmaI-digested pGL3-BV. All reporter constructs were end sequenced and checked by testing three different clones in transfection assays. The Ets-1 expression plasmid (provided by Drs. J. H. Ko and N. Taniguchi, Osaka, Japan) contains a neomycin resistance cassette and the human Ets-1 cDNA driven by the CAGGS promoter (19, 36).

Reporter assays. All transient transfection experiments were performed in triplicate using Superfect transfection reagent (Operon, Qiagen) as indicated by the manufacturer. Cells were incubated for 3 h in the presence of 2 µg of DNA per well and 10 µl of Superfect per well for six-well dishes or 10 µg of DNA per dish and 60 µl of Superfect for 100-mm dishes. Then cells were cultured as described for each experiment. Cells were harvested using Passive Lysis Buffer (Promega). Cell extracts were assayed for luciferase expression using Enhanced Luciferase Assay kit (Becton Dickinson, Franklin Lakes, NJ) and for protein amount using Coomassie Plus Protein Assay Reagent (Pierce, Rockford, IL) as indicated by the manufacturers. Relative promoter activities are expressed as luminescence relative units normalized for protein content in cell extracts. Results represent average data ± SD from a triplicate experiment. Results in Figs. 1– 3 and 5–8 are representative of at least three different experiments. Comparisons between treatment groups were analyzed by one-way ANOVA or Student's t-test. Differences between treatment groups for which P <= 0.05 were considered statistically significant (23).



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  		Fig. 1. Platelet-derived growth factor (PDGF)-BB repressed smooth muscle (SM) cell (SMC) marker promoter activity in subconfluent SMCs. SMCs were transfected with SM {alpha}-actin-luciferase, SM myosin heavy chain (MHC)-luciferase, SM22{alpha}-luciferase, aortic carboxypeptidase-like protein (ACLP)-luciferase, and c-fos-luciferase plasmids, serum starved, and then treated for 24 h with different concentrations of PDGF-BB. Promoter activities were normalized to that of cells transfected with promoterless construct and treated with PDGF vehicle. *Significantly different (P <= 0.05) from untreated cells (1-way ANOVA).

 


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  		Fig. 3. PDGF-{beta} receptor tyrosine kinase inhibition blocked PDGF-BB-induced repression of SM {alpha}-actin promoter activity. SMCs were transfected with SM {alpha}-actin-luciferase construct, serum starved, preincubated with different concentrations of the tyrosine kinase inhibitor AG-1295 for 40 min, and cotreated with PDGF-BB (30 ng/ml) and AG-1295 for 24 h (DMSO concentration was kept the same in all conditions). Promoter activities were normalized to that of cells treated with PDGF vehicle for each AG-1295 concentration. *Significantly different (P <= 0.05) from vehicle (paired t-test).

 


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  		Fig. 5. Overexpression of Ets-1 reduced activity of multiple SM promoter constructs in SMCs. SMCs were transfected with SM {alpha}-actin-luciferase, SM MHC-luciferase, SM22{alpha}-luciferase, ACLP-luciferase, or c-fos-luciferase constructs (1,000 ng/well) and Ets-1-expressing construct [phosphorylated Ets-1 (pEts-1) made up to 1,000 ng/well with the empty vector containing Ets-1 cDNA]. Cells were cultured for 40 h in insulin-free serum-free medium after transfection. Promoter activities were normalized for each construct to that of cells transfected with empty vector. *Significantly different (P <= 0.05) from control-transfected cells (1-way ANOVA).

 


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  		Fig. 8. Ets-1 overexpression decreased SM {alpha}-actin mRNA expression levels in stably transfected SMC clones. SMCs were transfected with the Ets-1-expressing plasmid carrying a neomycin resistance gene. Clones resistant to antibiotic treatment were isolated and propagated. Ets-1-transfected SMC clones were analyzed for expression of Ets-1 protein by Western blot (A) and SM {alpha}-actin, nonmuscle (NM) {beta}-actin, and Ets-1 mRNA by Northern blot (B) after culture for 48 h in insulin-free serum-free medium. Quantitative analysis of Northern blot was done using a PhosphorImager. C: results from a representative experiment show SM {alpha}-actin band intensity normalized to nonmuscle {beta}-actin band intensity.

 
Stable transfected clone preparation and analysis. SMCs were transfected using supercoiled Ets-1-expressing plasmid DNA. Stably transfected cells were selected using geneticin (Invitrogen; 200 µg/ml). Seven cell lines (A1–A4 and B1–B3) were recovered after cloning of the resistant cells. For characterization and analysis, cells were cultured in the absence of geneticin to avoid interference with experiments. Western blot analysis was performed as described elsewhere (18) using rabbit anti-human Ets-1 N-276 polyclonal antibody (Santa Cruz Biotechnology, Santa Cruz, CA). For Northern blot analysis, total cytoplasmic RNA was extracted, separated, and transferred to a nylon membrane as described previously (12). SM {alpha}-actin and nonmuscle {beta}-actin mRNA were detected with a 512-bp EcoRI fragment that encoded amino acids 202–374 of human skeletal muscle {alpha}-actin cDNA. Ets-1 mRNA was detected with a 324-bp SacI/EcoRI fragment that encoded amino acids 86–194 of human Ets-1 cDNA. cDNA probes were radiolabeled with [{alpha}-32P]dCTP by random priming. Blots were hybridized with probes in 6x SSC-2x Denhardt-0.1% SDS-0.1 mg/ml sperm DNA at 65°C overnight and washed with 0.1x SSC-0.1% SDS at 65°C for actins and with 0.5x SSC-0.05% SDS at 50°C for Ets-1. Signals were detected using PhosphorImager (Molecular Dynamics) after 27 h of exposition and quantified using Image Quant software (Molecular Dynamics).

RT-PCR analysis of RNA expression. SMCs were cultivated in 60-mm dishes. Then the cells were trypsinized, centrifuged, and dissolved in 150–400 µl of TRIzol reagent (Invitrogen). RNA extraction was performed according to the manufacturer's instructions. Two micrograms of total RNA were retrotranscribed in 40 µl final volume using Superscript II (Invitrogen) in the presence of RNase-Out RNase inhibitor (Invitrogen) and oligo(dT) oligonucleotide. cDNA was analyzed by PCR using 1 µl of cDNA template, AmpliTaq DNA polymerase (Perkin-Elmer), and oligonucleotides specific for each target gene (Operon, Qiagen). The linearity of RT-PCR amplification was checked by testing different numbers of cycles. The intensity of agarose gel-resolved PCR bands was quantified by densitometry using NIH Image software. The results of experiments shown in Figs. 4 and 9 were obtained using a number of cycles that resulted in exponential amplification of the PCR target. For ACLP, because only mouse and human cDNA sequences were available (24), we designed primers matching the mouse sequence, PCR-amplified rat cDNA, and purified and sequenced each DNA strand of the PCR product. We ended with a 564-bp rat cDNA sequence displaying 92% identity with the mouse cDNA sequence and designed PCR primers from that sequence for the RT-PCR analysis.



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  		Fig. 4. PDGF-BB treatment increased Ets-1 mRNA expression in SMCs. A: serum-starved SMCs were treated with PDGF-BB (30 ng/ml) or vehicle for 2, 4, 12, 24, or 36 h. All treatments were harvested at the same time. Total RNAs were analyzed by RT-PCR in linear amplification conditions for expression of Ets-1, SM {alpha}-actin, SM MHC, ACLP, and GAPDH mRNAs. B: serum-starved SMCs were treated with PDGF-BB (30 ng/ml) for 40 min or 2, 4, 8, or 24 h or with vehicle for 4 h. All treatments were harvested at the same time. Cell extracts were made, and proteins were analyzed by Western blot using an anti-Ets-1 antibody.

 


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  		Fig. 9. Ets-1 expression is upregulated after PDGF-BB treatment at low and high density. SMCs were plated at low (10,000 cells/cm2) or high (35,000 cells/cm2) density, cultured for 24 h in the presence of 10% serum, serum starved, and then treated with vehicle or PDGF-BB (30 ng/ml) for 2 or 24 h. Total RNAs were analyzed by RT-PCR in linear amplification conditions for expression of Ets-1, SM {alpha}-actin, and GAPDH mRNAs.

 

   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
PDGF-BB markedly repressed expression of multiple SMC promoter-luciferase constructs in cultured SMCs. We previously demonstrated that PDGF-BB markedly decreased expression of multiple SMC marker genes such as SM {alpha}-actin and SM MHC at the protein and mRNA levels (4, 8, 15). In addition, through a series of deletion studies in transgenic mice, we identified minimal regions of the SM {alpha}-actin (–2555 to +2813) and SM MHC (–4200 to +11600) genes required to recapitulate expression of these endogenous genes in vivo in transgenic mice (29, 31). These promoter-enhancer constructs contained regions that were required for expression in vivo but were absent from constructs used previously to test the possible effects of PDGF-BB on transcription (54, 58).

To determine whether reduced transcription contributes to PDGF-BB-induced repression of SMC marker genes, a series of transfection experiments were performed using luciferase reporter plasmids under the control of the –2555 to +2813 SM {alpha}-actin promoter enhancer, the –4200 to +11600 SM MHC promoter-enhancer construct, or the –447 to +47 SM22{alpha} promoter enhancer shown to drive expression in arterial SMCs in vivo in transgenic mice (17, 27). We also tested a –2500 to +186 mouse ACLP promoter construct that has previously been shown to drive high-level expression in SMCs and other non-SMC tissues in transgenic mice (25). The ACLP gene is activated in neointimal SMCs during vascular injury, suggesting that it may be induced by PDGF-BB. As a positive control, cells were transfected with a luciferase construct under control of the c-fos promoter (–356 to +109) (10), which is known to be activated by PDGF-BB (49). Results of these analyses indicated that PDGF-BB at 30 ng/ml repressed expression of SM {alpha}-actin, SM MHC, and SM22{alpha} promoter constructs by 60, 57, and 45%, respectively (Fig. 1). In contrast, PDGF-BB increased c-fos promoter activity under the same experimental conditions, whereas PDGF-BB had no effect on the ACLP promoter. PDGF-BB treatment also had no effect on expression from a simian virus 40 promoter-luciferase construct and a promoterless luciferase construct (data not shown). Taken together, these results provide clear evidence that PDGF-BB downregulates transcription of multiple SMC marker gene promoter-enhancer constructs and that effects were selective, in that PDGF-BB had no effect on transcription of several non-SMC genes.

PDGF-BB-induced repression of SM {alpha}-actin gene transcription was cell density dependent. As noted in the introduction, there is conflicting evidence whether PDGF BB-induced repression of SMC gene expression is mediated at the transcriptional (54, 58) or the posttranscriptional level (7). However, previous studies suggesting that PDGF-BB induced selective destabilization of mRNAs for SMC marker genes were done in postconfluent SMC cultures, whereas studies implicating decreased transcription were done in subconfluent cultures. Given that PDGF-BB responsiveness of cells has been found to vary as a function of cell density as a result of differential phosphorylation of the PDGF-{beta} receptor (55), we hypothesized that the underlying mechanisms whereby PDGF represses SMC gene expression differ as a function of cell density. Because transfection efficiencies are significantly decreased in confluent SMC cultures (unpublished data), these experiments necessarily required development of experimental protocols that allowed independent control of cell density yet also resulted in adequate efficiency of transfection. This was accomplished by subculturing cells at high or low cell density after transient transfection (Fig. 2A; see MATERIALS AND METHODS). Results clearly showed that PDGF-BB treatment had no effect on SM {alpha}-actin promoter activity when cells were plated at high density (Fig. 2B). In contrast, a decrease in promoter activity was observed after PDGF-BB treatment when cells were plated at low density. Similar results were observed using the SM MHC promoter-luciferase construct (data not shown). These results provide novel evidence that the repressive effect of PDGF-BB on transcription of the SM {alpha}-actin gene is cell density dependent.



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  		Fig. 2. PDGF-BB-induced repression was cell density dependent. SMCs were transfected with SM {alpha}-actin-luciferase plasmid, trypsinized, and seeded at low density (LD, 10,000 cells/cm2) or high density (HD, 35,000 cells/cm2) in the presence of 10% serum. On the next day, cells were serum starved and photographed at low magnification (A). Cells were then treated for 24 h with PDGF-BB (BB, 30 ng/ml) or vehicle (V) with or without 75 µM sodium orthovanadate (B). Promoter activities were normalized to that of cells transfected with promoterless construct and treated with PDGF vehicle. *Significantly different (P <= 0.05) from vehicle (paired t-test).

 
An increase in protein tyrosine phosphatase expression has previously been shown to be responsible for blocking PDGF-BB-induced autophosphorylation of the PDGF-{beta} receptor at high cell density (40, 55). Thus, to determine whether density-dependent increases in tyrosine phosphatases might contribute to loss of PDGF-BB-induced repression of SM {alpha}-actin in confluent SMCs, cells were treated with orthovanadate, a potent inhibitor of protein phosphatases, with or without PDGF-BB at low and high cell densities. Activity of the SM {alpha}-actin promoter was significantly repressed after treatment of high-density cultures with PDGF + orthovanadate (Fig. 2B), whereas at low density orthovanadate did not significantly affect PDGF-BB-induced repression of the SM {alpha}-actin promoter. The effect of orthovanadate at high density was dose dependent, with maximal effects at 75 µM (data not shown). Given that treatment with orthovanadate may induce nonspecific effects, we cannot conclude that induction of tyrosine phosphatase expression is responsible for the inhibitory effect of high cell density on the PDGF-BB response. Nevertheless, the finding that orthovanadate can rescue the response to PDGF-BB at high cell density, whereas it did not significantly affect the PDGF-BB response at low cell density, reinforces the observation that changes that prevent PDGF-BB-induced repression of SM {alpha}-actin gene expression occur at high cell density.

PDGF-{beta} receptor activation is required for PDGF-BB-induced repression of SM {alpha}-actin promoter. PDGF-BB has been shown to bind to PDGF-{alpha} and PDGF-{beta} receptors (14). To determine whether the repressive effect of PDGF-BB was dependent on PDGF-{beta} receptor phosphorylation, SMC cultures were pretreated with the tyrphostin AG-1295, a tyrosine kinase inhibitor that has been shown to be highly selective for the PDGF-{beta} receptor (21). AG-1295 has previously been shown to block PDGF-BB-induced autophosphorylation of the PDGF-{beta} receptor (2, 21, 48). AG-1295 was used at 0.5–10 µM [AG-1295 IC50 for PDGF-BB-induced DNA synthesis = 2.5 µM (21)]. At 2.5–10 µM, AG-1295 blocked PDGF-BB-induced repression of SM {alpha}-actin promoter activity, indicating that PDGF-BB-induced repression of SM {alpha}-actin promoter activity in subconfluent cultures was PDGF-{beta} receptor dependent (Fig. 3). In contrast, AG-1478, a tyrosine kinase inhibitor showing selectivity for the epidermal growth factor receptor, had no effect on the basal promoter activity or the response to PDGF-BB (data not shown). Interestingly, 5–10 µM AG-1478 increased SM {alpha}-actin promoter activity after PDGF-BB treatment. This observation suggests that 1) in the absence of PDGF-{beta} receptor tyrosine kinase activity, PDGF-BB may activate a subset of signaling pathways that lead to activation of SM promoters; 2) there is residual AG-1295-insensitive PDGF-{beta} receptor tyrosine kinase activity that can result in activation of SMC promoters; and/or 3) activation of non-PDGF-{beta} receptor pathways (e.g., PDGF-{alpha} receptors) may activate (rather than repress) SMC gene expression (see discussion). Taken together, these observations indicate that PDGF-BB-mediated repression of SM {alpha}-actin is dependent on PDGF-{beta} receptor activation but that PDGF-BB may induce differential effects on SMC gene expression through alternative signaling pathways.

Ets-1 transcription factor overexpression repressed SMC marker promoters. Several reports have shown that Ets-1 transcription factor expression is upregulated in PDGF-BB-stimulated SMCs and in neointimal SMCs in a rat model of carotid balloon injury (11, 16, 35). Given these observations, we hypothesized that Ets-1 may mediate PDGF-BB-induced transcriptional repression of SMC marker gene promoters. As an initial test of the possible role of endogenous Ets-1 in mediating the effects of PDGF-BB, we investigated the time course of expression of Ets-1 compared with that of SM {alpha}-actin, SM MHC, and ACLP in SMCs by RT-PCR (Fig. 4). Ets-1 mRNA and protein levels were upregulated within 2 h of treatment with PDGF-BB and remained increased at 4 h. By 8 h, Ets-1 returned to a basal level. SM {alpha}-actin and SM MHC mRNA levels started to decrease by 12 h of treatment, reached a minimum by 24 h, and began to return to pretreatment levels by 36 h. ACLP mRNA levels were not affected by PDGF-BB treatment. The effects of PDGF-BB on these three genes therefore were correlated with the effects of PDGF-BB on the promoter-reporter constructs. As reported previously, PDGF-BB also induced a slight increase in GAPDH mRNA levels (41).

To determine whether Ets-1 had the potential to modulate SMC promoter activity, we examined the effect of Ets-1 overexpression in cultured SMCs. Rat aortic SMCs were transfected with SMC reporter constructs and a plasmid construct expressing Ets-1. Results showed that –2555 to +2813 SM {alpha}-actin and –4200 to +11600 SM MHC promoters were strongly repressed by Ets-1 overexpression in a dose-dependent manner (Fig. 5). SM22{alpha} promoter activity was also significantly repressed. In contrast, activity of the ACLP promoter was not influenced by Ets-1 overexpression, whereas the c-fos promoter was activated. These results showed that Ets-1 induced the same pattern of changes in SMC gene expression that was induced by PDGF-BB.

To delineate the minimal region of the SM {alpha}-actin promoter that could mediate PDGF-BB- and Ets-1-induced repression, a series of deletion mutants of the SM {alpha}-actin promoter were constructed and tested in reporter assays. The –153 to +20 sequence contains the CArG-A and CArG-B and the TCE cis-regulatory elements and was shown in our laboratory to be the shortest sequence that retains transcriptional activity in cultured SMCs. The –153 to +2813 sequence contains the complete first exon and first intron of the SM {alpha}-actin gene and was used to test the role of the first intron. Whereas basal activities of the three constructs were different, all three constructs were repressed equivalently by PDGF-BB and Ets-1 (Fig. 6). These results indicate that the –153 to +20 region is sufficient to confer PDGF-BB- and Ets-1-induced repression.



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  		Fig. 6. The –153 to +20 region of the SM {alpha}-actin promoter was sufficient to confer PDGF-BB- and Ets-1-induced repression. A: SMCs were transfected with –2555 to +2813, –155 to +2813, or –155 to +20 SM {alpha}-actin-luciferase construct, serum starved, and treated for 24 h with vehicle or PDGF-BB (30 ng/ml). B: SMCs were cotransfected with the same constructs and Ets-1-expressing construct (1 g/well of each construct) and cultured for 40 h in serum-free medium. Promoter activities were normalized to basal {alpha}-actin promoter activity observed with cells transfected with the –2555 to +2813 construct. *Significant difference (P <= 0.01) between treated and untreated cells and between Ets-1-transfected and control-transfected cells (paired t-test).

 
Ets-1 overexpression activated, rather than repressed, activity of the –153 to +20 region of the SM {alpha}-actin gene in endothelial cells. The –153 to +20 region of the SM {alpha}-actin promoter was previously shown to exhibit promoter activity in endothelial cells, which do not normally express the endogenous SM {alpha}-actin gene or the –2555 to +2813 SM {alpha}-actin promoter-reporter construct (30, 52). To test whether Ets-1 overexpression exhibited cell type selectivity, a series of SM {alpha}-actin reporter/Ets-1 cotransfection studies were conducted in endothelial cells. Consistent with previous observations, the –153 to +20 SM {alpha}-actin promoter construct displayed substantial promoter activity (15.1 ± 3.2-fold over promoterless), whereas the –2555 to +2813 SM {alpha}-actin construct had a very low activity in endothelial cells (2.4 ± 0.2-fold over promoterless; Fig. 7). Ets-1 overexpression induced an increase in the –153 to +20 (4.3 ± 0.3-fold) and the –2555 to +2813 (3.6 ± 0.2-fold) SM {alpha}-actin constructs that was similar to the increase in luciferase expression from the promoterless construct (5.3 ± 0.4-fold; Fig. 7). Ets-1 overexpression in endothelial cells induced a general increase in transcription in endothelial cells, and Ets-1-induced repression of the SM {alpha}-actin promoter is selective for SMCs. ACLP and c-fos promoter constructs were activated upon Ets-1 overexpression to a much higher level (9.2 ± 0.8- and 32.8 ± 5.1-fold, respectively), suggesting specific activating effects in the case of these genes. These results show that Ets-1 effects were cell selective, having effects in SMCs that were completely opposite those observed in endothelial cells.



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  		Fig. 7. Ets-1 overexpression induced a general increase in promoter activities in endothelial cells. Endothelial cells were cotransfected with promoterless luciferase, –155 to +20 SM {alpha}-actin-luciferase, –2555/+2813 SM {alpha}-actin-luciferase, ACLP-luciferase, or c-fos-luciferase construct and Ets-1-expressing construct or empty vector (1 g/well of each construct). Cells were cultured for 40 h in the presence of 10% serum. Promoter activity was normalized to that of cells transfected with the promoterless construct to show activities of the different promoters in endothelial cells (open bars) and, for each reporter construct, to that of cells transfected with the empty vector construct to show the effect of Ets-1 on the different promoters. *Significant difference (P <= 0.01) between cells transfected with promoterless and promoter construct (paired t-test). #Significant difference (P <= 0.01) between cells transfected with Ets-1 plasmid and empty plasmid (paired t-test).

 
Ets-1 overexpression downregulated expression of the endogenous SM {alpha}-actin gene. Whereas the results of reporter analysis showed that Ets-1 repressed the activity of exogenous SMC promoters, DNA constructs of such promoters are episomal and may not accurately reflect effects of Ets-1 on expression of endogenous genes. For example, effects might be quite different because of effects of chromatin structure and/or complex protein-nucleic acid interactions. Because the transfection efficiency of cultured SMCs is so low that effects of Ets-1 overexpression on endogenous SMC marker gene expression cannot be tested directly, SMCs were stably transfected with Ets-1-expressing construct, and effects on SM {alpha}-actin expression were analyzed. Seven independent clones were obtained and initially analyzed for expression of Ets-1 protein by Western blotting: four clones (B2, A2, B3, and A3) expressed high levels of Ets-1, one clone (B1) expressed very low levels of Ets-1, and two clones (A4 and A1) expressed no detectable Ets-1 protein (Fig. 8A). Total RNAs were analyzed by Northern blot for Ets-1, nonmuscle {beta}-actin, and SM {alpha}-actin mRNA expression (Fig. 8B). Consistent with results from transiently transfected SMC promoter-reporter constructs, quantitative analysis of the Northern blot results indicated that SM {alpha}-actin mRNA levels were negatively correlated with Ets-1 mRNA expression levels within these various stable lines (Fig. 8C). Nonmuscle {beta}-actin mRNA levels did not significantly change in the different clones, suggesting that Ets-1-induced repression was selective for SM {alpha}-actin. The B1 clone, which expressed very low levels of Ets-1, displayed an intermediary reduction in SM {alpha}-actin expression between high- and low Ets-1-expressing clones, suggesting that SM {alpha}-actin expression is very sensitive to Ets-1 expression. These results suggest that Ets-1 overexpression in SMCs is capable of downregulating expression of the SM {alpha}-actin promoter-reporter construct in transient transfection assays as well as the endogenous SM {alpha}-actin gene.

PDGF-BB-induced Ets-1 upregulation is not affected by cell density. On the basis of observations that PDGF-BB-mediated downregulation of the SMC marker promoter was cell density dependent and that Ets-1 expression is upregulated by PDGF-BB, we hypothesized that PDGF-BB may be unable to increase Ets-1 expression when cells are cultured at high cell density. Ets-1 mRNA levels were also upregulated by PDGF-BB at high cell density (Fig. 9), whereas SM {alpha}-actin mRNA levels were decreased. The latter observation is consistent with our previous observations that PDGF-BB downregulates SMC marker gene expression in postconfluent SMCs at least in part by inducing destabilization of SM {alpha}-actin mRNA (4, 8, 15). These results indicate that PDGF-BB-induced repression of SMC gene expression is not a simple function of the level of expression of Ets-1 but involves mechanisms that are much more complex (see DISCUSSION).


   DISCUSSION
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 ABSTRACT
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 DISCUSSION
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The goals of the present study were to identify molecular mechanisms whereby PDGF-BB induces downregulation of expression of SMC marker genes in cultured SMCs and to identify factors and mechanisms involved in this process. Our results provide clear evidence that PDGF-BB selectively repressed transcription of multiple SMC marker genes and that the effects were cell density dependent and mediated through the PDGF-{beta} receptor. Moreover, we found that PDGF-BB induced expression of Ets-1 in SMCs and that overexpression of Ets-1 markedly repressed SMC marker gene expression and SMC marker promoter activity in cultured SMCs. In contrast, in endothelial cells, Ets-1 overexpression had effects that were completely opposite those in SMCs, in that Ets-1 increased SM {alpha}-actin promoter activity. These results advance our understanding of mechanisms whereby PDGF-BB inhibits expression of SMC marker genes and, in conjunction with results of previous studies, provide several lines of evidence in support of the possibility that PDGF-BB-induced expression of Ets-1 may play an important role in control of phenotypic modulation of SMCs in vivo. 1) PDGF-BB and Ets-1 selectively repressed SM {alpha}-actin, SM MHC, and SM22{alpha}, but not ACLP, expression, consistent with the pattern of change in expression of these genes in response to vascular injury in vivo (25, 42). 2) Expression of PDGF-B chain and PDGF-{beta} receptor is upregulated in vascular lesions after balloon injury (32, 57) and during atherosclerosis (47, 59). Moreover, vascular injury is known to result in increased release of PDGF-BB. 3) Several reports have indicated that blocking PDGF-B/PDGF-{beta} receptor signaling attenuates neointimal thickening in several models of vascular injury, suggesting that PDGF-BB plays an important role in the response to injury (2, 9, 50). 4) Expression of Ets-1 is increased within 2 h of injury in the rat carotid artery and aorta and remains increased for >=2 wk, a period that coincides with reduced expression of SMC marker genes (11, 16, 42). Moreover, Ets-1 has been shown to be a trans-activating factor of the genes of several matrix metalloproteinases (51, 60), which are known to play a role in vessel remodeling and vascular lesion development (3). Transcription of the Ets-1 gene has also been shown to be decreased, whereas transcription of SMC marker genes is increased, by exposure of cultured SMCs to mechanical strain (45, 60). Taken together, the results of this study and previous studies suggest that PDGF-BB-induced expression of Ets-1 may play a role in induction of phenotypic modulation of SMCs after vessel injury. However, there is no direct evidence that this is the case in vivo, and because PDGF-B chain and PDGF-{beta} receptor knockout mice die during embryonic development (26, 56) and Ets-1 knockout mouse embryos show variable lethality (34), further studies to directly test this hypothesis will likely depend on development of conditional transgenic knockout systems (43).

A potentially important observation in the present studies was that PDGF-BB-induced repression of SM {alpha}-actin promoter did not occur in SMC cultures when cells were grown to high density, suggesting that cell-cell contact, altered cell-matrix interactions, or other density-dependent changes influenced PDGF-BB-induced effects. Previous studies in our laboratory showed that expression of SMC differentiation marker genes such as SM {alpha}-actin is increased by growth arrest of subconfluent cells in serum-free defined media or by density-dependent growth inhibition resulting from increased cell-cell contact (5, 39). We previously showed that postconfluent cells respond to PDGF-BB by eliciting a posttranscriptional repressive effect through a very rapid and selective degradation of SMC marker mRNAs (7, 8, 15). Taken together, these results suggest that the nature of PDGF-BB effects on SMC marker gene expression (i.e., transcriptional vs. posttranscriptional) could differ in vivo depending on the nature of the cell-cell and cell-matrix contacts of SMCs. We previously showed that injury-induced decreases in expression of SMC marker genes in a mouse carotid injury model are mediated at least in part at the transcriptional level (42). It is also intriguing that PDGF-{beta} receptor/PDGF-B chain expression during vascular development correlates with an immature differentiated state of SMC precursors in blood vessels (53). Thus it is interesting to speculate that the effects we observed in low-density cultures of SMCs in some way mimic the effects observed in vivo after mechanical disruption of cell-cell and cell-matrix contacts during vessel injury and in low-SMC-populated vessels during vascular development. These results are also consistent with a possible role for Ets-1 and PDGF-BB in injury-induced decreases in transcription, although there is no direct evidence that PDGF-BB exerts direct effects on SMC differentiation marker gene expression in vivo at the transcriptional or posttranscriptional level.

We also observed a PDGF-BB-induced increase in Ets-1 expression in postconfluent SMCs, although we did not observe transcriptional repression of SMC marker genes under these conditions. On the basis of the finding that orthovanadate treatment resulted in PDGF-BB-induced transcriptional repression in high-density cells (Fig. 2), it is interesting to speculate that the density dependence of the response could occur through interaction of Ets-1 with one or more factors with activity that is sensitive to cell density through some tyrosine phosphatase-dependent pathway. Consistent with this possibility, although we found that Ets-1 repressed the activity of the –155 to +20 region of the SM {alpha}-actin promoter, we did not find any sequence in that region that could be a potential Ets-1 binding site. The Ets-1 consensus binding site has been shown to have the sequence 5'-A/GCCGGAA/TGT/C-3' (37). Given the extensive body of evidence that control of SMC gene expression is dependent on complex combinatorial interactions of many cis elements and trans-binding factors (for review see Ref. 22) and on the basis of these lines of evidence, we speculate that Ets-1- and PDGF-BB-mediated repression of SMC marker promoters may be regulated, at least in part, through complex interactions of multiple trans-regulatory factors and may or may not require Ets-1 binding to DNA. However, much additional work is needed to directly test this hypothesis, because little or nothing is known regarding potential interactions of Ets-1 with transcription factors known to be required for SMC gene expression in SMCs.

A provocative observation in the present studies was that selective inhibition of the PDGF-{beta} receptor using a PDGF tyrosine kinase receptor inhibitor completely reversed the effects of PDGF-BB, resulting in marked activation of the SM {alpha}-actin promoter. This result suggests that PDGF-BB may elicit very different effects on SMC marker transcription, depending on the receptor/signaling pathway activated. One possibility is that PDGF-BB-induced activation of the SM {alpha}-actin promoter is mediated through activation of the PDGF-{alpha} receptor. Indeed, cultured SMCs express the two known PDGF receptor subunits: {alpha} and {beta}. The {alpha}- and {beta}-subunits bind the PDGF-B chain, whereas the {alpha}-subunit only binds the PDGF-A chain. However, the PDGF-BB-mediated increase in SM {alpha}-actin promoter activity is probably not related to activation of the PDGF-{alpha} receptor, because we found that PDGF-AA had no effect on expression of any of the SMC promoter-reporter constructs tested (data not shown). Another possibility is that AG-1295 treatment only partially blocked PDGF-{beta} receptor signaling pathways, resulting in activation of a select subset of downstream signaling pathways, thereby altering effects on SMC gene transcription. Consistent with this possibility, Reusch et al. (44) showed that selective blockade of the phosphoinositide-3 kinase pathway associated with PDGF-BB stimulation increased SM MHC promoter activity through a Ras/Raf/MEK/ERK pathway. However, it is still possible that another PDGF-{beta} receptor may be involved in mediating the PDGF-BB-induced increase in SM {alpha}-actin promoter activity in the absence of PDGF-{beta} receptor signaling and that the simultaneous activation of both receptors may lead to a decrease in promoter activity. Although this question is of interest, supplementary work would first require identification of that new receptor for the PDGF-B chain. In any case, identification of mechanisms involved in mediating PDGF-BB-induced increases in SMC promoter activity is potentially very important, in that it may explain how, under some conditions, including early development, PDGF-BB could activate SMC differentiation. For example, Lu et al. (28) reported that PDGF-BB activated expression of SMC markers such as calponin, SM {gamma}-actin, and SM22{alpha} during SMC-mesenchymal transformation of proepicardial cells. Depending on signaling pathway availability and the level of tyrosine phosphatase in the cell, PDGF-BB may therefore trigger profoundly different effects on SMC gene expression. However, further studies involving selective inhibition or knockdown of the various PDGF signaling pathways will be important to directly ascertain how specific PDGF-activated signaling pathways are coupled to control of SMC marker gene expression.

Taken together, results of the present study provide evidence that PDGF-BB and Ets-1 induce potent coordinate repression of multiple SMC differentiation marker genes in cultured SMCs. However, many unresolved issues remain to be addressed, including 1) direct investigation of the role of Ets-1 and/or PDGF-{beta} receptor pathways in control of SMC phenotypic modulation after vascular injury in vivo using various knockout and overexpression models; 2) investigation of the possible role of other cytokines, including basic fibroblast growth factor and TNF-{alpha}, which are also increased within vascular lesions and known to induce expression of Ets-1 in cultured SMCs (11, 35); and 3) further elucidation of mechanisms by which Ets-1 induces coordinate repression of multiple SMC genes, including investigation of its interaction/effects on several common transcriptional activation pathways, such as CArG/SRF, which have been shown to be involved in controlling coordinate expression of virtually all the SMC marker genes that have been identified (17, 30, 33).


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grants RO1 HL-38854, R37 HL-57353, and PO1 HL-19242 to G. K. Owens, a postdoctoral fellowship grant from Fondation pour la Recherche Médicale (Paris, France) to F. Dandré, and American Heart Association Mid-Atlantic Affiliate Virginia Heart Association Postdoctoral Fellowship Grant 0020263U to F. Dandré.


   ACKNOWLEDGMENTS
 
We thank Dr. David Camerini (University of Virginia) and his group for use of their luminometer; Diane Raines, Marit Kingston, Doug Mullinex, and Rupanda Tripathi for expert technical assistance; and Brian Wamhoff for reading the manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. K. Owens, Cardiovascular Research Center, Univ. of Virginia, PO Box 801394, Charlottesville, VA 22908-1394 (E-mail: gko{at}virginia.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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Adam PJ, Regan CP, Hautmann MB, and Owens GK. Positive- and negative-acting Kruppel-like transcription factors bind a transforming growth factor-{beta} control element required for expression of the smooth muscle cell differentiation marker SM22{alpha} in vivo. J Biol Chem 275: 37798–37806, 2000.[Abstract/Free Full Text]
  2. Banai S, Wolf Y, Golomb G, Pearle A, Waltenberger J, Fishbein I, Schneider A, Gazit A, Perez L, Huber R, Lazarovichi G, Rabinovich L, Levitzki A, and Gertz SD. PDGF-receptor tyrosine kinase blocker AG1295 selectively attenuates smooth muscle cell growth in vitro and reduces neointimal formation after balloon angioplasty in swine. Circulation 97: 1960–1969, 1998.[Abstract/Free Full Text]
  3. Bendeck MP, Zempo N, Clowes AW, Galardy RE, and Reidy MA. Smooth muscle cell migration and matrix metalloproteinase expression after arterial injury in the rat. Circ Res 75: 539–545, 1994.[Abstract/Free Full Text]
  4. Blank RS and Owens GK. Platelet-derived growth factor regulates actin isoform expression and growth state in cultured rat aortic smooth muscle cells. J Cell Physiol 142: 635–642, 1990.[CrossRef][ISI][Medline]
  5. Blank RS, Thompson MM, and Owens GK. Cell cycle versus density dependence of smooth muscle {alpha}-actin expression in cultured rat aortic smooth muscle cells. J Cell Biol 107: 299–306, 1988.[Abstract/Free Full Text]
  6. Bochaton-Piallat ML, Ropraz P, Gabbiani F, and Gabbiani G. Phenotypic heterogeneity of rat arterial smooth muscle cell clones. Implications for the development of experimental intimal thickening. Arterioscler Thromb Vasc Biol 16: 815–820, 1996.[Abstract/Free Full Text]
  7. Corjay MH, Blank RS, and Owens GK. Platelet-derived growth factor-induced destabilization of smooth muscle {alpha}-actin mRNA. J Cell Physiol 145: 391–397, 1990.[CrossRef][ISI][Medline]
  8. Corjay MH, Thompson MM, Lynch KR, and Owens GK. Differential effect of platelet-derived growth factor- versus serum-induced growth on smooth muscle {alpha}-actin and non-muscle {beta}-actin mRNA expression in cultured rat aortic smooth muscle cells. J Biol Chem 264: 10501–10506, 1989.[Abstract/Free Full Text]
  9. Ferns GA, Raines EW, Sprugel KH, Motani AS, Reidy MA, and Ross R. Inhibition of neointimal smooth muscle accumulation after angioplasty by an antibody to PDGF. Science 253: 1129–1132, 1991.[Abstract/Free Full Text]
  10. Gilman MZ, Wilson RN, and Weinberg RA. Multiple protein-binding sites in the 5'-flanking region regulate c-fos expression. Mol Cell Biol 6: 4305–4316, 1986.[Abstract/Free Full Text]
  11. Goetze S, Kintscher U, Kaneshiro K, Meehan WP, Collins A, Fleck E, Hsueh WA, and Law RE. TNF{alpha} induces expression of transcription factors c-fos, Egr-1, and Ets-1 in vascular lesions through extracellular signal-regulated kinases 1/2. Atherosclerosis 159: 93–101, 2001.[CrossRef][ISI][Medline]
  12. Hautmann MB, Adam PJ, and Owens GK. Similarities and differences in smooth muscle {alpha}-actin induction by TGF-{beta} in smooth muscle versus non-smooth muscle cells. Arterioscler Thromb Vasc Biol 19: 2049–2058, 1999.[Abstract/Free Full Text]
  13. Hayashi K, Takahashi M, Kimura K, Nishida W, Saga H, and Sobue K. Changes in the balance of phosphoinositide 3-kinase/protein kinase B (Akt) and the mitogen-activated protein kinases (ERK/p38MAPK) determine a phenotype of visceral and vascular smooth muscle cells. J Cell Biol 145: 727–740, 1999.[Abstract/Free Full Text]
  14. Heldin CH and Westermark B. Mechanism of action and in vivo role of platelet-derived growth factor. Physiol Rev 79: 1283–1316, 1999.[Abstract/Free Full Text]
  15. Holycross BJ, Blank RS, Thompson MM, Peach MJ, and Owens GK. Platelet-derived growth factor-BB-induced suppression of smooth muscle cell differentiation. Circ Res 71: 1525–1532, 1992.[Abstract/Free Full Text]
  16. Hultgardh-Nilsson A, Cercek B, Wang JW, Naito S, Lovdahl C, Sharifi B, Forrester JS, and Fagin JA. Regulated expression of the Ets-1 transcription factor in vascular smooth muscle cells in vivo and in vitro. Circ Res 78: 589–595, 1996.[Abstract/Free Full Text]
  17. Kim S, Ip HS, Lu MM, Clendenin C, and Parmacek MS. A serum response factor-dependent transcriptional regulatory program identifies distinct smooth muscle cell sublineages. Mol Cell Biol 17: 2266–2278, 1997.[Abstract]
  18. Kitange G, Shibata S, Tokunaga Y, Yagi N, Yasunaga A, Kishikawa M, and Naito S. Ets-1 transcription factor-mediated urokinase-type plasminogen activator expression and invasion in glioma cells stimulated by serum and basic fibroblast growth factors. Lab Invest 79: 407–416, 1999.[ISI][Medline]
  19. Ko JH, Miyoshi E, Noda K, Ekuni A, Kang R, Ikeda Y, and Taniguchi N. Regulation of the GnT-V promoter by transcription factor Ets-1 in various cancer cell lines. J Biol Chem 274: 22941–22948, 1999.[Abstract/Free Full Text]
  20. Kocher O, Gabbiani F, Gabbiani G, Reidy MA, Cokay MS, Peters H, and Huttner I. Phenotypic features of smooth muscle cells during the evolution of experimental carotid artery intimal thickening. Biochemical and morphologic studies. Lab Invest 65: 459–470, 1991.[ISI][Medline]
  21. Kovalenko M, Gazit A, Bohmer A, Rorsman C, Ronnstrand L, Heldin CH, Waltenberger J, Bohmer FD, and Levitzki A. Selective platelet-derived growth factor receptor kinase blockers reverse cis-transformation. Cancer Res 54: 6106–6114, 1994.[Abstract/Free Full Text]
  22. Kumar MS and Owens GK. Combinatorial control of smooth muscle-specific gene expression. Arterioscler Thromb Vasc Biol 23: 737–747, 2003.[Abstract/Free Full Text]
  23. Kusuoka H and Hoffman JI. Advice on statistical analysis for circulation research. Circ Res 91: 662–671, 2002.[Abstract/Free Full Text]
  24. Layne MD, Endege WO, Jain MK, Yet SF, Hsieh CM, Chin MT, Perrella MA, Blanar MA, Haber E, and Lee ME. Aortic carboxypeptidase-like protein, a novel protein with discoidin and carboxypeptidase-like domains, is up-regulated during vascular smooth muscle cell differentiation. J Biol Chem 273: 15654–15660, 1998.[Abstract/Free Full Text]
  25. Layne MD, Yet SF, Maemura K, Hsieh CM, Liu X, Ith B, Lee ME, and Perrella MA. Characterization of the mouse aortic carboxypeptidase-like protein promoter reveals activity in differentiated and dedifferentiated vascular smooth muscle cells. Circ Res 90: 728–736, 2002.[Abstract/Free Full Text]
  26. Leveen P, Pekny M, Gebre-Medhin S, Swolin B, Larsson E, and Betsholtz C. Mice deficient for PDGF B show renal, cardiovascular, and hematological abnormalities. Genes Dev 8: 1875–1887, 1994.[Abstract/Free Full Text]
  27. Li L, Miano JM, Mercer B, and Olson EN. Expression of the SM22{alpha} promoter in transgenic mice provides evidence for distinct transcriptional regulatory programs in vascular and visceral smooth muscle cells. J Cell Biol 132: 849–859, 1996.[Abstract/Free Full Text]
  28. Lu J, Landerholm TE, Wei JS, Dong XR, Wu SP, Liu X, Nagata K, Inagaki M, and Majesky MW. Coronary smooth muscle differentiation from proepicardial cells requires RhoA-mediated actin reorganization and p160 Rho-kinase activity. Dev Biol 240: 404–418, 2001.[CrossRef][ISI][Medline]
  29. Mack CP and Owens GK. Regulation of smooth muscle {alpha}-actin expression in vivo is dependent on CArG elements within the 5' and first intron promoter regions. Circ Res 84: 852–861, 1999.[Abstract/Free Full Text]
  30. Mack CP, Thompson MM, Lawrenz-Smith S, and Owens GK. Smooth muscle {alpha}-actin CArG elements coordinate formation of a smooth muscle cell-selective, serum response factor-containing activation complex. Circ Res 86: 221–232, 2000.[Abstract/Free Full Text]
  31. Madsen CS, Regan CP, Hungerford JE, White SL, Manabe I, and Owens GK. Smooth muscle-specific expression of the smooth muscle myosin heavy chain gene in transgenic mice requires 5'-flanking and first intronic DNA sequence. Circ Res 82: 908–917, 1998.[Abstract/Free Full Text]
  32. Majesky MW, Reidy MA, Bowen-Pope DF, Hart CE, Wilcox JN, and Schwartz SM. PDGF ligand and receptor gene expression during repair of arterial injury. J Cell Biol 111: 2149–2158, 1990.[Abstract/Free Full Text]
  33. Manabe I and Owens GK. CArG elements control smooth muscle subtype-specific expression of smooth muscle myosin in vivo. J Clin Invest 107: 823–834, 2001.[ISI][Medline]
  34. Muthusamy N, Barton K, and Leiden JM. Defective activation and survival of T cells lacking the Ets-1 transcription factor. Nature 377: 639–642, 1995.[CrossRef][Medline]
  35. Naito S, Shimizu S, Maeda S, Wang J, Paul R, and Fagin JA. Ets-1 is an early response gene activated by ET-1 and PDGF-BB in vascular smooth muscle cells. Am J Physiol Cell Physiol 274: C472–C480, 1998.[Abstract/Free Full Text]
  36. Niwa H, Yamamura K, and Miyazaki J. Efficient selection for high-expression transfectants with a novel eukaryotic vector. Gene 108: 193–199, 1991.[CrossRef][ISI][Medline]
  37. Nye JA, Petersen JM, Gunther CV, Jonsen MD, and Graves BJ. Interaction of murine Ets-1 with GGA-binding sites establishes the ETS domain as a new DNA-binding motif. Genes Dev 6: 975–990, 1992.[Abstract/Free Full Text]
  38. Owens GK. Molecular control of vascular smooth muscle cell differentiation. Acta Physiol Scand 164: 623–635, 1998.[ISI][Medline]
  39. Owens GK, Loeb A, Gordon D, and Thompson MM. Expression of smooth muscle-specific {alpha}-isoactin in cultured vascular smooth muscle cells: relationship between growth and cytodifferentiation. J Cell Biol 102: 343–352, 1986.[Abstract/Free Full Text]
  40. Pallen CJ and Tong PH. Elevation of membrane tyrosine phosphatase activity in density-dependent growth-arrested fibroblasts. Proc Natl Acad Sci USA 88: 6996–7000, 1991.[Abstract/Free Full Text]
  41. Ranganna K and Yatsu FM. Inhibition of platelet-derived growth factor BB-induced expression of glyceraldehyde-3-phosphate dehydrogenase by sodium butyrate in rat vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 17: 3420–3427, 1997.[Abstract/Free Full Text]
  42. Regan CP, Adam PJ, Madsen CS, and Owens GK. Molecular mechanisms of decreased smooth muscle differentiation marker expression after vascular injury. J Clin Invest 106: 1139–1147, 2000.[ISI][Medline]
  43. Regan CP, Manabe I, and Owens GK. Development of a smooth muscle-targeted Cre recombinase mouse reveals novel insights regarding smooth muscle myosin heavy chain promoter regulation. Circ Res 87: 363–369, 2000.[Abstract/Free Full Text]
  44. Reusch HP, Zimmermann S, Schaefer M, Paul M, and Moelling K. Regulation of Raf by Akt controls growth and differentiation in vascular smooth muscle cells. J Biol Chem 276: 33630–33637, 2001.[Abstract/Free Full Text]
  45. Reusch P, Wagdy H, Reusch R, Wilson E, and Ives HE. Mechanical strain increases smooth muscle and decreases nonmuscle myosin expression in rat vascular smooth muscle cells. Circ Res 79: 1046–1053, 1996.[Abstract/Free Full Text]
  46. Ross R. The pathogenesis of atherosclerosis: a perspective for the 1990s. Nature 362: 801–809, 1993.[CrossRef][Medline]
  47. Ross R, Masuda J, Raines EW, Gown AM, Katsuda S, Sasahara M, Malden LT, Masuko H, and Sato H. Localization of PDGF-B protein in macrophages in all phases of atherogenesis. Science 248: 1009–1012, 1990.[Abstract/Free Full Text]
  48. Saito Y, Haendeler J, Hojo Y, Yamamoto K, and Berk BC. Receptor heterodimerization: essential mechanism for platelet-derived growth factor-induced epidermal growth factor receptor transactivation. Mol Cell Biol 21: 6387–6394, 2001.[Abstract/Free Full Text]
  49. Salhany KE, Robinson-Benion C, Candia AF, Pledger WJ, and Holt JT. Differential induction of the c-fos promoter through distinct PDGF receptor-mediated signaling pathways. J Cell Physiol 150: 386–395, 1992.[CrossRef][ISI][Medline]
  50. Sano H, Sudo T, Yokode M, Murayama T, Kataoka H, Takakura N, Nishikawa S, Nishikawa SI, and Kita T. Functional blockade of platelet-derived growth factor receptor-{beta} but not of receptor-{alpha} prevents vascular smooth muscle cell accumulation in fibrous cap lesions in apolipoprotein E-deficient mice. Circulation 103: 2955–2960, 2001.[Abstract/Free Full Text]
  51. Sato Y, Teruyama K, Nakano T, Oda N, Abe M, Tanaka K, and Iwasaka-Yagi C. Role of transcription factors in angiogenesis: Ets-1 promotes angiogenesis as well as endothelial apoptosis. Ann NY Acad Sci 947: 117–123, 2001.[Abstract/Free Full Text]
  52. S