This Article Abstract 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 Pickering, J. G. Articles by Chan, B. M. C. Search for Related Content PubMed PubMed Citation Articles by Pickering, J. G. Articles by Chan, B. M. C.

(Circulation Research. 1997;80:627-637.)
© 1997 American Heart Association, Inc.

Articles

Fibroblast Growth Factor-2 Potentiates Vascular Smooth Muscle Cell Migration to Platelet-Derived Growth Factor

Upregulation of ₂ß₁ Integrin and Disassembly of Actin Filaments

J. Geoffrey Pickering, Shashi Uniyal, Carol M. Ford, Thu Chau, Mary Ann Laurin, Lawrence H. Chow, Christopher G. Ellis, Jonathan Fish, , Bosco M. C. Chan

From the John P. Robarts Research Institute and London Health Sciences Centre, Departments of Medicine (Cardiology) (J.G.P., C.M.F., M.A.L., L.H.C.), Medical Biophysics (J.G.P., C.G.E., J.F.), Biochemistry (J.G.P.), and Microbiology and Immunology (S.U., T.C., B.M.C.C.), University of Western Ontario, London, Canada.

Correspondence to J. Geoffrey Pickering, London Health Sciences Centre, 339 Windermere Rd, London, Ontario N6A 5A5, Canada. E-mail gpickrng{at}rri.uwo.ca

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Abstract Fibroblast growth factor-2 (FGF-2) has been implicated in vascular smooth muscle cell (SMC) migration, a key process in vascular disease. We demonstrate here that FGF-2 promotes SMC motility by altering ß₁ integrin–mediated interactions with the extracellular matrix (ECM). FGF-2 significantly increased surface expression of ₂ß₁, ₃ß₁, and ₅ß₁ integrins on human SMCs, as assessed by flow cytometry. The greatest increase was for the collagen-binding ₂ß₁ integrin. Despite this, FGF-2 did not increase SMC adhesion to type I collagen but instead promoted SMC elongation and SMC motility. The latter was evaluated by using a microchemotaxis chamber and by digital time-lapse video microscopy. Although FGF-2 was not chemotactic for human SMCs, cells preincubated with FGF-2 displayed a 3.1-fold increase in migration to the undersurface of porous type I collagen–coated membranes and a 2.1-fold increase in migration speed on collagen. Furthermore, chemotaxis to platelet-derived growth factor-BB on collagen was significantly greater in SMCs exposed to FGF-2. FGF-2–induced elongation and migration on collagen were inhibited by a blocking anti-₂ß₁ antibody; however, SMC adhesion to collagen was unaffected. SMC migration on fibronectin was also enhanced by FGF-2, although less prominently: migration through porous membranes increased 1.8-fold, and migration speed increased 1.3-fold. Also, FGF-2 completely disassembled the smooth muscle -actin–containing stress fiber network contemporaneously with the change in integrin expression and cell shape. We conclude that (1) exogenous FGF-2 promotes SMC migration and potentiates chemotaxis to PDGF-BB; (2) the promigratory effect of FGF-2 is especially prominent on type I collagen and is mediated by upregulation of ₂ß₁ integrin; and (3) FGF-2 disassembles actin stress fibers, which may promote differential utilization of ₂ß₁ integrin for motility but not adhesion. This dynamic SMC-ECM interplay may be an important mechanism by which FGF-2 facilitates SMC motility in vivo.

Key Words: smooth muscle cell • integrin • collagen • actin • fibroblast growth factor

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Migration of vascular SMCs is an important means by which the arterial intima thickens during the course of vascular diseases.^{1 2} The mechanisms mediating SMC migration are incompletely understood, but potential regulators include chemotactic cytokines and growth factors³ as well as ECM molecules.^{4 5 6 7} A number of physical changes to the cell and surrounding ECM are also features of cell migration, including restructuring of the actin cytoskeleton,⁸ dynamic attachment to and release from the ECM,^{9 10} and local turnover of the ECM.^{11 12}

FGF-2 (basic FGF) is a potent mitogen for SMCs^{13 14} and a proven mediator of SMC proliferation after injury to the rat carotid artery.¹⁵ A role for FGF-2 in mediating SMC migration has also been suggested by in vivo studies. Injection of FGF-2 following gentle deendothelialization of the rat carotid artery significantly stimulated SMC migration from the media to the intima. Furthermore, SMC migration induced by balloon disruption of the arterial media was abrogated by a blocking antibody to FGF-2.¹⁶ However, the mechanism by which FGF-2 mediates SMC migration is unclear. Unlike PDGF, which is a potent chemoattractant for SMCs,^{3 17} FGF-2 has no,³ or at best weak,^{17 18} chemotactic activity for SMCs. A potential role for FGF-2 in modulating the interactions between SMCs and the ECM has been suggested¹⁶ but not explored.

Integrins are cell surface receptors that serve to bridge the ECM with the cell cytoskeleton.¹⁹ In SMCs, integrins of the ß₁ family appear to be the dominant integrins responsible for cell adhesion to the ECM, and several ß₁ integrins have been identified on SMCs.^{10 20 21 22 23 24 25} Migration of SMCs has been noted to be dependent on ß₃ integrins,^{26 27} but a number of recent studies have also implicated ß₁ integrins in SMC migration. Antisera against ß₁ integrin blocked fibronectin-promoted migration of rat SMCs,²⁷ and ₂ß₁ integrin has been implicated in PDGF-induced chemotaxis on collagen.²⁵ On the other hand, excessive ß₁ integrin–mediated adhesion may impair SMC migration, as demonstrated by recent studies with a function-activating anti–ß₁ integrin antibody.^{10 28}

Expression of ß₁ integrins is regulatable by growth factors. Transforming growth factor-ß₁ upregulated ₁ and ₅ integrin subunits in rabbit SMCs,²⁹ and PDGF-BB has been reported to increase the ₅ integrin subunit in bovine and rabbit SMCs.^{29 30} Klein et al³¹ have demonstrated that FGF-2 increases the expression of ₂ß₁, ₃ß₁, ₅ß₁, and ₆ß₁ integrins in microvascular endothelial cells. However, the effect of FGF-2 on ß₁ integrin expression in vascular SMCs is unknown.

In the present study, we have examined the mechanism by which FGF-2 controls migration of human SMCs, focusing particularly on the interplay between SMCs and the ECM. Using flow cytometry, we found that FGF-2 upregulates the expression of a number of ß₁ integrins, most prominently the collagen-binding ₂ß₁ integrin. Associated with this altered integrin repertoire was morphological elongation of the cell and complete disassembly of the actin stress fiber–containing cytoskeleton. Consistent with other reports,³ FGF-2 was not chemotactic for human SMCs. However, SMCs treated with FGF-2 displayed enhanced migration on type I collagen, including augmented chemotaxis to PDGF-BB, that was inhibited by a blocking antibody to ₂ß₁ integrin. This ₂ß₁ integrin–mediated increase in migration was not accompanied by an increase in adhesion to collagen, suggesting a complex response to FGF-2 designed to selectively enhance motility. The findings indicate that coordinated alterations in the ß₁ integrin–cytoskeletal axis may be a critical means by which FGF-2 promotes SMC migration and potentiates chemotaxis to PDGF.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Antibodies and Reagents
mAbs used included the ₁ß₁ integrin–specific mAb, Ts2/7³² ; the ₂ß₁ integrin–specific mAb, BHA2.1³³ (Chemicon Inc); the ₃ß₁ integrin–specific mAb, P1B5 (GIBCO/BRL); an ₅ß₁ integrin–specific mAb, HA5; a ß₁ integrin subunit–specific mAb, HB1.1(Chemicon Inc); and an _v integrin subunit–specific mAb, 23C6 (Serotec). Isotype-matched control antibodies were used for flow cytometry and blocking experiments. The F(ab')₂ fragment of FITC-conjugated anti-mouse antibody was purchased from Becton-Dickinson.

Human recombinant FGF-2 and PDGF-BB were purchased from GIBCO/BRL. Rat-tail collagen solubilized in acetic acid was used as a source of type I collagen and was prepared as previously described.³⁴ Human fibronectin was isolated from human plasma by gelatin-Sepharose chromatography,³⁵ and EHS tumor–derived laminin was obtained from GIBCO/BRL.

SMC Culture
Primary cultures of human arterial SMCs were initiated by explant outgrowth from unused segments of internal mammary artery retrieved at the time of coronary artery bypass surgery, as previously described.^{36 37} The identity of vascular SMCs was confirmed morphologically and by positive immunostaining for smooth muscle -actin (clone 1A4, Dako). Cells were grown in medium (M199, GIBCO/BRL) supplemented with the designated concentration of FBS. All experiments were performed using human SMCs in the third or fourth subculture.

Immunofluorescence
SMCs seeded on to multiwell slides were fixed in ice-cold acetone and stained for smooth muscle -actin (clone 1A4, Dako) as previously described.³⁶ F-actin stress fibers were also visualized using FITC-conjugated phalloidin (Sigma Chemical Co). Cells were coverslipped using glycerol/PBS (9:1) containing Hoechst 33258 (2.5 µg/mL, Sigma) to identify cell nuclei and evaluated by fluorescence microscopy.

Flow Cytometry
Flow cytometry for detection of integrin expression was carried out by indirect immunofluorescence staining as described previously.^{33 38} Early confluent SMCs were trypsinized and washed in cold PBS with 1% BSA. Cells were incubated for 30 minutes on ice with control or integrin-specific mAbs at predetermined saturating concentrations. Washed cells were incubated with FITC-labeled anti-mouse F(ab')₂ fragment, and fluorescence staining was analyzed using a Becton-Dickinson FacScan.

SMC Adhesion Assay
Cell adhesion was assessed using SMCs labeled with BCECF (Sigma), as described previously.^{33 39} Briefly, 5x10⁴ BCECF-labeled cells were allowed to adhere to matrix-coated wells for 20 minutes at 37°C. Unbound cells were removed by gentle washing. Bound fluorescence was measured by a Fluorescence Concentrator Analyzer (IDEXX). Bound fluorescence from adhesion to BSA-coated wells served as control for background adhesion. Net fluorescence was determined by subtraction of background and expressed as a percentage of the total fluorescence yielded from 5x10⁴ labeled SMCs minus background fluorescence. For inhibition experiments, mAb BHA2.1 was used at 10 to 20 µg/mL, a concentration based on prior titering to inhibit adhesion of HT1080 cells to collagen.³³

SMC Elongation Assay
SMC shape change after seeding was evaluated using a Boyden-type microchemotaxis chamber (Neuroprobe, Cabot John), as previously described.²⁸ Polycarbonate filters (Nucleopore) were precoated overnight at 4°C with 10 µg/mL of either fibronectin or collagen type I in PBS. Control or FGF-2–treated SMCs were washed in PBS and trypsinized, and 2000 SMCs (40 000 cells/mL in DME with 1% BSA) were placed in the upper well. The lower well was filled with the same medium with or without FGF-2. The chamber was incubated at 37°C for 2 hours, and SMCs adherent to the upper surface of the filter were fixed in methanol and stained with hematoxylin. Prior experiments demonstrated that cell migration to the undersurface of the membrane was negligible (<1 cell/HPF) under these conditions for both control and FGF-2–treated SMCs. The lengths of the major and minor axes transecting the nucleus were measured from digitized calibrated images, using morphometry software (Jandel Scientific).

SMC Migration Assay
Bulk migration was measured using a microchemotaxis chamber, as previously described.^{25 40} Control or FGF-2–treated SMCs were washed in PBS and trypsinized, and 25 000 cells (500 000 cells/mL in DME with 1% BSA) were added to the upper well of the microchemotaxis chamber. Polycarbonate filters with 10-µm pores were precoated with fibronectin or collagen type I as described above. The lower well of the chamber was filled with DME with 1% BSA with or without growth factor. Migration was allowed to proceed for 6 hours at 37°C under 5% CO₂. Cells remaining on the upper surface of the filters were mechanically removed, and then filters were fixed in methanol and stained with Harris's hematoxylin. The number of cells that had migrated to the lower surface was determined by counting under high-power microscopy (x40 objective). All conditions for a given experiment were performed in triplicate.

Time-Lapse Video Microscopy
SMC migration speed was evaluated using a digital time-lapse video recording system. Cells incubated with or without FGF-2 were seeded onto culture dishes coated with matrix proteins at 10 µg/mL. Migration was monitored with an inverted microscope (Diaphot 300, Nikon) using a x10 objective and a halogen light source. A CCD video camera (C72, Dage-MTI) attached to the microscope was used to generate video images, which were digitally acquired over a 5- to 8-hour recording period using a Silicon Graphics Indy workstation and custom-written time-lapse software. Ambient temperature was maintained at 37°C by mounting the culture dish in a Styrofoam sleeve with transparent heating elements (MINCO Products, Inc) placed above and below the dish. Cells were incubated in bicarbonate-reduced medium (M199 with Hanks' salts and 25 mmol/L HEPES, GIBCO/BRL) to maintain physiological pH in room air. Migration was measured from digital images by tracking the location of cell centroids at hourly intervals. Migration speed was determined as the sum of hourly distances divided by the total time, as described previously.⁴¹

Statistics
Data are expressed as mean±SD. Comparisons were made by t test or ANOVA with Scheffé's post hoc test. Statistical significance was set at P<.05.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
FGF-2 Increases Expression of ß₁ Integrins in SMCs
To evaluate the effect of FGF-2 on ß₁ integrin expression in human SMCs, early confluent cells were incubated in starvation medium (M199, 1% FBS) overnight and stimulated with FGF-2 (0 to 25 ng/mL) for 48 hours. Expression of ß₁ integrins was examined by performing flow cytometry. As shown in Fig 1, the total ß₁ integrin cell surface pool, detected using the pan-ß₁ integrin mAb HB1.1, was greater in SMCs treated with 25 ng/mL FGF-2 than in untreated control SMCs. However, the effect of FGF-2 on expression of individual ß heterodimers differed. FGF-2 produced either no effect or a slight reduction in ₁ß₁ integrin level, whereas it increased the expression of ₂ß₁, ₃ß₁, and ₅ß₁ integrins. Expression of these integrins in cells exposed to 10 ng/mL FGF-2 was intermediate, indicating that the effect was dose dependent (data not shown). ₄ß₁ integrin was not detected in the cultured human SMCs, similar to the results of Skinner et al.²⁵ Expression of the _v integrin subunit was detected and was increased by FGF-2 treatment (data not shown).

View larger version (28K): [in this window] [in a new window]   Figure 1. A, Expression of ß1 integrins in human SMCs cultured in the presence or absence of FGF-2. SMCs were incubated with specific anti-integrin antibodies for 45 minutes, labeled with FITC-conjugated anti-mouse IgG, and analyzed by flow cytometry. Anti-integrin antibodies used were as follows: HB1.1 (anti-ß1), TS2/7 (anti-1ß1), BHA2.1 (anti-2ß1), P1B5 (anti-3ß1), and HA5 (anti-5ß1). P3 was used as an isotype-matched control IgG, and the corresponding fluorescence signal appears on the left of each histogram. Of the other two tracings, the thin and the thick lines depict the specific antibody-bound signal from SMCs incubated for 48 hours with vehicle or 25 ng/mL FGF-2, respectively. B, Time course of FGF-2–mediated expression of 2ß1 integrin by SMCs. Cells were cultured with 25 ng/mL FGF-2 for the designated length of time and harvested simultaneously. Integrin expression was assessed by flow cytometry using mAb BHA2.1, and the net mean fluorescence index (MFI) was determined. Graph is representative of two separate experiments. C, Effect of ECM substrate on FGF-2–mediated expression of 2ß1 integrin. Cells were seeded onto culture dishes coated with fibronectin, laminin, or type I collagen, and FGF-2 (25 ng/mL) was simultaneously added. After 48 hours, integrin expression was evaluated by flow cytometry. The absolute values for the MFI are plotted. Two further experiments showed the same findings.

The quantitatively greatest change in expression induced by FGF-2 was observed for ₂ß₁ integrin; thus, this response was characterized further. The kinetics of the increase in ₂ß₁ integrin was evaluated by harvesting SMCs at various times after the addition of 25 ng/mL FGF-2. As shown in Fig 1B, increased expression was detected at 12 hours and was maximal by 35 hours.

₂ß₁ integrin is known to bind collagens and laminin.¹⁹ To determine if the ability of FGF-2 to increase ₂ß₁ integrin surface expression was dependent on specific interaction with the ECM, SMCs were seeded onto dishes coated with either fibronectin, laminin, or type I collagen and stimulated for 48 hours with FGF-2. The relative increase in ₂ß₁ integrin expression induced by FGF-2 was not different for the three substrates: 3.7±0.7-fold for cells on fibronectin, 2.2±0.5-fold for cells on laminin, and 2.4±1.3-fold for cells on type I collagen (P=NS). However, in absolute terms, SMCs seeded on collagen expressed more ₂ß₁ integrin under both basal and FGF-2–stimulated conditions than did cells seeded on fibronectin or laminin (P<.05) (Fig 1C).

FGF-2 Increases SMC Elongation but Not Adhesion to Type I Collagen via ₂ß₁ Integrin
As shown in Fig 2, neither acute exposure to FGF-2 (25 ng/mL) nor 48 hours of treatment with FGF-2 at this dose had a significant effect on SMC adhesion to type I collagen. This was interesting in light of the observation that FGF-2 increased surface expression of the collagen-binding ₂ß₁ integrin. Of note, however, addition of mAb BHA2.1, which blocks ₂ß₁ integrin function,³³ had no detectable effect on adhesion of SMCs to collagen in either untreated or FGF-2–treated SMCs (Fig 2).

View larger version (59K): [in this window] [in a new window]   Figure 2. Effect of FGF-2 on SMC adhesion to type I collagen. Untreated SMCs and SMCs exposed acutely or for 48 hours to 25 ng/mL FGF-2 were allowed to adhere for 20 minutes in the presence of either control IgG (P3, 10 µg/mL) or mAb BHA2.1 (10 µg/mL). The proportion of adherent cells was determined as described in "Materials and Methods" and was not significantly different among the various conditions.

The effect of FGF-2 on the morphology of SMCs was assessed first by evaluating changes in cell shape over 48 hours. As shown in Fig 3, SMCs incubated with 25 ng/mL FGF-2 acquired a distinct spindle-like shape that was evident after 24 hours of treatment and most prominent after 48 hours. This shift to a spindle-like morphology was observed in SMCs growing on collagen type I, fibronectin, or laminin. To quantitatively examine the effect of FGF-2 on SMC elongation, SMCs were treated with FGF-2 (25 ng/mL) for 48 hours and then trypsinized and allowed to spread on matrix-coated filters for 2 hours. The length and width of 80 SMCs were determined for each condition, and data were obtained in triplicate. As shown in Fig 3C and 3D, the ratio of length to width was significantly higher in SMCs treated with FGF-2, indicating that FGF-2 promoted elongation of SMCs. This effect of FGF-2 was observed for cells seeded on collagen (P<.05) and fibronectin (P<.01). mAb BHA2.1 inhibited the FGF-2–mediated increase in elongation of SMCs on collagen (P<.05) but had no effect on elongation on fibronectin (Fig 3). Further evidence for the role of ₂ß₁ integrin in mediating shape changes came from visual assessment of the SMCs seeded onto matrix. FGF-2–treated SMCs seeded on collagen with mAb BHA2.1 displayed atypical thin cytoplasmic processes in an arborized pattern, suggesting a failed attempt at reshaping the cell (data not shown).

View larger version (78K): [in this window] [in a new window]   Figure 3. Effect of FGF-2 on SMC morphology. A and B, Phase-contrast photomicrographs of control cells (A) and cells treated for 48 hours with 25 ng/mL FGF-2 (B). C and D, Ratio of length to width 2 hours after seeding onto type I collagen (C) or fibronectin (D). SMCs were seeded in the presence of either control IgG (P3, 10 µg/mL) or mAb BHA2.1 (10 µg/mL), and measurements were taken from 80 cells for each condition. Data represent mean of triplicate values.

FGF-2 Disassembles the Actin Stress Fiber Cytoskeleton
Actin microfilaments are a major component of the cytoskeleton and are known to interact with integrins on the inner face of the cell membrane. To determine if FGF-2 influenced cytoskeletal organization, and hence its relationship with integrins in focal adhesions, SMCs were fixed and immunostained for smooth muscle -actin. As shown in Fig 4, there was a gradual loss of smooth muscle -actin microfilament bundles traversing the cell body. This was evident by 24 hours and complete in almost all SMCs after 48 to 72 hours. The disassembly of smooth muscle -actin–containing stress fibers was seen in cells growing on either collagen or fibronectin. Furthermore, the response did not require ₂ß₁ integrin function, as mAb BHA2.1 failed to prevent actin disassembly. Similar results were seen when F-actin was visualized using FITC-phalloidin (data not shown).

View larger version (49K): [in this window] [in a new window]   Figure 4. Fluorescence micrographs of SMCs cultured on either type I collagen (A through D) or fibronectin (E through H) and immunostained for smooth muscle -actin. A and E, Untreated SMCs. B and F, SMCs incubated with 25 ng/mL FGF-2 for 24 hours. C and G, SMCs incubated with 25 ng/mL FGF-2 for 48 hours in the presence of control IgG (P3, 10 µg/mL). D and H, SMCs incubated with 25 ng/mL FGF-2 for 48 hours in the presence of mAb BHA2.1 (20 µg/mL).

FGF-2 Is Not Chemotactic for Human SMCs but Enhances Migration Through Type I Collagen–Coated Membranes and Increases Migration Speed on Type I Collagen
Having identified that FGF-2 alters ß₁ integrin expression, SMC shape, and the actin cytoskeleton, we determined whether SMCs incubated with FGF-2 would also manifest altered migratory properties. Control SMCs and SMCs treated for 48 hours with 25 ng/mL FGF-2 were seeded onto ECM-coated porous polycarbonate filters, and migration to the lower surface was evaluated after 6 hours. For these experiments, FGF-2 (0 to 25 ng/mL) was placed in the lower chamber. As illustrated in Fig 5, several observations were made. First, there was no evidence for chemotaxis of human SMCs toward FGF-2; ie, for a given set of experimental conditions, the number of cells that migrated to the undersurface of the membrane when FGF-2 was in the lower well was not different than when vehicle alone was in the lower chamber. This lack of chemotaxis to FGF-2 was observed with SMCs seeded on membranes precoated with fibronectin or type I collagen. PDGF-BB (10 µg/mL) served as a positive control for chemotaxis and produced SMC migration when placed in the bottom but not upper well of the chamber (eg, see Fig 8). FGF-2 in the upper well also had no effect on migration. A second observation was that SMCs pretreated with FGF-2 migrated through collagen-coated filters in significantly greater numbers than did untreated control SMCs. The number of FGF-treated SMCs that migrated to the lower surface, in the absence of growth factor in the lower chamber, was 3.1±0.8-fold greater than control SMCs (P<.001, n=5 experiments). Similar results were obtained when FGF-2 (1 to 25 ng/mL) was present in the bottom well of the chamber. The third observation was that the FGF-2–induced augmentation of migration was more prominent for SMCs on type I collagen than on fibronectin. Migration through fibronectin-coated membranes was increased 1.8±0.2-fold by FGF-2 pretreatment (P<.05, n=3 experiments), but this increase was significantly less than that for SMCs migrating on collagen-coated membranes (P<.05).

View larger version (30K): [in this window] [in a new window]   Figure 5. Effects of FGF-2 on SMC migration through type I collagen-coated (top) and fibronectin-coated (bottom) polycarbonate membranes. Before the migration assay, SMCs were incubated for 48 hours with or without 25 ng/mL FGF-2. The migration assay was performed in a chemotaxis chamber with designated concentrations of FGF-2 in the lower well. Results are representative of three experiments.

View larger version (41K): [in this window] [in a new window]   Figure 8. Effect of FGF-2 on PDGF-induced chemotaxis of SMCs on type I collagen. Untreated SMCs and cells treated for 48 hours with 25 ng/mL FGF-2 were placed in the upper well of the migration chamber together with either control IgG (P3, 10 µg/mL) or mAb BHA2.1 (10 µg/mL). PDGF-BB (0 to 30 ng/mL) was in the lower well, and the number of cells that migrated to the lower membrane surface was counted after 6 hours. *P<.01 vs untreated SMCs incubated with control IgG, untreated SMCs incubated with mAb BHA2.1, and FGF-2 treated SMCs incubated with mAb BHA2.1; P<.01 vs untreated SMCs incubated with mAb BHA2.1 and FGF-2–treated SMCs incubated with mAb BHA2.1.

To directly measure the speed of SMC migration, SMC movement was monitored by digital time-lapse video microscopy. The results of image quantification including representative migration paths are shown in Fig 6. SMCs preincubated for 48 hours with FGF-2 migrated on collagen at speeds that, on average, were 2.1-fold higher than those of untreated SMCs (19.4±5.2 versus 9.3±3.8 µm/h, P<.05). FGF-2–treated SMCs also migrated on fibronectin faster than did untreated SMCs, although the relative increase was only 1.3-fold (11.2±0.4 versus 8.4±0.8 µm/h, P<.05). These findings thus corroborate those of the chemotaxis chamber assay and implicate augmented crawling speed, likely together with cell shape changes, as components of the enhanced migratory response of SMCs to FGF-2.

View larger version (23K): [in this window] [in a new window]   Figure 6. Effect of FGF-2 on SMC migration path and migration speed on type I collagen and fibronectin. SMCs treated with FGF-2 (25 ng/mL) or vehicle for 48 hours were monitored by digital time-lapse video microscopy. Migration paths (A through D) depict hourly translocations of eight cells per experimental condition, over a total of 5 hours. Migration speed over the monitoring period (E) was determined for each cell, and the results are the mean of three separate experiments on each of collagen and fibronectin. *P<.05 vs untreated SMCs on collagen; P<.01 vs untreated SMCs on fibronectin.

FGF-2–Mediated Enhancement of SMC Migration on Collagen Is Inhibited by Anti–₂ß₁ Integrin Antibody
Although inhibition of ₂ß₁ integrin had no significant effect on SMC adhesion to collagen, ₂ß₁ integrin function was important for FGF-2–stimulated morphological changes. Therefore, the role of ₂ß₁ integrin in mediating the FGF-2–induced increase in SMC migration was evaluated using the blocking mAb BHA2.1. As shown in Fig 7, the addition of 10 µg/mL mAb BHA2.1 to the upper well prevented the FGF-2–mediated increase in migration through collagen-coated membrane (P<.01), whereas the isotype-matched control antibody had no effect. The same dose of mAb BHA2.1 had no effect on basal or FGF-2–enhanced migration through fibronectin-coated membranes.

View larger version (37K): [in this window] [in a new window]   Figure 7. Effect of mAb BHA2.1 (anti–2ß1 integrin) on FGF-2–mediated SMC migration on collagen-coated (top) and fibronectin-coated (bottom) polycarbonate membranes. Untreated SMCs and cells treated for 48 hours with 25 ng/mL FGF-2 were placed in the upper well of the migration chamber together with either control IgG (P3, 10 µg/mL) or mAb BHA2.1 (10 µg/mL). Cells that migrated to the lower membrane surface were counted after 6 hours. *P<.01 vs untreated SMCs; P<.01 vs FGF-2–treated SMCs in the presence of control IgG.

FGF-2 Enhances Chemotaxis to PDGF via ₂ß₁ Integrin
Both FGF-2¹⁶ and PDGF¹⁷ have been found to regulate SMC migration in vivo. Therefore, we next determined if exposure of SMCs to exogenous FGF-2 altered the chemotactic response to PDGF. As illustrated in Fig 8, PDGF induced dose-dependent chemotaxis through collagen-coated filters, as has been previously described.^{3 25} When cells were pretreated for 48 hours with 25 ng/mL FGF-2, the migratory response to 1 and 10 ng/mL PDGF was significantly increased (P<.01). At these doses of PDGF, migration of SMCs not preincubated with FGF-2 was unaffected by the anti–₂ß₁ integrin mAb BHA2.1. However, mAb BHA2.1 blocked the incremental increase provided by FGF-2 treatment (P<.01). Thus, FGF-2 treatment potentiated the chemotactic response to PDGF on collagen, by effectively supplementing an ₂ß₁ integrin–independent process with an ₂ß₁ integrin–dependent process. The response to high-dose PDGF (30 ng/mL) differed in that FGF-2 did not augment the chemotactic response in this setting. Furthermore, migration of SMCs not preincubated with FGF-2 to high-dose PDGF was partially inhibited by mAb BHA2.1 (P<.01), indicating that an ₂ß₁-mediated mechanism already existed.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
A role for FGF-2 in vascular SMC migration has been demonstrated in cell culture⁴² and in vivo.¹⁶ However, the mechanism by which FGF-2 promotes SMC migration is not well defined. A number of studies,^3,17,18 including the present report, indicate that FGF-2 provides chemotactic stimulus to SMCs; no, or at best a weak induction of directed migration is therefore unlikely to be a mechanism of action. Induction of endogenous FGF-2 has been implicated in migration to PDGF,⁴² and Bilato et al¹⁸ recently found that FGF-2 was necessary to activate calcium/calmodulin-dependent protein kinase II, which is required for PDGF-mediated chemotaxis. The data presented here provide evidence for a very different mechanism by which FGF-2 promotes SMC migration, namely, by altering the nature of the interaction between SMCs and the ECM. The promigratory effect of FGF-2 was apparent after sustained exposure and was chemokinetic in nature, since a gradient of FGF-2 was not required for the effect. The effect was especially pronounced for SMCs migrating on a substrate of type I collagen. As well, exposure to FGF-2 served to potentiate directed migration of SMCs (chemotaxis) to PDGF-BB on this substrate. This enhanced motility was mediated by ₂ß₁ integrin, the expression of which was increased by over 2-fold by FGF-2. Finally, FGF-2–induced motility was also associated with postintegrin alterations in the cytoskeleton, including disassembly of actin stress fibers.

The regulation of cell migration by growth factors is often considered and studied in a time frame of a few hours. SMC chemotaxis, for example, typically is manifested in vitro within 4 to 6 hours.^{12 18 25} However, SMC migration within the arterial wall follows a more prolonged time course. Indeed, the rationale for examining the consequences of a more prolonged exposure to FGF-2 comes from in vivo studies. After balloon injury to the rat carotid artery, SMC proliferation begins after 24 hours,⁴³ and SMC migration is detectable by 3 days.¹⁶ Both responses have been shown to be regulated by FGF-2,^{15 16} which implies that in this setting SMCs may be exposed to FGF-2 over a matter of days. The fact that FGF-2 binds with low affinity to the ECM is also compatible with sustained delivery of FGF-2 to the SMC, during conditions of active remodeling.⁴⁴

FGF-2 increased the surface expression of several ß₁ integrins, including ₂ß₁, ₃ß₁, and ₅ß₁ integrins. However, the effect was not generalized, since ₁ß₁ integrin expression was not increased. The quantitatively greatest increase in expression was seen for ₂ß₁ integrin. ₂ß₁ integrin is capable of binding collagen types I and IV as well as laminin,¹⁹ and there is evidence that expression of ₂ß₁ integrin is dependent on SMC phenotype. Specifically, expression of ₂ß₁ integrin was not observed in SMCs within the aortic media but was induced after growth in culture, suggesting that ₂ß₁ integrin expression is a feature of an "activated" SMC phenotype.²⁵ Although one study did not detect ₂ß₁ integrin after injury to the rat carotid artery,⁴⁵ neoexpression of ₂ß₁ integrin has been identified in human SMC tumors.⁴⁶ The effect of growth factors on ₂ß₁ integrin expression by SMCs has not previously been reported, although it is of interest that FGF-2 has been shown to increase ₂ß₁ integrin expression in microvascular endothelial cells.³¹

It was noteworthy that the FGF-2–mediated increase in the expression of ₂ß₁ integrin was not associated with increased adhesion to collagen but was associated with cell elongation and increased motility on this substrate. Under basal or FGF-2–stimulated conditions, adhesion to collagen was not inhibited by mAb BHA2.1, an anti-₂ß₁ antibody previously shown to block adhesion to type I collagen.³³ Similar results were obtained using another ₂ß₁ integrin mAb (data not shown). Skinner et al²⁵ also did not observe a role for ₂ß₁ integrin on SMC attachment to type I collagen. On the other hand, Lee et al²⁴ observed a 31% decrease in adhesion of human venous SMCs to type I collagen by an anti–₂ß₁ integrin antibody, implying that the ₂ß₁ receptor is capable of at least partly mediating adhesion of SMCs to type I collagen. Other receptors besides ₂ß₁ integrin may be involved in binding to type I collagen, including ₁ß₁ and ₃ß₁ integrins,^{19 38} and cooperativity has been demonstrated between ₁ß₁ and ₂ß₁ integrins.²⁴ Therefore, it is conceivable that inhibition of ₂ß₁ integrin alone may be insufficient to impair adhesion and that inhibition of multiple receptors is necessary before a functional consequence on adhesion is manifested. Even so, it is of interest that in the face of a 2.4-fold increase in ₂ß₁ integrin expression, FGF-2 did not increase SMC adhesion to collagen whatsoever. FGF-2 did, however, promote SMC elongation and migration on collagen in a manner that was dependent on the increase in ₂ß₁ integrin. The ability of the cell to modulate its shape and extend over its substrate is fundamental to locomotion and morphogenesis, and a role for ₂ß₁ integrin in these processes has been suggested in a few cell lines.^{47 48} For example, in a mammary epithelial cell line, inhibition of ₂ß₁ integrin significantly reduced cell spreading and migration speed as well as the ability of these cells to wrap around collagen fibers.⁴⁷ The present studies support a similar role for ₂ß₁ integrin in human SMCs, specifically in the setting of sustained stimulation by FGF-2.

The ability of FGF-2 to induce integrin-mediated changes in cell shape without augmenting adhesivity may be precisely the scenario required to optimize cell locomotion. Theoretical analysis has suggested that maximum migration speed will occur at an intermediate adhesiveness,⁴⁹ and subsequent experimental evaluation of SMC migration on fibronectin and type IV collagen has supported this concept.⁵ Furthermore, using an anti–ß₁ integrin antibody that augments the binding affinity of ß₁ integrins, Seki et al¹⁰ and Koyama et al²⁸ observed enhanced adhesion of SMCs to ECM substrates but decreased SMC migration to PDGF on Matrigel. Thus, if integrin-mediated adhesion is excessive, the cell may be effectively "frozen" in location. Therefore, it appears that utilization of ₂ß₁ integrin for shape change (elongation) but not for adhesion is a coordinated response to FGF-2 designed to facilitate motility.

What is the potential mechanism by which a FGF-2–mediated increase in ₂ß₁ integrin increases SMC elongation and migration but does not increase adhesion to collagen? Presumably, there are other effects of FGF-2 that effectively dissociate the expression of ₂ß₁ integrin from its adhesive function. In this regard, it was noteworthy that incubation of SMCs with FGF-2 led to disassembly of the actin stress fiber network, evidenced by staining for smooth muscle -actin and F-actin. It is well established that integrins associate with the actin-based cytoskeleton,¹⁹ including the actin stress fibers that traverse the cell. These actin microfilament bundles are structurally linked to integrins in focal adhesion contacts, and their presence is felt to be an important feature of the stationary cell phenotype⁵⁰ ; eg, actin stress fibers are prominent in corneal fibroblasts attached and flattened over a planar substrate but are absent in fibroblasts migrating in three-dimensional collagen gels⁵¹ or in vivo.⁵² In vivo, actin stress fibers are present in endothelial cells subjected to high shear stress,⁵³ where strong attachment to the underlying ECM is required. Given the association between actin stress fiber network and strong surface adhesion, disassembly of actin stress fibers may be one means by which the cell liberates itself for movement. We speculate that disassembly of actin stress fibers in SMCs by FGF-2 minimizes or even prevents increased adhesivity in the setting of increased integrin expression, by disrupting the intracellular structural network required for stable attachment. Other possible mechanisms underlying the differential response include selective activation of intracellular signaling cascades that mediate migration but not adhesion or selective modulation of integrin function or activation state through conformational changes in specific receptor domains.^{10 54}

Disassembly of actin stress fibers by FGF-2 was observed in cells cultured on collagen as well as on fibronectin, and the process did not require ₂ß₁ integrin function. Therefore, this particular effect of FGF-2 does not appear to be specific for a given ECM environment. However, actin disassembly could reflect an overall reduced level of interaction between the cell and the ECM. It has recently been shown that cellular interaction with ECM is necessary for stress fiber attachment to focal adhesion contacts.⁵⁵ Furthermore, we have recently determined that FGF-2 induces expression of matrix metalloproteinase-1 (collagenase) in human SMCs.⁵⁶ This raises the possibility that local degradation of collagen, induced by FGF-2, may reduce the available ECM ligands for integrins, with disorganization of actin stress fibers as a downstream consequence. This remains speculative, and whether a similar degradation of fibronectin occurs in response to FGF-2 is not known. However, the interplay between ECM degradation and integrin function has recently been highlighted by the observation that signaling via ₂ß₁ integrin induces matrix metalloproteinase-1 expression.⁵⁷ Other potentially relevant mediators of stress fiber integrity include members of the Rho family of GTP-binding proteins.^{58 59} Some growth factors, such as PDGF and epidermal growth factor, induce stress fiber assembly via functional Rho.⁵⁸ The effect of FGF-2 on Rho function is unexplored.

Both FGF-2¹⁶ and PDGF¹⁷ have been shown to mediate SMC migration following balloon injury to the rat carotid artery. Previous studies have indicated that PDGF-mediated release of FGF-2 plays a role in chemotaxis to PDGF.^{18 42} However, during vigorous vascular remodeling, eg, after mechanical injury, FGF-2 may also be liberated from dead or injured cells as well as from ECM depot sites. The present findings of augmented chemotaxis to PDGF following FGF-2 exposure may be particularly relevant to such circumstances. By endowing the SMC with the capacity to migrate faster on collagen, FGF-2 induced a functional augmentation of PDGF-induced migration. Interestingly, this cooperative effect was dose dependent and apparent when SMCs were exposed to 1 to 10 ng/mL PDGF but not to high-dose PDGF (30 ng/mL). In the absence of FGF-2, SMC migration to the lower concentrations of PDGF was largely independent of ₂ß₁ integrin function. However, preincubation of SMCs with FGF-2 effectively supplemented an ₂ß₁ integrin–independent process with an ₂ß₁ integrin–dependent process. In contrast, migration to high-dose PDGF in the absence of FGF-2 was partly dependent on ₂ß₁ integrin, consistent with that reported by Skinner et al.²⁵ Under these circumstances, FGF-2 did not provide an additional effect, likely because the ₂ß₁-mediated component of migration had already been recruited. Other effects of PDGF relevant to cell motility have also been shown to depend on PDGF concentration. In particular, the effects of low-dose PDGF (5 ng/mL) on membrane ruffling, actin organization, and tyrosine phosphorylation of focal adhesion kinase and phosphatidylinositol 3-kinase were found to differ from those of high-dose (30 ng/mL) PDGF.⁶⁰ Furthermore, we have observed that high-dose PDGF induced a small increase in ₂ß₁ integrin expression, but this was not detectable at lower concentrations (data not shown).

Although the promigratory effect of FGF-2 was greatest for SMCs on collagen, an enhancement in migration was also seen with SMCs on fibronectin. The mechanism for this response is not presently defined; however, both disassembly of actin stress fibers and cell elongation occurred on fibronectin and may be contributing factors. Furthermore, integrins other than ₂ß₁ may be relevant. FGF-2 effected a modest increase in expression of ₅ß₁ integrin and the _v integrin subunit, both of which are capable of interacting with fibronectin.¹⁹ A role for _vß₃ integrin in SMC migration and vascular lesion formation has been previously suggested.^{61 62}

In summary, we have determined that FGF-2 increases the speed with which SMCs migrate and have identified two associated subcellular processes: upregulation of ₂ß₁ integrin and disassembly of the actin stress fiber network. The former process is necessary for the increase in migration on collagen; the latter may underlie the differential utilization of ₂ß₁ integrin for migration but not for adhesion. These coordinated changes in the ß₁ integrin–cytoskeletal axis may be a critical means by which FGF-2, and possibly other growth factors, regulate SMC migration in vivo. Furthermore, they provide a unique basis of cooperativity between PDGF and FGF-2 in the control of SMC migration.

	Selected Abbreviations and Acronyms

 

DME = Dulbecco's modified minimum essential medium ECM = extracellular matrix FGF = fibroblast growth factor HPF = high-power field mAb = monoclonal antibody PDGF = platelet-derived growth factor SMC = smooth muscle cell

	Acknowledgments

 
This study was supported by grants from the Heart and Stroke Foundation of Canada (Drs Pickering, Chan, and Ellis), the Medical Research Council of Canada (Drs Pickering and Chan), and the Natural Sciences and Engineering Research Council of Canada (Dr Chan). Drs Pickering and Chan are Research Scholars of the Medical Research Council of Canada.

Received September 18, 1996; accepted February 20, 1997.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

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

2. Casscells W. Endothelial and smooth muscle cell migration: critical factors in restenosis. Circulation. 1992;86:723-729.[Free Full Text]

3. Grotendorst CR, Chang T, Seppa HEJ, Kleinman HK, Martin GR. Platelet-derived growth factor is a chemoattractant for vascular smooth muscle cells. J Cell Physiol. 1982;113:261-266.[Medline] [Order article via Infotrieve]

4. Boudreau N, Turley E, Rabinovitch M. Fibronectin, hyaluronan, and a hyaluronan binding protein contribute to increased ductus arteriosus smooth muscle cell migration. Dev Biol. 1991;143:235-247.[Medline] [Order article via Infotrieve]

5. 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 intermediate attachment strength. J Cell Biol. 1993;122:729-737.[Abstract/Free Full Text]

6. 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.

7. Molossi S, Elices M, Arrhenius T, Diaz R, Coulber C, Rabinovitch M. Blockade of very late antigen-4 integrin binding to fibronectin with connecting segment-1 peptide reduces accelerated coronary arteriopathy. J Clin Invest. 1995;95:2601-2610.

8. Mitchison JT, Cramer LP. Actin-based cell motility and cell locomotion. Cell. 1996;84:371-379.[Medline] [Order article via Infotrieve]

9. Gumbiner BM. Cell adhesion: the molecular basis of tissue architecture and morphogenesis. Cell. 1996;84:345-357.[Medline] [Order article via Infotrieve]

10. 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]

11. Zempo N, Koyama N, Kenagy RD, Lea HJ, Clowes AW. Regulation of vascular smooth muscle cell migration and proliferation in vitro and in injured rat arteries by a synthetic matrix metalloproteinase inhibitor. Arterioscler Thromb Vasc Biol. 1996;16:28-32.[Abstract/Free Full Text]

12. Pauly RP, Passaniti A, Bilato C, Monticone R, Cheng L, Papadopoulos N, Gluzband YA, Smith L, Weinstein C, Lakatta EG, Crow MT. Migration of cultured vascular smooth muscle cells through a basement membrane barrier requires type IV collagenase activity and is inhibited by cellular differentiation. Circ Res. 1994;75:41-54.[Abstract/Free Full Text]

13. Gospodarowicz D. Fibroblast growth factor. Crit Rev Oncog. 1989;1:1-26.[Medline] [Order article via Infotrieve]

14. Lindner V, Lappi DA, Baird A, Majack RA, Reidy MA. Role of basic fibroblast growth factor in vascular lesion formation. Circ Res. 1991;68:106-113.[Abstract/Free Full Text]

15. Lindner V, Reidy M. Proliferation of smooth muscle cells after vascular injury is inhibited by an antibody against basic fibroblast growth factor. Proc Natl Acad Sci U S A. 1991;88:3739-3743.[Abstract/Free Full Text]

16. Jackson CL, Reidy MA. Basic fibroblast growth factor: its role in the control of smooth muscle cell migration. Am J Pathol. 1993;143:1024-1031.[Abstract]

17. Ferns GAA, Raines EW, Sprugel K, Motani AS, Reidy MA, Ross R. Inhibition of neointimal smooth muscle accumulation after angioplasty by an antibody to PDGF. Science. 1991;253:1129-1132.[Abstract/Free Full Text]

18. Bilato C, Pauly RR, Melillo G, Monticone R, Gorelick-Feldman D, Gluzband Y, Sollott SS, Ziman B, Lakatta EG, Crow MT. Intracellular signaling pathways required for rat vascular smooth muscle cell migration: interaction between basic fibroblast growth factor and platelet-derived growth factor. J Clin Invest. 1995;96:1905-1915.

19. Hynes RO. Integrins: versatility, modulation, and signalling in cell adhesion. Cell. 1992;69:11-24.[Medline] [Order article via Infotrieve]

20. Clyman RI, McDonald KA, Kramer RH. Integrin receptors on aortic smooth muscle cells mediate adhesion to fibronectin, laminin, and collagen. Circ Res. 1990;67:175-186.[Abstract/Free Full Text]

21. Clyman RI, Turner DC, Kramer RH. An ₁/ß₁-like integrin receptor on rat aortic smooth muscle cells mediates adhesion to laminin and collagen types I and IV. Arteriosclerosis. 1990;10:402-409.[Abstract/Free Full Text]

22. Belkin VM, Belkin AM, Koteliansky VE. Human smooth muscle cell VLA-1 integrin: purification, substrate specificity, localization in aorta, and expression during development. J Cell Biol. 1990;111:2159-2170.[Abstract/Free Full Text]

23. Bottger BA, Hedin U, Johansson S, Thyberg J. Integrin-type fibronectin receptors of rat arterial smooth muscle cells: isolation, partial characterization and role in cytoskeletal organization and control of differentiated properties. Differentiation. 1989;41:158-167.[Medline] [Order article via Infotrieve]

24. Lee RT, Berditchevski F, Cheng GC, Hemler ME. Integrin-mediated collagen matrix reorganization by cultured human vascular smooth muscle cells. Circ Res. 1995;76:209-214.[Abstract/Free Full Text]

25. Skinner MP, Raines EW, Ross R. Dynamic expression of 1ß1 and 2ß2 integrin receptors by human vascular smooth muscle cells: 2ß1 integrin is required for chemotaxis across type I collagen coated membranes. Am J Pathol. 1994;145:1070-1081.[Abstract]

26. Clyman RI, Mauray F, Kramer RH. ß₁ and ß₃ 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]

27. Yue T-L, McKenna PJ, Ohlstein EH, Farah-Carson MC, Butler WT, Johanson K, McDevitt P, Feuerstein GZ, Stadel JM. Osteopontin-stimulated vascular smooth muscle cell migration is mediated by ß₃ integrin. Exp Cell Res. 1994;214:459-464.[Medline] [Order article via Infotrieve]

28. Koyama N, Seki J, Vergel S, Mattsson EJ, 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]

29. Janat MF, Argraves WS, Liau G. Regulation of vascular smooth muscle cell integrin expression by transforming growth factor ß1 and by platelet-derived growth factor-BB. J Cell Physiol. 1992;151:588-595.[Medline] [Order article via Infotrieve]

30. Basson CT, Kocher O, Basson MD, Asis A, Madri JA. Differential modulation of vascular cell integrin and extracellular matrix in vitro by TGF-ß1 correlates with reciprocal effects on cell migration. J Cell Physiol. 1992;153:118-128.[Medline] [Order article via Infotrieve]

31. Klein S, Giancotti FP, Presta M, Albelda SM, Buck CA, Rifken D. Basic fibroblast growth factor modulates integrin expression in microvascular endothelial cells. Mol Biol Cell. 1993;4:973-982.[Abstract]

32. Hemler ME, Sanchez-Madrid F, Flotte TJ, Krensky AM, Burakoff SJ, Bhan AK, Springer TA, Strominger JL. Glycoproteins of 210,000 and 130,000 m.w. on activated T cells: cell distribution and antigenic relation to components on resting cells and T cell lines. J Immunol. 1984;132:3011-3018.[Abstract]

33. Hangan D, Uniyal S, Morris V, MacDonald I, von Ballestrem C, Chau T, Schmidt E, Chambers A, Groom A, Chan BMC. Integrin VLA-2 (2ß1) function in post-extravasation movement of human rhabdomyosarcoma cells in the liver. Cancer Res. 1996;56:3142-3149.[Abstract/Free Full Text]

34. Bell E, Ivarsson B, Merrill C. Production of a tissue-like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential in vitro. Proc Natl Acad Sci U S A. 1979;76:1274-1278.[Abstract/Free Full Text]

35. Ruoslahti E, Hayman EG, Pierschbacher M, Engvall E.Fibronectin: purification, immunohistochemical properties, and biological activities. Methods Enzymol. 1982;82:803-831.

36. Pickering JG, Weir L, Rosenfield K, Stetz J, Jekanowski J, Isner JM. Smooth muscle cell outgrowth from human atherosclerotic plaque: implications for the assessment of lesion biology. J Am Coll Cardiol. 1992;20:1430-1439.[Abstract]

37. Pickering JG, Bacha P, Weir L, Jekanowski J, Nichols JC, Isner JM. Prevention of smooth muscle cell outgrowth from human atherosclerotic plaque by a recombinant fusion protein specific for the epidermal growth factor receptor. J Clin Invest. 1993;91:724-729.

38. Elices MJ, Osborn L, Takada Y, Crouse C, Luhowskyj S, Hemler ME, Lobb RR. VCAM-1 on activated endothelium interacts with the leukocyte integrin VLA-4 at a site distinct from the VLA-4/fibronectin binding site. Cell. 1990;60:577-584.[Medline] [Order article via Infotrieve]

39. Chan BMC, Elices MJ, Murphy E, Hemler ME. Adhesion to vascular cell adhesion molecule 1 and fibronectin: comparison of 4ß1 (VLA-4) and 4ß7 on the human B cell line JY. J Biol Chem. 1992;267:8366-8370.[Abstract/Free Full Text]

40. Grotendorst GR, Seppa HEJ, Kleinman HK, Martin GR.Attachment of smooth muscle cells to collagen and their migration toward platelet-derived growth factor. Proc Natl Acad Sci U S A. 1981;78:3669-3672.[Abstract/Free Full Text]

41. Chan BMC, Kassner PD, Shiro JA, Byers R, Kupper TS, Hemler ME. Distinct cellular functions mediated by different VLA integrin subunit cytoplasmic domains. Cell. 1992;68:1051-1060.[Medline] [Order article via Infotrieve]

42. Sato YR, Hamanaka R, Ono J, Kuwano M, Rifken DB, Takaki R. The stimulatory effect of PDGF on vascular smooth muscle cell migration is mediated by the induction of endogenous basic FGF. Biochem Biophys Res Commun. 1991;174:1260-1266.[Medline] [Order article via Infotrieve]

43. Clowes A, Reidy M, Clowes M. Kinetics of cellular proliferation after arterial injury, I: smooth muscle growth in the absence of endothelium. Lab Invest. 1983;49:327-333.[Medline] [Order article via Infotrieve]

44. Biro S, Yu Z-X, Fu Y-M, Smale G, Sasse J, Sanchez J, Ferrans VJ, Casscells W. Expression and subcellular distribution of basic fibroblast growth factor are regulated during migration of endothelial cells. Circ Res. 1994;74:485-494.[Abstract/Free Full Text]

45. Gotwals PJ, Chi-Rosso G, Lindner V, Yang J, Ling L, Fawell SE, Koteliansky VE. The 1ß1 integrin is expressed during neointima formation in rat arteries and mediates collagen matrix reorganization. J Clin Invest. 1996;97:2469-2477.[Medline] [Order article via Infotrieve]

46. Mechtersheimer G, Barth T, Quentmeier A, Moller P. Differential expression of ß1 integrins in nonneoplastic smooth and striated muscle cells and in tumors derived from these cells. Am J Pathol. 1994;144:1172-1182.[Abstract]

47. Berdichevsky F, Gilbert C, Shearer M, Taylor-Papadimitriou J. Collagen-induced rapid morphogenesis of human mammary epithelial cells: the role of ₂ß₁ integrin. J Cell Sci. 1992;102:437-446.[Abstract/Free Full Text]

48. Keely PJ, Fong AM, Zutter MM, Santoro SA. Alteration of collagen-dependent adhesion, motility, and morphogenesis by the expression of antisense 2 integrin mRNA in mammary cells. J Cell Sci. 1995;108:595-607.[Abstract]

49. DiMilla PA, Barbee K, Lauffenburger DA. Mathematical model for the effects of adhesion and mechanics on cell migration speed. Biophys J. 1991;60:15-37.[Medline] [Order article via Infotrieve]

50. Humphries MJ, Mould AP, Yamada KM. Matrix receptors in cell migration. In: Mecham, R, ed. Receptors for Extracellular Matrix. New York, NY: Academic Press Inc; 1991.

51. Tomasek JJ, Hay ED, Fujiwara K. Collagen modulates cell shape and cytoskeleton of embryonic corneal and fibroma fibroblasts: distribution of actin, -actinin, and myosin. Dev Biol. 1982;92:107-122.[Medline] [Order article via Infotrieve]

52. Hay ED, Revel JP. Fine structure of the developing avian cornea. Monogr Dev Biol. 1969;1:1-144.[Medline] [Order article via Infotrieve]

53. Walpola PL, Gotlieb AI, Langille BL. Monocyte adhesion and changes in endothelial cell number, morphology, and F-actin distribution elicited by low shear stress in vivo. Am J Pathol. 1993;142:1392-1400.[Abstract]

54. Huttenlocher A, Ginsburg MH, Horwitz AF. Modulation of cell migration by integrin-mediated cytoskeletal linkages and ligand-binding affinity. J Cell Biol. 1996;134:1551-1562.[Abstract/Free Full Text]

55. Hotchin NA, Hall A. The assembly of integrin adhesion complexes requires both extracellular matrix and intracellular rho/rac GTPases. J Cell Biol. 1995;131:1857-1865.[Abstract/Free Full Text]

56. Pickering JG, Ford CM, Tang B, Chow LH. Coordinated effects of fibroblast growth factor-2 on expression of fibrillar collagens, matrix metalloproteinases, and tissue inhibitors of matrix metallopro-teinases by human vascular smooth muscle cells: evidence for repressed collagen production and activated degradative capacity. Arterioscler Thromb Vasc Biol. 1997;17:475-482.[Abstract/Free Full Text]

57. Langholz O, Rockel D, Mauch C, Kozlowska E, Bank I, Krieg T. Collagen and collagenase gene expression in three-dimensional collagen lattices are differentially regulated by 1ß1 and 2ß1 integrins. J Cell Biol. 1995;131:1903-1915.[Abstract/Free Full Text]

58. Ridley AJ, Hall A. The small GTP-binding protein rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell. 1992;70:389-399.[Medline] [Order article via Infotrieve]

59. Nobes CD, Hall A. Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell. 1995;81:53-62.[Medline] [Order article via Infotrieve]

60. Rankin S, Rozengurt E. PDGF modulation of focal adhesion kinase p125^FAK and paxillin tyrosine phosphorylation in Swiss 3T3 cells: bell-shaped dose response and cross-talk with bombesin. J Biol Chem. 1994;269:704-710.[Abstract/Free Full Text]

61. Choi ET, Engel L, Callow AD, Sun S, Trachtenberg J, Santoro S, Ryan US. Inhibition of neointimal hyperplasia by blocking _vß₃ integrin with a small peptide antagonist GpenGRGDSPCA. J Vasc Surg. 1994;19:125-134.[Medline] [Order article via Infotrieve]

62. Matsuno H, Stassen JM, Vermylen J, Deckmyn H. Inhibition of integrin function by a cyclic RGD-containing peptide prevents neointimal formation. Circulation. 1994;90:2203-2206.[Abstract/Free Full Text]

This article has been cited by other articles:

				M. J. Frontini, C. O'Neil, C. Sawyez, B. M.C. Chan, M. W. Huff, and J. G. Pickering Lipid Incorporation Inhibits Src-Dependent Assembly of Fibronectin and Type I Collagen by Vascular Smooth Muscle Cells Circ. Res., April 10, 2009; 104(7): 832 - 841. [Abstract] [Full Text] [PDF]

				K. Schroder, I. Helmcke, K. Palfi, K.-H. Krause, R. Busse, and R. P. Brandes Nox1 Mediates Basic Fibroblast Growth Factor-Induced Migration of Vascular Smooth Muscle Cells Arterioscler Thromb Vasc Biol, August 1, 2007; 27(8): 1736 - 1743. [Abstract] [Full Text] [PDF]

				S. Filippov, G. C. Koenig, T.-H. Chun, K. B. Hotary, I. Ota, T. H. Bugge, J. D. Roberts, W. P. Fay, H. Birkedal-Hansen, K. Holmbeck, et al. MT1-matrix metalloproteinase directs arterial wall invasion and neointima formation by vascular smooth muscle cells J. Exp. Med., September 6, 2005; 202(5): 663 - 671. [Abstract] [Full Text] [PDF]

				C. H. O'Neil, M. B. Boffa, M. A. Hancock, J. G. Pickering, and M. L. Koschinsky Stimulation of Vascular Smooth Muscle Cell Proliferation and Migration by Apolipoprotein(a) Is Dependent on Inhibition of Transforming Growth Factor-{beta} Activation and on the Presence of Kringle IV Type 9 J. Biol. Chem., December 31, 2004; 279(53): 55187 - 55195. [Abstract] [Full Text] [PDF]

				E. Fera, C. O'Neil, W. Lee, S. Li, and J. G. Pickering Fibroblast Growth Factor-2 and Remodeled Type I Collagen Control Membrane Protrusion in Human Vascular Smooth Muscle Cells: BIPHASIC ACTIVATION OF Rac1 J. Biol. Chem., August 20, 2004; 279(34): 35573 - 35582. [Abstract] [Full Text] [PDF]

				S. Li, C. Van Den Diepstraten, S. J. D'Souza, B. M. C. Chan, and J. G. Pickering Vascular Smooth Muscle Cells Orchestrate the Assembly of Type I Collagen via {alpha}2{beta}1 Integrin, RhoA, and Fibronectin Polymerization Am. J. Pathol., September 1, 2003; 163(3): 1045 - 1056. [Abstract] [Full Text] [PDF]

				E. P. Moiseeva Adhesion receptors of vascular smooth muscle cells and their functions Cardiovasc Res, December 1, 2001; 52(3): 372 - 386. [Abstract] [Full Text] [PDF]

				A. Watanabe, M. Kurabayashi, M. Arai, K. Sekiguchi, and R. Nagai Combined effect of retinoic acid and basic FGF on PAI-1 gene expression in vascular smooth muscle cells Cardiovasc Res, July 1, 2001; 51(1): 151 - 159. [Abstract] [Full Text] [PDF]

				R. M. Korah, V. Sysounthone, Y. Golowa, and R. Wieder Basic Fibroblast Growth Factor Confers a Less Malignant Phenotype in MDA-MB-231 Human Breast Cancer Cells Cancer Res., February 1, 2000; 60(3): 733 - 740. [Abstract] [Full Text]

				J. G. Pickering, L. H. Chow, S. Li, K. A. Rogers, E. F. Rocnik, R. Zhong, and B. M. C. Chan {alpha}5{beta}1 Integrin Expression and Luminal Edge Fibronectin Matrix Assembly by Smooth Muscle Cells after Arterial Injury Am. J. Pathol., February 1, 2000; 156(2): 453 - 465. [Abstract] [Full Text] [PDF]

				A Facchiano, F De Marchis, E Turchetti, F Facchiano, M Guglielmi, A Denaro, R Palumbo, M Scoccianti, and M. Capogrossi The chemotactic and mitogenic effects of platelet-derived growth factor-BB on rat aorta smooth muscle cells are inhibited by basic fibroblast growth factor J. Cell Sci., January 8, 2000; 113(16): 2855 - 2863. [Abstract] [PDF]

				E Stringa, V Knauper, G Murphy, and J Gavrilovic Collagen degradation and platelet-derived growth factor stimulate the migration of vascular smooth muscle cells J. Cell Sci., January 6, 2000; 113(11): 2055 - 2064. [Abstract] [PDF]

				A Scherberich, M Campos-Toimil, P Ronde, K Takeda, and A Beretz Migration of human vascular smooth muscle cells involves serum-dependent repeated cytosolic calcium transients J. Cell Sci., January 2, 2000; 113(4): 653 - 662. [Abstract] [PDF]

				X.-Q. Wang, F. P. Lindberg, and W. A. Frazier Integrin-Associated Protein Stimulates {alpha}2{beta}1-Dependent Chemotaxis via GI-Mediated Inhibition of Adenylate Cyclase and Extracellular-Regulated Kinases J. Cell Biol., October 18, 1999; 147(2): 389 - 400. [Abstract] [Full Text] [PDF]

				S. Li, S. Sims, Y. Jiao, L. H. Chow, and J. G. Pickering Evidence From a Novel Human Cell Clone That Adult Vascular Smooth Muscle Cells Can Convert Reversibly Between Noncontractile and Contractile Phenotypes Circ. Res., August 20, 1999; 85(4): 338 - 348. [Abstract] [Full Text] [PDF]

				C. Brown, X. Pan, and A. Hassid Nitric Oxide and C-Type Atrial Natriuretic Peptide Stimulate Primary Aortic Smooth Muscle Cell Migration via a cGMP-Dependent Mechanism : Relationship to Microfilament Dissociation and Altered Cell Morphology Circ. Res., April 2, 1999; 84(6): 655 - 667. [Abstract] [Full Text] [PDF]

				D. Palmer, K. Tsoi, and D. H. Maurice Synergistic Inhibition of Vascular Smooth Muscle Cell Migration by Phosphodiesterase 3 and Phosphodiesterase 4 Inhibitors Circ. Res., May 4, 1998; 82(8): 852 - 861. [Abstract] [Full Text] [PDF]

				S. Li, L. H. Chow, and J. G. Pickering Cell Surface-bound Collagenase-1 and Focal Substrate Degradation Stimulate the Rear Release of Motile Vascular Smooth Muscle Cells J. Biol. Chem., November 3, 2000; 275(45): 35384 - 35392. [Abstract] [Full Text] [PDF]

				S. Li, Y.-S. Fan, L. H. Chow, C. Van Den Diepstraten, E. van der Veer, S. M. Sims, and J. G. Pickering Innate Diversity of Adult Human Arterial Smooth Muscle Cells: Cloning of Distinct Subtypes From the Internal Thoracic Artery Circ. Res., September 14, 2001; 89(6): 517 - 525. [Abstract] [Full Text] [PDF]

This Article Abstract 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 Pickering, J. G. Articles by Chan, B. M. C. Search for Related Content PubMed PubMed Citation Articles by Pickering, J. G. Articles by Chan, B. M. C.