Heparan Sulfate Proteoglycans Mediate a Potent Inhibitory Signal for Migration of Vascular Smooth Muscle Cells

From the Departments of Surgery (N.K., U.H., A.W.C.) and Pathology (M.G.K., T.N.W.), University of Washington, Seattle.

Correspondence to Alexander W. Clowes, MD, Department of Surgery, Box 356410, University of Washington, Seattle, WA 98195-6410. E-mail clowes{at}u.washington.edu

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Abstract—Migration of vascular smooth muscle cells (SMCs) is a key step in vascular remodeling and formation of pathological lesions in diseased arteries and may be controlled by extracellular matrix (ECM) and by factors that regulate ECM composition, such as platelet-derived growth factor (PDGF). In culture, PDGF-AB and -BB enhance but PDGF-AA (although having no effect alone) suppresses SMC migration stimulated by other PDGF isoforms. To determine whether the migration-inhibitory mechanism of PDGF-AA was mediated by ECM composition, we examined baboon SMC migration in a Boyden chamber assay using filters coated with different ECM proteins. PDGF-AA suppressed the PDGF-BB–induced migration of baboon SMCs on a filter coated with basement membrane proteins (Matrigel) and fibronectin but failed to inhibit cell migration on a type I collagen (Vitrogen)-coated filter. Fibronectin and fibronectin fragments that contain heparin-binding domains permitted PDGF-AA inhibition of cell migration, but a fragment lacking heparin-binding domains did not. Treatment of SMCs with heparin lyases II and III, but not with chondroitin ABC lyase, diminished the PDGF-AA–mediated inhibition of migration. PDGF-AA stimulated accumulation of proteoglycan (PG) in the cell layer more potently than did PDGF-BB, whereas the turnover of cell layer PG was unaffected by either PDGF-AA or -BB. Northern blot analysis revealed that PDGF-AA increased syndecan-1 mRNA expression more than did PDGF-BB, whereas both PDGF isoforms decreased perlecan expression. The changes in cell migration and PG synthesis induced by PDGF-AA were accompanied by changes in the morphology of SMCs. PDGF-AA dramatically induced the spreading of SMCs, whereas the heparin lyase treatment of PDGF-AA–stimulated cultures diminished cell spreading. The data suggest that PDGF-AA selectively modifies heparan sulfate PG accumulation on SMCs and thereby influences the interactions of SMCs with heparin-binding ECM proteins. These interactions, in turn, generate signals that suppress SMC migration.

Key Words: heparan sulfate • smooth muscle cell • platelet-derived growth factor • extracellular matrix

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
The migration of vascular SMCs is a crucial event in the formation of intimal thickening in injured vessels, healing grafts and atherosclerotic lesions.^{1 2} SMC migration is regulated in part by PDGF. Three isoforms of PDGF (PDGF-AA, -AB, and -BB) and 2 PDGF-Rs (PDGF-R and -Rß) have been identified. Numerous observations suggest that signal transduction by PDGF requires receptor dimerization. Because of the ligand-binding specificity of the 2 receptors, PDGF-AA binds only to dimers, PDGF-AB binds to or ß dimers, and PDGF-BB binds to , ß, or ßß dimers.³

Both mRNA and protein of PDGF-A and -B chains have been detected in a variety of human and animal atherosclerotic tissues.^{2 4} In addition, both PDGF-A and -B chain mRNAs are expressed in human restenotic tissues after angioplasty.⁵ Infusion of anti–PDGF-A chain antibody abolishes normal cardiovascular development in murine embryos,⁶ and mice carrying a null mutation in the gene encoding the PDGF-B chain have phenotypes that include gross abnormalities of the heart and aorta.⁷ These reports suggest that PDGF has an important role in vascular morphogenesis and pathogenesis.

PDGF isoforms differ in their effects on vascular SMC migration. PDGF-AB and -BB enhance rat SMC migration in a modified Boyden chamber assay, whereas PDGF-AA inhibits the migration induced by PDGF-AB and PDGF-BB.⁸ Our studies using anti–PDGF-R and anti–PDGF-Rß antibodies indicate that PDGF-R generates an inhibitory signal for baboon SMC migration, regardless of the ligand.⁹ In another study, porcine endothelial cells transfected with native and mutant PDGF-R were used to identify 3 tyrosine residues (tyr 768, 993, and 1018) in the cytoplasmic domain of PDGF-R that are critical for the signaling that results in the inhibition of cell migration.¹⁰ These data provide evidence that PDGF-AA selectively and specifically suppresses PDGF-BB–induced cell migration.

ECM is an important component in the regulation of SMC migration.¹¹ SMC migration is suppressed when cells adhere tightly to fibronectin but not to type IV collagen.¹² Cell-associated HSPGs mediate and modulate some aspects of the interactions of cells with their ECM. The principal plasma membrane–associated HSPGs, which may function as matrix receptors,¹³ are those of the syndecan family of transmembrane PGs and the glypican-related PGs that are intercalated in the plasma membrane via a phosphatidylinositol anchor.¹⁴ ECM-associated HSPGs, such as perlecan, are present primarily in basement membranes and laminae adjacent to specific cell types, such as SMCs and endothelial cells.¹⁵ Plasma membrane–associated HSPGs interact with heparin-binding ECM proteins via their heparan sulfate chains, and perlecan, in addition to charge interactions of glycosaminoglycan chains, also is integrated within the ECM by the interactions of specific core protein domains.^{13 16}

A number of studies regarding the involvement of HSPG in development and pathology suggest a significant role for HSPG in the control of SMC proliferation and migration.^{13 17} It is well documented that the content and composition of HSPGs change during naturally occurring and experimentally induced atherosclerotic lesion formation.^{18 19 20 21} Although it is not clear which factors regulate HSPG expression in the pathogenesis of atherosclerosis, certain growth factors, such as PDGF, which is known to be involved in the pathogenesis of this disease, also modulate HSPG metabolism by SMCs.^{21 22}

Therefore, we have investigated the possibility that the inhibition of SMC migration by PDGF-AA operates through a mechanism that involves SMC HSPGs. Our results show that PDGF-AA selectively affects the expression of a specific HSPG (syndecan-1) and that HSPGs influence the facility with which SMCs migrate on a fibronectin-rich ECM in a Boyden chamber assay. Thus, we conclude that one signal that regulates SMC migration operates through a mechanism involving cell-associated HSPGs.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Materials
Recombinant human PDGF-AA (short variant) and PDGF-BB were kindly supplied by Dr Charles Hart (Zymogenetics Inc, Seattle, Wash). Plasma fibronectin and heparin lyases II and III were purchased from Sigma Chemical Co. Fibronectin fragments, which were affinity-purified from chymotrypsin-digested fibronectin, were from GIBCO BRL. Pepsin-digested type I collagen (Vitrogen) was obtained from Celtrix Co. Matrigel, a basement membrane matrix extract whose major components are laminin and type IV collagen, was from Collaborative Research. Chondroitin ABC lyase was from ICN Biochemicals.

Cell Culture
Baboon vascular SMCs were isolated by the explant method. Briefly, aortic explants were obtained from the thoracic aorta of baboons and cultured in DMEM supplemented with 10% FBS. After 2 weeks, the cells that had migrated out of the explant were removed by trypsinization and seeded in T-75 flasks. Confluent SMCs at the second passage were subcultured successively at a 1:2 split ratio. SMCs were used up to the 15th passage. Serum-free medium used for the preparation of SMC suspensions consisted of DMEM supplemented with insulin (10 µg/mL), transferrin (5 µg/mL), and ovalbumin (1 mg/mL).

Migration and Adhesion Assays and Morphometric Analysis of Cultured SMCs
Migration of SMCs was assayed in a modification of the Boyden chamber method using 48-well microchemotaxis chambers (Neuro Probe Inc) and polycarbonate filters (Nucleopore Corp) with pores of 10.0-µm diameter. The filters were precoated with 0.5 µg/well of basement membrane proteins (Matrigel), type I collagen (Vitrogen), fibronectin, or fibronectin fragment peptides, as indicated below. Cultured SMCs were trypsinized and suspended in serum-free DMEM; 20 000 cells were placed in the upper chamber; and 25 µL of serum-free medium containing PDGF-AA, PDGF-BB, or both was placed in the lower chamber. In experiments in which enzymatic digestion of HSPGs and CSPGs was required, the SMC suspensions were incubated with 20 U/mL of heparin lyase II and 0.2 U/mL of heparin lyase III or 2.0 U/mL of chondroitin ABC lyase for 1 hour before introduction into the upper chamber of the Boyden apparatus. To minimize deposition of intact PG during the experiment, the cell suspension also included these enzymes during the assay. Microchemotaxis chambers seeded with cells were incubated at 37°C under 5% CO₂ in air for 6 hours. At the end of the assay period, filters were removed from the chamber, and SMCs remaining on the upper side of the filter were removed. The SMCs that had migrated to the lower side of the filter were fixed in methanol, stained with Diff-Quick staining solution (Baxter) and counted under a microscope (x100) to quantify SMC migration. Migration activity was expressed as the mean number of cells that had migrated per high-power field. For the adhesion assay, 2000 cells were seeded into the upper chamber, and after a 2-hour incubation period, the SMCs that attached to the upper-side of the ECM-coated filter were fixed and counted, as above. For morphometric measurements, cell perimeters were measured by using a digitizing pad to trace the edges of SMCs as projected from a microscope image (x400). Fifty single cells were analyzed for each condition.

[³⁵S]Sulfate Radiolabeling and Characterization of PG
Cultured SMCs were trypsinized, suspended in serum-free and sulfate-free DMEM supplemented with carrier-free [³⁵S]sulfate (ICN-Radiochemicals) at 100 µCi/mL, and seeded at 3x10⁵ cells/well into 48-well plates (Corning Co) precoated with 100 µg/mL of Matrigel. After incubation for the indicated times with PDGF-AA or PDGF-BB at 10 ng/mL, the cell layers were washed with PBS twice, and ³⁵S-labeled PG was harvested with 8 mol/L urea, 0.2% Triton X-100, and 0.25% SDS, containing proteinase inhibitors, including 100 mmol/L 6-aminohexanoic acid, 5 mmol/L benzamidine, 10 mmol/L N-ethylmaleimide, and 1 mmol/L phenylmethylsulfonyl fluoride. Determinations of total labeled PGs were obtained from duplicate determinations on 50 µL aliquots of samples using a cetylpyridinium chloride precipitation assay.²³ Proportions of ³⁵S label that were incorporated into HSPG and CSPG subclasses were determined by digestion with chondroitin ABC lyase.

RNA Extraction and Northern Blot Analysis
Cultured SMCs were trypsinized, suspended in serum-free medium, and seeded. After a 6-hour incubation with PDGF-AA or PDGF-BB at 10 ng/mL, total RNA was isolated from cells by the single-step extraction method.²⁴ Total RNA (10 µg) was loaded per lane and resolved by electrophoresis overnight on 1% (wt/vol) agarose-formaldehyde gels. After electrophoresis, RNA was transferred to Zetaprobe GT (Bio-Rad Laboratories) and UV–cross-linked (Stratagene Cloning Systems). Before hybridization, filters were prehybridized for at least 2 hours at 42°C in a solution containing 50% (vol/vol) formamide (Life Technologies, Inc), 6x SSPE, 5x Denhardt's solution, 0.5% SDS, 5% dextran sulfate, and 100 µg/mL salmon sperm DNA (Sigma). For the preparation of hybridization probes, a partial human perlecan cDNA (HS-1)²⁵ was kindly supplied by Dr R.V. Iozzo, Thomas Jefferson University, Philadelphia, Pa, and a partial mouse syndecan-1 cDNA was provided by Dr M. Jalkanen, University of Turku, Turku, Finland. Probes were ³²P-labeled by random priming (Amersham) using 5'-[-³²P]dCTP (Amersham). Hybridizations with ³²P-labeled cDNA probes were carried out at 42°C for at least 16 hours, after which the filters were washed 3 times with 2x SSPE/0.1% SDS at 42°C and twice with 0.3x SSPE/0.1% SDS at 65°C. Autoradiographs were prepared by exposure on Kodak XAR2 film at -70°C and then developed. Quantification of radiolabeled bands was by scanning densitometry of the fluorograph.²⁶ Northern blots were normalized for loading by comparison with ethidium bromide staining of the 28S rRNA band.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
ECM-Dependent Regulation of SMC Migration by PDGF-AA
We determined the inhibitory activity of PDGF-AA on SMC migration induced by 10 ng/mL PDGF-BB on a filter coated with ECM containing fibronectin, basement membrane proteins (Matrigel), type I collagen (Vitrogen), and collagen/fibronectin (1:1) (Figure 1). PDGF-AA dose-dependently suppressed SMC migration on a filter coated with fibronectin, Matrigel, or collagen/fibronectin. At PDGF concentrations of 10 ng/mL, the inhibition of migration was maximal. In contrast, PDGF-AA at concentrations of up to 20 ng/mL failed to suppress migration on a filter coated only with type I collagen. PDGF-AA alone did not affect the basal migration, regardless of the ECM protein present on the filter (not shown), in agreement with earlier studies.^{8 9} These data suggest that the SMC migration-inhibitory signal generated by PDGF-AA depends on the nature of the ECM.

View larger version (22K): [in this window] [in a new window]   Figure 1. The effect of ECM on SMC migration regulated by PDGF-AA. The Boyden chamber filter was coated with ECM, including type I collagen (), Matrigel (), fibronectin (), and a 1:1 mixture of collagen and fibronectin (). Migration was induced by 10 ng/mL of PDGF-BB, and the activity was assayed in the absence and presence of PDGF-AA at indicated concentrations, as shown in Materials and Methods. Migration activity was expressed as the mean number of cells per high-power field (HPF).

Boyden chamber filters were coated with collagen and fibronectin, mixed in different ratios, and the effect of these substrates on SMC migration was determined (Figure 2). On ECM mixtures containing 25% and 50% fibronectin, basal and PDGF-BB–induced migrations were identical to migration on substrates without fibronectin, but PDGF-AA inhibition of PDGF-BB–stimulated migration increased as the proportion of fibronectin increased. PDGF-AA inhibited migration by 42% on ECM containing 25% fibronectin, and inhibition reached 80% at 50% fibronectin. At the highest proportions of fibronectin, migration in response to PDGF-BB was reduced, and inhibition by PDGF-AA was greater. Control experiments indicated that cell attachment was identical at 0% to 50% fibronectin and increased by 20% at 75% and 100% fibronectin (data not shown). These data indicate that PDGF-AA suppressed PDGF-BB–induced SMC migration on fibronectin but not on type I collagen.

View larger version (35K): [in this window] [in a new window]   Figure 2. The effect of the ratio of collagen and fibronectin on SMC migration regulated by PDGF-AA and PDGF-BB. The filter of Boyden chamber was coated with an indicated ratio mixture of collagen and fibronectin. Migration activity was assayed in the absence (open bars) or presence of 10 ng/mL of PDGF-BB (hatched bars) or PDGF-BB plus 10 ng/mL of PDGF-AA (solid bars) as shown in Materials and Methods. The ratio of collagen and fibronectin is shown as a percentage of fibronectin, and results are expressed as the mean number of cells (±SD) per high-power field (HPF). Statistical analysis of data using the unpaired t test (2-tailed) compared PDGF-BB+PDGF-AA vs PDGF-BB alone. *P<0.01.

Inhibition of SMC Migration by PDGF-AA Involves the Heparin-Binding Domain of Fibronectin
Fibronectin has both N- and C-terminus heparin-binding domains,²⁷ which may be involved in modifying or mediating cell-adhesive interactions,²⁸ as well as an internal domain that includes an integrin binding site. Therefore, chymotrypsin-generated peptides that include the heparin-binding domains of fibronectin were used to coat Boyden chamber filters to test whether these domains in fibronectin are involved in the inhibition of SMC migration by PDGF-AA (Figure 3). In these experiments, fibronectin peptides were combined with an equal proportion of type I collagen to prevent decreased cell adhesion to the filter. Intact fibronectin, as well as the 45-kDa and 40-kDa peptides containing, respectively, the N- and C-terminal fibronectin heparin-binding domains, promoted the inhibition of PDGF-BB–stimulated cell migration by PDGF-AA. PDGF-AA inhibited migration 59% on the 45-kDa N-terminal peptide, 66% on the 40-kDa C-terminal peptide, and 71% on a filter coated with intact fibronectin. In contrast, induced cell migration was not inhibited by PDGF-AA when cells were assayed on a filter coated with a 120-kDa fibronectin peptide, which includes an integrin-dependent cell-binding domain and lacks heparin-binding domains. Cell attachment was not affected by different fibronectin peptide:type I collagen substrata (data not shown), and basal migration was identical, suggesting that the heparin-binding domains of fibronectin were necessary for the inhibition of PDGF-BB–stimulated SMC migration by PDGF-AA, but not cell adhesion, in this assay.

View larger version (37K): [in this window] [in a new window]   Figure 3. The effect of fibronectin (Fn) peptides on SMC migration regulated by PDGF-AA and PDGF-BB. The filter of Boyden chamber was coated with the 1:1 mixture of type I collagen with intact Fn or peptides derived from Fn by chymotryptic digestion, containing an N-terminal 45-kDa heparin-binding fragment (N-45k), a central 120-kDa cell-binding RGD-containing fragment (120k), and a C-terminal 40-kDa heparin-binding fragment (C-40k). Migration was assayed in the absence (open bars) or presence of 10 ng/mL of PDGF-BB (hatched bars) or PDGF-BB plus 10 ng/mL of PDGF-AA (solid bars). Migration was expressed as the mean number of cells (±SD) per high-power field (HPF). Statistical analyses were made of data with PDGF-BB+PDGF-AA vs PDGF-BB alone by unpaired t test (2-tailed). *P<0.01.

HSPGs, but Not CSPGs, Are Involved in the Regulation of SMC Migration by PDGF-AA
PGs are known to affect the interaction of cells with their ECM and to influence cell migration. The observation that heparin-binding, but not integrin-binding, fibronectin domains mediate the inhibition of SMC migration by PDGF-AA suggests that PGs that interact with these domains may be present on SMCs. To differentiate between the involvement of HSPGs and CSPGs in the modulation of SMC migration, cells were treated with heparin lyases II and III or chondroitin ABC lyase and placed onto a Matrigel-coated filter, and migration in response to PDGF isoforms was assayed. The heparin lyase treatment completely blocked the inhibitory effect of PDGF-AA on cell migration stimulated by PDGF-BB (Figure 4A), whereas basal and PDGF-BB–induced migrations were not affected. In contrast, the digestion of CSPG by chondroitin ABC lyase did not affect SMC migration (Figure 4B). Cell adhesion to the filter was not affected by the treatment of cells with these enzymes (data not shown). These results indicate that HSPGs are required for the inhibition of PDGF-BB–induced cell migration by PDGF-AA.

View larger version (33K): [in this window] [in a new window]   Figure 4. The effect of digestion of PG on SMC migration regulated by PDGF-AA and PDGF-BB. The Boyden chamber filter was coated with Matrigel. Cells in suspension were treated for 30 minutes before seeding with heparin lyase II and III (A) or chondroitin ABC lyase (B). Migration activity was assayed in the absence (open bars) or presence of 10 ng/mL of PDGF-BB (hatched bars) or PDGF-BB plus 10 ng/mL of PDGF-AA (solid bars). Migration activity was expressed as the mean number of cells (±SD) per high-power field (HPF). Control indicates no enzyme treatment. Statistical analyses were made of data with PDGF-BB+PDGF-AA vs PDGF-BB alone by unpaired t test (2-tailed). *P<0.01.

Differential Regulation of PG Synthesis by PDGF Isoforms
To determine whether PDGF-AA and -BB have different effects on the PGs deposited around SMCs, we measured the incorporation of [³⁵S]SO₄ into cell layer–associated PG. Both PDGF-AA and -BB dose-dependently increased PG synthesis by SMCs (Figure 5). PDGF-AA increased [³⁵S]SO₄ incorporation into PG 27% over control at 10 ng/mL. At 2 to 10 ng/mL, PDGF-AA was more potent than PDGF-BB, but at 20 ng/mL, the relative increase in [³⁵S]SO₄ incorporation was greater with PDGF-BB stimulation. [³⁵S]SO₄-labeled PG was isolated from cell layers of untreated cultures and from cultures treated with 10 ng/mL of PDGF isoforms, either alone or in combination, and relative HSPG content was determined after chondroitin ABC lyase digestion in 4 independent experiments. The proportion of HSPG in cell layer samples (18%) was not changed after treatment of cells with any PDGF isoform. The accumulation of [³⁵S]SO₄-labeled PG in cell layers increased in a time-dependent manner (Figure 6). At 24 hours, PDGF-AA increased cell layer–associated PG by 2.4-fold over control; PDGF-BB enhanced cell layer PG accumulation by 1.5-fold. Since this baboon SMC line expresses more PDGF-Rß than PDGF-R and since PDGF-BB is more potent in inducing mitogenesis than is PDGF-AA,⁹ the stimulation of PG synthesis in response to PDGF-AA cannot be explained by differences in receptor number or proliferative activity of this factor.

View larger version (18K): [in this window] [in a new window]   Figure 5. The dose-dependent effect of PDGF-AA () and PDGF-BB () on PG synthesis in SMC cell layers. PG produced by SMCs was labeled with [35S]SO4 and determined using a cetylpyridinium chloride precipitation assay. [35S]PG labeled in the absence of PDGF was 2.7x10 counts/well.5 Results were the average value of 2 independent wells and shown as the increase over control. Representative data for 4 experiments are shown.

View larger version (31K): [in this window] [in a new window]   Figure 6. The time-dependent change of PG synthesis in SMCs induced by PDGF-AA and PDGF-BB. PG produced by SMCs was labeled with [35S]SO4 for the indicated period in the absence (open bars) or presence of PDGF-AA (solid bars) or PDGF-BB (hatched bars) and determined using cetylpyridinium chloride precipitation assay. Results are the average value of 2 independent wells and are shown as counts per well. Representative data are shown for 3 experiments.

To examine whether the increase in cell layer PG after PDGF treatment was due to a decrease in PG turnover, pulse-chase experiments were performed in the absence or presence of PDGF-AA and PDGF-BB (Figure 7). After PG was labeled with a 6-hour pulse of [³⁵S]SO₄ in the absence of PDGF, cultures were rinsed and chased in fresh DMEM containing PDGF for 6 and 18 hours before the incorporated [³⁵S]SO₄ remaining in the cell layer was determined. The time-dependent decrease in PG accumulation in the cell layer was not different between PDGF-AA–and PDGF-BB–treated cultures (46.6% and 47.8% at 6 hours and 74.7% and 71.9% at 18 hours, respectively). These data indicate that the increase in cell layer PG accumulation caused by PDGF is not due to changes in PG turnover.

View larger version (24K): [in this window] [in a new window]   Figure 7. The effect of PDGF-AA and PDGF-BB on turnover of cell layer PG. PGs produced by SMCs were labeled with [35S]SO4 for 6 hours. After the cell layer was thoroughly washed with PBS, unlabeled DMEM was added, and cultures were incubated for the indicated period in the absence (open bars) or presence of PDGF-AA (solid bars) or PDGF-BB (hatched bars). Incorporated [35S]SO4 in the cell layer was determined using a cetylpyridinium chloride precipitation assay. Results are the average value of 2 independent wells and are shown as counts per well. Representative data are shown for 3 experiments.

Regulation of HSPG mRNA Expression by PDGF Isoforms
In our previous work, syndecan-1 and perlecan mRNA expression was induced after balloon injury of rat carotid arteries.²⁰ Northern blot analysis of syndecan-1 and perlecan mRNA expression was used to determine whether the increase in PG synthesis by SMCs treated with PDGF was accompanied by the induction of HSPG mRNA expression (Figure 8). PDGF-AA and -BB had different effects on syndecan-1 and perlecan expression. Syndecan-1 mRNA was upregulated 41% by PDGF-AA, whereas PDGF-BB increased the level of this transcript only 15%. In contrast, perlecan mRNA was downregulated 31% by PDGF-AA, 42% by PDGF-BB, and 49% by their combination. Since the induction of syndecan-1 mRNA by PDGF-AA and -BB was consistent with the increase of ³⁵S-labeled HSPG, it is possible that the increase of HSPG by PDGF-AA was in part due to the upregulation of syndecan-1 mRNA.

View larger version (41K): [in this window] [in a new window]   Figure 8. Effect of PDGF-AA and -BB on the mRNA expression of syndecan-1 and perlecan. A, Total RNA was isolated from SMCs cultured for 6 hours without or with PDGF-AA and/or PDGF-BB at 10 ng/mL. mRNA expression for syndecan-1 and perlecan was determined by Northern blotting. The ethidium bromide (EtBr)–stained band of 28S rRNA is shown to compare relative loading among lanes. B, The bands of syndecan-1, perlecan, and 28S rRNA were quantified by scanning densitometry. The mRNA expression of syndecan-1 (hatched bars) and perlecan (open bars) normalized to 28S were indicated as a relative value to control (untreated SMCs). Similar results were obtained in 2 experiments.

Modulation of Cell Morphology by PDGF-AA and HSPG
Cell-ECM interaction affects cytoskeletal architecture and cell migration. We determined the effect of PDGF isoforms and heparin lyase treatment on SMC morphology on a Matrigel-coated filter to examine whether the PDGF-AA–modulated cell-ECM interaction involved HSPG (Figure 9). Attached unstimulated SMCs were spindle-shaped (Figure 9A). PDGF-AA induced dramatic cell spreading and elongation, whereas PDGF-BB had only a minor effect on cell shape (Figure 9B and 9C). Morphometric analysis was used to quantify SMC spreading (Figure 10). In the absence of PDGF, the distribution of SMC perimeters showed a sharp peak with the average value of 92 µm. PDGF-AA dramatically increased the mean perimeter of SMCs (161 µm), whereas PDGF-BB treatment did not significantly affect the SMC mean perimeter (115 µm). Treatment of SMCs with heparin lyase II and III significantly suppressed SMC spreading by PDGF-AA (Figure 9D, 9E, and 9F) and decreased the mean SMC perimeter by 55%, whereas no significant changes were observed in control and PDGF-BB–treated SMCs (Figure 10). In contrast, treatment of cells with chondroitin ABC lyase did not affect SMC spreading by PDGF-AA (data not shown). Similar effects on SMC spreading were induced by PDGF-AA or heparin lyase treatment when cells were seeded on a fibronectin-coated filter or seeded on an uncoated filter in the presence of 10% FBS (data not shown). These observations suggest that PDGF-AA induces SMC spreading by affecting SMC-ECM interactions that are mediated through HSPG.

View larger version (152K): [in this window] [in a new window]   Figure 9. Morphological change in SMCs induced by PDGF-AA and PDGF-BB. Cells were seeded without (A and D) or with PDGF-AA (B and E) or PDGF-BB (C and F) on a Matrigel-coated filter with 10-µm pores. After 2 hours of incubation, cells were fixed, stained, and observed with a microscope. In some experiments (D to F), cells in suspension were treated with heparin lyases before seeding.

View larger version (27K): [in this window] [in a new window]   Figure 10. The quantitative analysis of the morphological change of SMCs. Cells were seeded onto a Matrigel-coated filter and incubated without () or with PDGF-AA () or PDGF-BB () for 6 hours either without (A) or with (B) heparin lyase treatment. The morphological change of SMCs was quantified as described in Materials and Methods. Fifty cells were analyzed for each condition, and results are shown as perimeter (x-axis) of each cell and frequency (y-axis). Results are shown in panel C as mean±SE for cells treated without (open bars) or with heparin lyases (solid bars). Statistical analyses were made of data of SMCs treated with PDGF-AA or PDGF-BB vs control SMCs (no PDGF and no heparin lyase treatment) by unpaired t test (2-tailed) (n=50). **P<0.001; *P<0.01.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
We previously reported that PDGF-AA, which alone does not affect rat and baboon SMC basal migration, inhibits PDGF-BB–induced chemotaxis in Boyden chamber assays.^{8 9} We now show that PDGF-AA suppresses PDGF-BB–stimulated SMC migration by a mechanism that involves endogenously synthesized HSPGs and requires heparin-binding ECM as a migration substratum. Moreover, the suppression of cell migration by PDGF-AA is accompanied by an HSPG-dependent increase in cell spreading and an increased cell layer PG accumulation that are consistent with increased expression of syndecan.

The finding that selective induction of HSPG by PDGF-AA correlates with increased cell spreading and inhibition of induced chemotaxis agrees with other studies that suggest that decreased cell adhesion and induction of cell migration are accompanied by a relative decrease in the synthesis of HSPG.^{29 30 31 32} For example, the expression of the transmembrane HSPG, syndecan-1, is downregulated on circulating B lymphocytes and increased during their differentiation into immobilized cells.³⁰ A B-lymphoid cell line transfected with syndecan-1 gene shows reduced migration.³¹ Conversely, downregulation of syndecan-1 with expressed anti-sense has been shown to result in a conversion of epithelial cells to a nonanchorage-dependent fibroblastoid morphology.³² Proteolytic cleavage and release of syndecans from the cell surface may also be important for the regulation of cell function. Ectodomains of syndecan-1 and -4 are shed in response to signals from receptors for thrombin and heparin-binding epidermal growth factor.³³ Syndecans released from the cell membrane appear to enhance SMC migration in vitro^{34 35} and may diminish the antimigratory activity of membrane-associated syndecans. Cleavage of the glycosaminoglycan chains on HSPGs may also diminish this effect. For example, heparan sulfate–degrading enzymes induce neutrophil migration.³⁶ Other HSPGs are associated with decreased cell proliferation and migration. The large basement membrane HSPG, perlecan, is expressed at high levels by quiescent SMCs, whereas little is synthesized by growing cells, either in vitro or during embryonic development of the rat arterial wall.³⁷ Perlecan also inhibits the induction of the growth-related transcription factor Oct-1 and supports growth arrest by cultured SMCs.³⁸ Recent experiments that have used perlecan anti-sense constructs to limit endogenous perlecan expression by fibrosarcoma cells have demonstrated that perlecan may suppress cell proliferation and migration.³⁹ Thus, like syndecans, perlecan may inhibit cell migration and proliferation. However, in the present study, the inhibition of PDGF-BB–induced SMC migration by PDGF-AA correlated more closely with the selective upregulation of syndecan-1 expression, whereas perlecan is downregulated regardless of PDGF isoform (Figure 8), suggesting that syndecan-1 may be essential to this inhibitory signal.

Cell layer HSPGs, such as syndecans, may modulate cell migration and cell morphology by regulating the type of cell-ECM adhesion site that is formed. Syndecan-1 transfected into Schwann cells associates with actin and induces cell spreading.⁴⁰ Moreover, syndecan-4 has recently been specifically localized to focal adhesion sites,⁴¹ and syndecan-1 is associated with fibronectin in focal contacts and with stress fibers in fibroblasts.⁴² Fibronectin heparin-binding domains are required for the formation of focal adhesions in fibroblasts and endothelial cells,^{43 44} suggesting that cell surface HSPGs may regulate integrin-mediated attachment to the ECM. Taken together with our observation, these studies strongly support a role for HSPGs in the regulation of SMC migration by PDGF-AA. It is interesting to note that melanoma cell migration on type I collagen is dependent on a chondroitin sulfate–bearing variant of CD44 and can be abolished by chondroitin ABC lyase digestion.⁴⁵ Our observation that chondroitin ABC lyase had no effect on the migration of SMCs on fibronectin suggests that different cell surface PGs may regulate cell migration when cells are in contact with different ECM proteins.

The effect of HSPG in the suppression of SMC migration is substrate specific. In the present study, the inhibition of PDGF-BB–induced SMC migration by PDGF-AA depends on the presence of heparin-binding domains of fibronectin (Figure 5) but not on the RGD-containing fibronectin domain that mediates integrin-dependent cell attachment. The ß₁ subunit–containing integrins of SMCs are the dominant receptors for the cell attachment domain of fibronectin, and integrin expression and conformation are important in SMC migration, both in vitro and in vivo.⁴⁶ However, it is unlikely that PDGF-AA and -BB exert their effects on SMC migration through these receptors, since neither PDGF isoform affects the expression or activation of ß₁ integrins,⁴⁷ although expression of _vß₃ integrin is stimulated by PDBF-BB⁴⁸ and is important for SMC migration on other matrices.⁴⁹ A role for fibronectin in the inhibition of cell migration has been documented in other systems. For example, an enrichment of fibronectin in the ECM suppressed the migration of NIH 3T3 cells,⁵⁰ and overexpression of fibronectin in fibrosarcoma cells blocked their invasion.⁵¹ Our data that PDGF-AA failed to suppress SMC migration on a Vitrogen-coated filter are consistent with the report that HSPG binds to intact fibrillar type I collagen but not to pepsin-digested type I collagen, such as Vitrogen.⁵² Clearly, other matrix macromolecules and matrix receptors also influence SMC migration. For example, SMC migration in vitro is enhanced by hyaluronan through a receptor for hyaluronan-mediated motility (RHAMM),⁵³ whereas hyaluronidase diminishes PDGF-BB–induced migration.⁵⁴ Hyaluronan production is stimulated by PDGF-BB and PDGF-Rß, but not by PDGF-AA,⁵⁵ consistent with the stimulatory effect of hyaluronan on cell migration. A combination of diverse stimulatory and inhibitory signals may ultimately serve to regulate the migratory response of SMCs.

The mechanisms by which PDGF isoforms induce different effects on cellular migration and phenotype is complex and not well understood. However, recent studies using endothelial cells transfected with wild and mutant PDGF-R clearly indicate that PDGF-R activates 2 signaling pathways that differentially affect cell migration.¹⁰ Both PDGF-AA and -BB activate MAP kinase and induce SMC proliferation, with PDGF-BB more potent than PDGF-AA, consistent with the larger number of PDGF-Rß present on these cells.⁹ However, PDGF-BB stimulates the phosphorylation of p125 focal adhesion kinase and tensin in SMCs and induces migration, whereas PDGF-AA does not.⁵⁶ These observations suggest that differential signaling pathways activated by PDGF-R and PDGF-Rß may be responsible for the different effects of PDGF isoforms on SMC migration. We previously reported that PDGF-AA suppresses SMC migration induced by chemoattractants, including PDGF-BB, but does not inhibit unstimulated migration,⁹ in agreement with reports by other laboratories (eg, see References 56 and 57^{56 57} ). However, others have reported that PDGF-AA induces chemotaxis in several cell types, including vascular SMCs.^{58 59 60} There are several observations that may ultimately reconcile these disparate results. One possibility is that ligand binding or receptor signaling by PDGF-R may be modulated by interaction with other variably expressed matrix proteins or signaling through other receptors. For example, PDGF isoforms bind differentially to the extracellular glycoprotein SPARC, an interaction that affects the activity of the bound mitogen.⁶¹ The report that antibodies against the cell surface receptor NG-2, which is involved in SMC mitogenesis and chemotaxis, decrease the cellular responsiveness to PDGF-AA and that PDGF-BB responsiveness is unaffected⁶² suggests that subtle interactions of signaling pathways induced by other receptors may influence PDGF-R signaling as well. Finally, Ferns et al⁶⁰ demonstrated that signaling through PDGF receptors in SMCs may vary with receptor number, as exemplified by the augmentation of migration by PDGF-R–overexpressing cells in response to PDGF-AA on Vitrogen-coated filters. This observation suggests that the low numbers of PDGF-R present on some SMCs are insufficient to induce migration to the degree seen by the stimulation of the more abundant PDGF-Rß. However, in the present study, PDGF-AA promoted HSPG production and syndecan-1 mRNA expression by SMCs more potently than did PDGF-BB. Since HSPG-mediated interactions with ECM are involved in the inhibition of SMC migration by PDGF-AA, the possibility remains that differential induction by different PDGF isoforms of either HSPG matrix receptors or heparin-binding ECM proteins gives rise to different effects of PDGF-AA and -BB on SMC migration. Interestingly, PDGF-AA induces cAMP and activates protein kinase A.⁶³ Since cAMP both potently suppresses SMC migration⁸ and induces the expression of syndecan-1 in peritoneal macrophages and NIH 3T3 cells,^{64 65} induction of syndecan expression by cAMP may be a mechanism for the PDGF-AA–mediated inhibition of PDGF-BB–induced SMC migration.

Although PDGF-AA is mitogenic for SMCs, PDGF-AA is expressed in nonproliferating cells in human atherosclerotic tissues.⁴ For example, a recent report by Murry et al⁶⁶ used quantitative PCR to demonstrate that PDGF-AA mRNA levels were 100 times higher in quiescent human aortic tissues than in advanced atherosclerotic plaques. HSPGs, which we and others²² have shown are induced by PDGF, support maintenance of a differentiated cellular phenotype in SMCs and other cells.^{67 68} Analysis of promoter sequences of syndecan-1 and -4 genes reveals that expression is controlled in part by E-box nuclear factors, which are also involved in regulation of gene expression during muscle cell differentiation.^{69 70} HSPG genes, including syndecan-1 and perlecan, are activated in injured rat arteries and may play a role in intimal thickening.^{20 21} In addition, HSPG isolated from rat arterial wall suppressed the expansion of the neointima when introduced into injured arteries.⁷¹ Taken together, PDGF-AA and HSPG may function to regulate SMC migration and maintain vascular structure rather than to induce proliferation. Conversely, decreased expression of PDGF-AA and HSPG may be crucial for the progression of atherosclerotic lesions.

	Selected Abbreviations and Acronyms

 

CSPG = chondroitin sulfate PG ECM = extracellular matrix HSPG = heparan sulfate PG PDGF-R = PDGF receptor PDGF = platelet-derived growth factor PG = proteoglycan SMC = smooth muscle cell

	Acknowledgments

 
This study was supported by grants HL-18645 and HL-616775 from the National Institutes of Health. We thank Christina K. Tsoi for assistance in PG analysis, Dr Susan D. Perigo for technical suggestions and scientific discussions, Trevina W. Wang for performing cell culture, and Sachiko Koyama for preparation of the manuscript.

Received August 29, 1997; accepted April 16, 1998.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 
1. Clowes AW, Reidy MA, Clowes MM. Kinetics of cellular proliferation after arterial injury, I: smooth muscle cell growth in the absence of endothelium. Lab Invest. 1983;49:327–333.[Medline] [Order article via Infotrieve]

2. Ross R. The pathogenesis of atherosclerosis: a perspective for the 1990s. Nature. 1993;362:801–809.[Medline] [Order article via Infotrieve]

3. Claesson-Welsh L. Platelet-derived growth factor receptor signals. J Biol Chem. 1994;269:32023–32026.[Free Full Text]

4. Rekhter MD, Gordon D. Does platelet-derived growth factor-A chain stimulate proliferation of arterial mesenchymal cells in human atherosclerotic plaques? Circ Res. 1994;75:410–417.[Abstract/Free Full Text]

5. Ueda M, Becker A, Kasayuki N, Kojima A, Morita Y, Tanaka S. In situ detection of platelet-derived growth factor-A and -B chain mRNA in human coronary arteries after percutaneous transluminal coronary angioplasty. Am J Pathol. 1996;149:831–843.[Abstract]

6. Schattemann GC, Loushin C, Li T, Hart CE. PDGF-A is required for normal murine cardiovascular development. Dev Biol. 1996;176:133–142.[Medline] [Order article via Infotrieve]

7. Levéen P, Pekny M, Gebre-Medhin S, Swolin B, Larsson E, Betsholtz C. Mice deficient for PDGF B show renal, cardiovascular, and hematological abnormalities. Genes Dev. 1994;8:1875–1887.[Abstract/Free Full Text]

8. Koyama N, Morisaki N, Saito Y, Yoshida S. Regulatory effects of platelet-derived growth factor-AA homodimer on migration of vascular smooth muscle cells. J Biol Chem. 1992;267:22806–22812.[Abstract/Free Full Text]

9. Koyama N, Hart CE, Clowes AW. Different functions of platelet-derived growth factor- and -ß receptors for the migration and proliferation of baboon vascular smooth muscle cells. Circ Res. 1994;75:682–691.[Abstract/Free Full Text]

10. Yokote K, Mori S, Siegbahn A, Ronnstrand L, Wernstedt C, Heldin C-H, Claesson-Welsh L. Structural determinants in the platelet-derived growth factor -receptor implicated in modulation of chemotaxis. J Biol Chem. 1996;271:5101–5111.[Abstract/Free Full Text]

11. Schor SL. Cytokine control of cell motility: modulation and mediation by the extracellular matrix. Prog Growth Factor Res. 1994;5:223–248.[Medline] [Order article via Infotrieve]

12. DiMilla PA, Stone JA, Quinn JA, Albelda SM, Lauffenburger DA. Maximal migration of human smooth muscle cells on fibronectin and type IV collagen occurs at an intermediate attachment strength. J Cell Biol. 1993;122:729–737.[Abstract/Free Full Text]

13. Bernfield M, Kokenyesi R, Kato M, Hinkes MT, Spring J, Gallo RL, Lose EJ. Biology of the syndecans: a family of transmembrane heparan sulfate proteoglycans. Annu Rev Cell Biol. 1992;8:365–393.

14. David G. Integral membrane heparan sulfate proteoglycans. FASEB J. 1993;7:1023–1030.[Abstract]

15. Noonan DM, Hassell JR. Perlecan, the large low-density proteoglycan of basement membranes: structure and variant forms. Kidney Int. 1993;43:53–60.[Medline] [Order article via Infotrieve]

16. Wight TN, Kinsella MG, Qwarnström EE. The role of proteoglycans in cell adhesion, migration and proliferation. Curr Opin Cell Biol. 1992;4:793–801.[Medline] [Order article via Infotrieve]

17. David G, Bai X, Van der Schueren B, Cassiman JJ, Van den Berghe H. Developmental changes in heparan sulfate expression: in situ detection with monoclonal antibodies. J Cell Biol. 1992;119:961–975.[Abstract/Free Full Text]

18. Alavi MZ, Wasty F, Li Z, Galis ZS, Ismail N, Moore S. Enhanced incorporation of [¹⁴C]glucosamine into glycosaminoglycans of aortic neointima of balloon-injured and cholesterol-fed rabbits in vitro. Atherosclerosis. 1992;95:59–67.[Medline] [Order article via Infotrieve]

19. Wasty F, Alavi MZ, Moore S. Distribution of glycosaminoglycans in the intima of human aortas: changes in atherosclerosis and diabetes mellitus. Diabetologia. 1993;36:316–322.[Medline] [Order article via Infotrieve]

20. Nikkari ST, Jarvelainen HT, Wight TN, Ferguson M, Clowes AW. Smooth muscle cell expression of extracellular matrix genes after arterial injury. Am J Pathol. 1994;144:1348–1356.[Abstract]

21. Cizmeci-Smith G, Langan E, Youkey J, Showalter LJ, Carey DJ. Syndecan-4 is a primary-response gene induced by basic fibroblast growth factor and arterial injury in vascular smooth muscle cells. Arterioscler Thromb Vasc Biol. 1997;17:172–180.[Abstract/Free Full Text]

22. Cizmeci-Smith G, Stahl RC, Showalter LJ, Carey DJ. Differential expression of transmembrane proteoglycans in vascular smooth muscle cells. J Biol Chem. 1993;268:18740–18747.[Abstract/Free Full Text]

23. Glimelius B, Norling B, Westermark B, Wasteson A. Turnover of cell surface associated glycosaminoglycans in cultures of human normal and malignant glial cells. Exp Cell Res. 1978;117:179–189.[Medline] [Order article via Infotrieve]

24. Chomczynski P, Sacchi N. Single step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem. 1987;162:156–159.[Medline] [Order article via Infotrieve]

25. Murdoch AD, Dodge GR, Cohen I, Tuan RS, Iozzo RV. Primary structure of the human heparan sulfate proteoglycan from basement membrane (HSPG2/perlecan): a chimeric molecule with multiple domains homologous to the low density lipoprotein receptor, laminin, neural cell adhesion molecules, and epidermal growth factor. J Biol Chem. 1992;267:8544–8557.[Abstract/Free Full Text]

26. Kinsella MG, Tsoi CK, Järveläinen HT, Wight TN. Selective expression and processing of biglycan during migration of bovine aortic endothelial cells: the role of endogenous basic fibroblast growth factor. J Biol Chem. 1997;272:318–325.[Abstract/Free Full Text]

27. Hynes RO. Molecular biology of fibronectin. Annu Rev Cell Biol. 1985;1:67–90.

28. Woods A, Couchman JR, Johansson S, Höök M. Adhesion and cytoskeletal organisation of fibroblasts in response to fibronectin fragments. EMBO J. 1986;5:665–670.[Medline] [Order article via Infotrieve]

29. Kinsella MG, Wight TN. Modulation of sulfated proteoglycan synthesis by bovine aortic endothelial cells during migration. J Cell Biol. 1986;102:679–687.[Abstract/Free Full Text]

30. Sanderson RD, Lalor P, Bernfield M. B lymphocytes express and lose syndecan at specific stages of differentiation. Cell Regul. 1989;1:27–35.[Medline] [Order article via Infotrieve]

31. Liebersbach BF, Sanderson RD. Expression of syndecan-1 inhibits cell invasion into type I collagen. J Biol Chem. 1994;269:20013–20019.[Abstract/Free Full Text]

32. Kato M, Saunders S, Nguyen H, Bernfield M. Loss of cell surface syndecan-1 causes epithelia to transform into anchorage-independent mesenchyme-like cells. Mol Biol Cell. 1995;6:559–576.[Abstract]

33. Subramanian SV, Fitzgerald ML, Bernfield M. Regulated shedding of syndecan-1 and -4 ectodomains by thrombin and growth factor receptor activation. J Biol Chem. 1997;272:14713–14720.[Abstract/Free Full Text]

34. Higashiyama S, Abraham JA, Klagsbrun M. Heparin-binding EGF-like growth factor stimulation of smooth muscle cell migration: dependence on interactions with cell surface heparan sulfate. J Cell Biol. 1993;122:933–940.[Abstract/Free Full Text]

35. Noda-Heiny H, Sobel BE. Vascular smooth muscle cell migration mediated by thrombin and urokinase receptor. Am J Physiol. 1995;268:C1195–C1201.[Abstract/Free Full Text]

36. Hoogewerf AJ, Leone JW, Reardon IM, Howe WJ, Asa D, Heinrikson RL, Ledbetter SR. CXC chemokines, connective tissue activating peptide-III and neutrophil activating peptide-2 are heparin-heparan sulfate-degrading enzymes. J Biol Chem. 1995;270:3268–3277.[Abstract/Free Full Text]

37. Weiser MCM, Grieshaber SS, Belknap JK, Kinsella MG, Majack RA. Developmental regulation of perlecan gene expression in aortic smooth muscle cells. Matrix Biol. 1996;15:331–340.[Medline] [Order article via Infotrieve]

38. Weiser MCM, Grieshaber NA, Schwartz PE, Majack RA. Perlecan regulates Oct-1 gene expression in vascular smooth muscle cells. Mol Biol Cell. 1997;8:999–1011.[Abstract]

39. Mathiak M, Yenisey C, Grant DS, Sharma B, Iozzo RV. A role for perlecan in the suppression of growth and invasion in fibrosarcoma cells. Cancer Res. 1997;57:2130–2136.[Abstract/Free Full Text]

40. Carey DJ, Stahl RC, Cizmeci-Smith G, Asundi VK. Syndecan-1 expressed in Schwann cells causes morphological transformation and cytoskeletal reorganization and associates with actin during cell spreading. J Cell Biol. 1994;124:161–170.[Abstract/Free Full Text]

41. Woods A, Couchman JR. Syndecan 4 heparan sulfate proteoglycan is a selectively enriched and widespread focal adhesion component. Mol Biol Cell. 1994;5:183–192.[Abstract]

42. Yamagata M, Saga S, Kato M, Bernfield M, Kimata K. Selective distributions of proteoglycans and their ligands in pericellular matrix of cultured fibroblasts: implications for their roles in cell-substratum adhesion. J Cell Sci. 1993;106:55–65.[Abstract]

43. Woods A, McCarthy JB, Furcht LT, Couchman JR. A synthetic peptide from the COOH-terminal heparin-binding domain of fibronectin promotes focal adhesion formation. Mol Biol Cell. 1993;4:605–613.[Abstract]

44. Huebsch JC, McCarthy JB, Diglio CA, Mooradian DL. Endothelial cell interactions with synthetic peptides from the carboxy-terminal heparin-binding domains of fibronectin. Circ Res. 1995;77:43–53.[Abstract/Free Full Text]

45. Faassen AE, Schrager JA, Klein DJ, Oegema TR, Couchman JR, McCarthy JB. A cell surface chondroitin sulfate proteoglycan, immunologically related to CD44, is involved in type I collagen-mediated melanoma cell motility and invasion. J Cell Biol. 1992;116:521–531.[Abstract/Free Full Text]

46. Koyama N, Seki J, Vergel S, Mattsson EJR, Yednock T, Kovach NL, Harlan JM, Clowes AW. Regulation and function of an activation-dependent epitope of the ß1 integrins in vascular cells after balloon injury in baboon arteries and in vitro. Am J Pathol. 1996;148:749–761.[Abstract]

47. Seki J, Koyama N, Kovach NL, Yednock T, Clowes AW, Harlan JM. Regulation of ß₁-integrin function in cultured human vascular smooth muscle cells. Circ Res. 1996;78:596–605.[Abstract/Free Full Text]

48. Janat MF, Argraves WS, Liau G. Regulation of vascular smooth muscle cell integrin expression by transforming growth factor beta 1 and by platelet-derived growth factor-BB. J Cell Physiol. 1992;151:588–595.[Medline] [Order article via Infotrieve]

49. Clyman RI, Mauray F, Kramer RH. Beta 1 and beta 3 integrins have different roles in the adhesion and migration of vascular smooth muscle cells on extracellular matrix. Exp Cell Res. 1992;200:272–284.[Medline] [Order article via Infotrieve]

50. Corbett SA, Wilson CL, Schwarzbauer JE. Changes in cell spreading and cytoskeletal organization are induced by adhesion to a fibronectin-fibrin matrix. Blood. 1996;88:158–166.[Abstract/Free Full Text]

51. Akamatsu H, Ichihara K, Ozono K, Kamiike W, Matsuda H, Sekiguchi K. Suppression of transformed phenotypes of human fibrosarcoma cells by overexpression of recombinant fibronectin. Cancer Res. 1996;56:4541–4546.[Abstract/Free Full Text]

52. Koda JE, Bernfield M. Heparan sulfate proteoglycans from mouse mammary epithelial cells. J Biol Chem. 1984;259:11763–11770.[Abstract/Free Full Text]

53. Savani RC, Wang C, Yang B, Zhang S, Kinsella MG, Wight TN, Stern R, Nance DM, Turley EA. Migration of bovine aortic smooth muscle cells after wounding injury: the role of hyaluronan and RHAMM. J Clin Invest. 1995;95:1158–1168.

54. Ellis I, Banyard J, Schor SL. Differential response of fetal and adult fibroblasts to cytokines: cell migration and hyaluronan synthesis. Development. 1997;124:1593–1600.[Abstract]

55. Heldin P, Laurent TC, Heldin CH. Effects of growth factors on hyaluronan synthesis in cultured human fibroblasts. Biochem J. 1989;258:919–922.[Medline] [Order article via Infotrieve]

56. Jiang B, Yamamura S, Nelson PR, Mureebe L, Kent KC. Differential effect of platelet-derived growth factor isotypes on human smooth muscle cell proliferation and migration are mediated by distinct signaling pathways. Surgery. 1996;120:427–431.[Medline] [Order article via Infotrieve]

57. Siegbahn A, Hammacher A, Westermark B, Heldin C-H. Differential effects of the various isoforms of platelet-derived growth factor on chemotaxis of fibroblasts, monocytes, and granulocytes. J Clin Invest. 1990;85:916–920.

58. Hayashi N, Takehara K, Soma Y. Differential chemotactic responses mediated by platelet-derived growth factor alpha- and beta-receptors. Arch Biochem Biophys. 1995;322:423–428.[Medline] [Order article via Infotrieve]

59. Uren A, Yu JC, Gholami NS, Pierce JH, Heidaran MA. The alpha PDGFR tyrosine kinase mediates locomotion of two different cell types through chemotaxis and chemokinesis. Biochem Biophys Res Commun. 1994;204:628–634.[Medline] [Order article via Infotrieve]

60. Ferns GAA, Sprugel KH, Seifert RA, Bowen-Pope DF, Kelly JD, Murray M, Raines EW, Ross R. Platelet-derived growth factor receptor subunit expression determines cell migration to different dimeric forms of PDGF. Growth Factors. 1990;3:315–324.[Medline] [Order article via Infotrieve]

61. Raines EW, Lane TF, Iruela-Arispe ML, Ross R, Sage EH. The extracellular glycoprotein SPARC interacts with platelet-derived growth factor (PDGF)-AB and -BB and inhibits the binding of PDGF to its receptors. Proc Natl Acad Sci U S A. 1992;89:1281–1285.[Abstract/Free Full Text]

62. Grako KA, Stallcup WB. Participation of the NG-2 proteoglycan in rat aortic smooth muscle cell responses to platelet-derived growth factor. Exp Cell Res. 1995;221:231–240.[Medline] [Order article via Infotrieve]

63. Graves LM, Bornfeldt KE, Sidhu JS, Argast GM, Raines EW, Ross R, Leslie CC, Krebs EG. Platelet-derived growth factor stimulates protein kinase A through a mitogen-activated protein kinase dependent pathway in human arterial smooth muscle cells. J Biol Chem. 1996;271:505–511.[Abstract/Free Full Text]

64. Gallo RL, Povsic TJ, Bernfield M. PR-39, an antimicrobial peptide, induces syndecans, binds a receptor and increases cAMP in mesenchymal cells [abstract]. Mol Biol Cell. 1995;6a:162.

65. Yeaman C, Rapraeger AC. Post-transcriptional regulation of syndecan-1 expression by cAMP in peritoneal macrophages. J Cell Biol. 1993;122:941–950.[Abstract/Free Full Text]

66. Murry CE, Bartosek T, Giachelli CM, Alpers CE, Schwartz SM. Platelet-derived growth factor-A mRNA expression in fetal, normal adult, and atherosclerotic human aortas. Circulation. 1996;93:1095–1106.[Abstract/Free Full Text]

67. Campbell JH, Rennick RE, Kalevitch SG, Campbell GR. Heparan sulfate-degrading enzymes induce modulation of smooth muscle phenotype. Exp Cell Res. 1992;200:156–167.[Medline] [Order article via Infotrieve]

68. Leppa S, Mali M, Miettinen HM, Jalkanen M. Syndecan expression regulates cell morphology and growth of mouse mammary epithelial tumor cells. Proc Natl Acad Sci U S A. 1992;89:932–6.[Abstract/Free Full Text]

69. Takagi A, Kojima T, Tsuzuki S, Katsumi A, Yamazaki T, Sugiura I, Hamaguchi M, Saito H. Structural organization and promoter activity of the human ryudocan gene. J Biochem. 1996;119:979–984.[Abstract/Free Full Text]

70. Hinkes MT, Goldberger OA, Neumann PE, Kokenyesi R, Bernfield M. Organization and promoter activity of the mouse syndecan-1 gene. J Biol Chem. 1993;268:11440–11448.[Abstract/Free Full Text]

71. Bingley JA, Campbell JH, Hayward IP, Campbell GR. Inhibition of neointimal formation by natural heparan sulfate proteoglycans of the arterial wall. Ann N Y Acad Sci. 1997;811:238–244.[Medline] [Order article via Infotrieve]

This article has been cited by other articles:

				J. M. Ricono, B. Wagner, Y. Gorin, M. Arar, A. Kazlauskas, G. G. Choudhury, and H. E. Abboud PDGF receptor-{beta} modulates metanephric mesenchyme chemotaxis induced by PDGF AA Am J Physiol Renal Physiol, February 1, 2009; 296(2): F406 - F417. [Abstract] [Full Text] [PDF]

				A. C. Doran, N. Meller, and C. A. McNamara Role of Smooth Muscle Cells in the Initiation and Early Progression of Atherosclerosis Arterioscler Thromb Vasc Biol, May 1, 2008; 28(5): 812 - 819. [Abstract] [Full Text] [PDF]

				W. T. Gerthoffer Mechanisms of Vascular Smooth Muscle Cell Migration Circ. Res., March 16, 2007; 100(5): 607 - 621. [Abstract] [Full Text] [PDF]

				A. Abramsson, S. Kurup, M. Busse, S. Yamada, P. Lindblom, E. Schallmeiner, D. Stenzel, D. Sauvaget, J. Ledin, M. Ringvall, et al. Defective N-sulfation of heparan sulfate proteoglycans limits PDGF-BB binding and pericyte recruitment in vascular development Genes & Dev., February 1, 2007; 21(3): 316 - 331. [Abstract] [Full Text] [PDF]

				W. Xiao, Y. Liu, and D. M. Templeton Ca2+/calmodulin-dependent protein kinase II inhibition by heparin in mesangial cells Am J Physiol Renal Physiol, January 1, 2005; 288(1): F142 - F149. [Abstract] [Full Text] [PDF]

				A. Segev, N. Nili, and B. H Strauss The role of perlecan in arterial injury and angiogenesis Cardiovasc Res, September 1, 2004; 63(4): 603 - 610. [Abstract] [Full Text] [PDF]

				Z. Zhou, J. Wang, R. Cao, H. Morita, R. Soininen, K. M. Chan, B. Liu, Y. Cao, and K. Tryggvason Impaired Angiogenesis, Delayed Wound Healing and Retarded Tumor Growth in Perlecan Heparan Sulfate-Deficient Mice Cancer Res., July 15, 2004; 64(14): 4699 - 4702. [Abstract] [Full Text] [PDF]

				P.-K. Tran, K. Tran-Lundmark, R. Soininen, K. Tryggvason, J. Thyberg, and U. Hedin Increased Intimal Hyperplasia and Smooth Muscle Cell Proliferation in Transgenic Mice With Heparan Sulfate-Deficient Perlecan Circ. Res., March 5, 2004; 94(4): 550 - 558. [Abstract] [Full Text] [PDF]

				C.-m. Li, D. Newman, J. Khosla, and P. L. Sannes Heparin Inhibits DNA Synthesis and Gene Expression in Alveolar Type II Cells Am. J. Respir. Cell Mol. Biol., September 1, 2002; 27(3): 345 - 352. [Abstract] [Full Text] [PDF]

				T. M. Grzeszkiewicz, V. Lindner, N. Chen, S. C.-T. Lam, and L. F. Lau The Angiogenic Factor Cysteine-Rich 61 (CYR61, CCN1) Supports Vascular Smooth Muscle Cell Adhesion and Stimulates Chemotaxis through Integrin {alpha}6{beta}1 and Cell Surface Heparan Sulfate Proteoglycans Endocrinology, April 1, 2002; 143(4): 1441 - 1450. [Abstract] [Full Text] [PDF]

				M. Ishigami, D. K. Swertfeger, M. S. Hui, N. A. Granholm, and D. Y. Hui Apolipoprotein E Inhibition of Vascular Smooth Muscle Cell Proliferation but Not the Inhibition of Migration Is Mediated Through Activation of Inducible Nitric Oxide Synthase Arterioscler Thromb Vasc Biol, April 1, 2000; 20(4): 1020 - 1026. [Abstract] [Full Text] [PDF]

				L. Paka, I. J. Goldberg, J. C. Obunike, S. Y. Choi, U. Saxena, I. D. Goldberg, and S. Pillarisetti Perlecan Mediates the Antiproliferative Effect of Apolipoprotein E on Smooth Muscle Cells. AN UNDERLYING MECHANISM FOR THE MODULATION OF SMOOTH MUSCLE CELL GROWTH? J. Biol. Chem., December 17, 1999; 274(51): 36403 - 36408. [Abstract] [Full Text] [PDF]

				T. Christen, M.-L. Bochaton-Piallat, P. Neuville, S. Rensen, M. Redard, G. van Eys, and G. Gabbiani Cultured Porcine Coronary Artery Smooth Muscle Cells : A New Model With Advanced Differentiation Circ. Res., July 9, 1999; 85(1): 99 - 107. [Abstract] [Full Text] [PDF]

				D. K. Swertfeger and D. Y. Hui Apolipoprotein E Receptor Binding Versus Heparan Sulfate Proteoglycan Binding in Its Regulation of Smooth Muscle Cell Migration and Proliferation J. Biol. Chem., June 29, 2001; 276(27): 25043 - 25048. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Koyama, N. Articles by Clowes, A. W. Search for Related Content PubMed PubMed Citation Articles by Koyama, N. Articles by Clowes, A. W.