American Journal of Respiratory Cell and Molecular Biology. Vol. 28, pp. 436-442, 2003
© 2003 American Thoracic Society
DOI: 10.1165/rcmb.4754
Mechanical Stress Increases RhoA Activation in Airway Smooth Muscle Cells
Paul G. Smith,
Chaity Roy,
Ying Ning Zhang and
Subhenu Chauduri
Department of Pediatrics, Case Western Reserve University, Cleveland, Ohio
Address correspondence to: Paul G. Smith, D.O., 11000 Euclid Avenue, Rainbow Babies and Children's Hospital, Department of Pediatrics, Case Western Reserve University, Cleveland, OH 44106. E-mail: pgs3{at}po.cwru.edu
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Abstract
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Cultured airway smooth muscle cells subjected to cyclic strain respond with increased cytoskeletal organization and contractility resembling effects described with RhoA activation. To test the hypothesis that strain increases cell cytoskeletal organization through RhoA, cells were subjected to strain in the presence of known activators or inhibitors of RhoA. Ten percent cyclic deformational strain (serum-free conditions) increased F-actin staining (152% over control), and this effect was enhanced by serum or lysophosphatidic acid (180%), but decreased (68%) with Clostridium botulinum toxin inhibition of RhoA or with the Rho kinase inhibitor Y27632 (67%). When cells expressing the dominant negative N17–RhoA isoform were subjected to strain, F-actin staining was disorganized and cells failed to elongate or migrate relative to strain direction. When cells expressing a green fluorescent protein (GFP)-RhoA fusion protein were subjected to strain, GFP showed up to 25% greater cell membrane staining than control cells. Finally, strain caused a 4-fold increase in RhoA activation (Rhotekin binding assay), and a 3-fold increase myosin phosphatase phosphorylation that was inhibited by Y27632. We conclude that mechanical stress activates RhoA, an event that may increase airway smooth muscle contractility.
Abbreviations: airway smooth muscle, ASM • fetal bovine serum, FBS • fluorescein isothiocyanate, FITC • green fluorescent protein, GFP • lysophosphatidic acid, LPA • myosin-binding subunit, MBS • myosin light chain kinase, MLCK • phosphate-buffered saline, PBS • reverse transcription polymerase chain reaction, RT-PCR • sodium dodecyl sulfate, SDS
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Introduction
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When tissue containing smooth muscle is subjected to chronic increases in mechanical stress, smooth muscle content and contractility is increased (1–3). Such changes are maladaptive as seen in chronic obstructive lung diseases, pulmonary hypertension, and hypertensive vasculopathy. To study the consequences of mechanical stress on airway smooth muscle (ASM) independent of other in vivo factors, we have developed an in vitro system exposing cultured ASM cells to cyclic deformational strain approximating levels seen in vivo. With this system, we have shown that cyclic deformational strain increases cytoskeletal organization, expression of smooth muscle-specific contractile proteins, and the contractility of cultured ASM cells (3–5). These studies suggest that mechanical stress is a major determinant of smooth muscle phenotype in some diseases.
The molecular pathways transducing mechanical signals into cellular responses are therefore of interest to understand factors contributing to phenotypic modulation and develop means to treat maladaptive states. One group of signaling molecules influenced by signaling from the cell membrane is the Ras family of small GTPases. RhoA, a member of this family, is of particular interest because its activation mediates a number of cell processes that are identical to stress-induced changes we have described in our system, and because RhoA activation influences smooth muscle contractility. RhoA is activated when it cycles from the GDP- to the GTP-bound state under the influence of guanidine exchange factors. RhoA activation could increase smooth muscle contractility by promoting contractile protein production (6), actin filament assembly (7), myosin light chain phosphorylation (8), inhibition of myosin phosphatase (9), and calcium sensitivity of force production (10). Similar changes are seen when mechanical stress is placed on cultured ASM cells: increased contractile protein production (5), increased stress fibers and focal adhesion formation (11), myosin phosphatase inhibition (12), and increased calcium sensitivity of force production (4). Therefore, we studied whether strain activates RhoA in ASM cells by comparing levels of RhoA activation (RhoA-GTP) and by detecting RhoA localization to the membrane during strain. Because some of the effects of RhoA enhancement of smooth muscle contractility are mediated through Rho kinase, a downstream effector of RhoA (10), we also studied whether inhibition of Rho kinase altered the morphologic responses of ASM cells to strain.
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Materials and Methods
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Cell Culture and Mechanical Stress Protocol
Canine trachealis muscle was harvested and digested in collagenase and elastase with soy trypsin inhibitor as previously described (3). Freshly dissociated cells were seeded into flasks at a density of 5 x 104 cells/cm2 in Ham's F-12/Dulbecco's modified Eagle's medium with 10% fetal bovine serum (FBS), penicillin, streptomycin, and amphotericin and maintained at 37°C in 5% CO2 in air in a humidified incubator. Cell passages 2–4 were used for all experiments. Cells were passaged to collagen type I–coated silastic membranes in six well plates (Bioflex plate, Cat. # 3001C; Flexcell Inc., McKeesport, PA) at a density of 5 x 105 cells/10 cm2 well. Cells were subjected to mechanical stress by positioning the plates over a manifold connected to a vacuum source. The vacuum was programmed by computer software (Flexcell Inc.) to cause a 10% increase in surface area of the membranes for 2 s followed by 2 s of relaxation (cyclic deformational strain). The duration of this strain protocol varied as described for individual experiments.
Reagents Modifying RhoA Activation during Strain
In studies where the various agents were used to alter RhoA activation, cells were grown to  50% confluence and the media changed to serum-free media for 24 h before experiments. Cells were then exposed to the following drugs before and during strain using the protocol described. To activate RhoA, cells were treated either with lysophosphatidic acid (LPA) 10 µM (Sigma, St. Louis, MO), or serum (10% FBS). To inhibit RhoA, cells were pretreated with Clostridium botulinum C3 exotoxin 6 µg/ml overnight (List Biological Inc., Campbell, CA). To study the downstream effector Rho kinase, the inhibitor Y-27632 (1 µM) (kindly provided by Welfide Corporation, Osaka, Japan) was studied in a similar fashion. Cells were then subjected to strain while drug concentrations were maintained (media changed daily) to assess changes in cell morphology. Images of cells were acquired and analyzed as described below.
Vectors and Methods for Transient Transfection of Smooth Muscle Cells
The bacterial expression vector pCEV co-expressing the AU5 reporter gene with either wild-type RhoA or the dominant-negative isoform RhoA-N17 were propagated in Escherichia coli (kind gifts of Silvio Gutkind, Ph.D., Scripts Institute, La Jolla, CA). ASM cells were plated as described and allowed to reach 50% confluent growth of the monolayer. Media was changed to serum-free, antibiotic-free media and transfection performed with the various expression vectors using Lipofectamine (Lipofectamine Plus; Gibco BRL, Gaithersburg, MD). Media containing 10% FBS was then re-added to wells and transfection performed overnight. For control, another set of cells was transfected with a pCEV plasmid containing only the AU5 construct. The transfection efficiency was 15% with the pCEV/AU5 and pCEV/AU5/Val14-RhoA vectors and between 10 and 16% using the pCEV/AU5/N17-RhoA vector determined by indirect immunocytochemical staining with rabbit antibody against AU5 (Gibco BRL) and fluorescein isothiocyanate (FITC)-labeled goat anti-rabbit antibodies (Sigma). After determination of expression in sample wells, other wells were subjected to strain for 48 h before fixing and staining for AU5 as above in combination with rhodamine-conjugated phalloidin (Sigma) for F-actin.
Previous studies have determined that RhoA localizes to smooth muscle membranes upon activation (13). To study whether mechanical strain causes RhoA localization to the membrane, a plasmid expressing a green fluorescent protein (GFP)-RhoA fusion product was constructed. The coding sequence of RhoA was amplified by reverse transcription polymerase chain reaction (RT-PCR) of RNA from a human cell line. RT-PCR was performed by random priming with hexamers and PCR using primers 5'Rho/XhoI 5'-tggactcgagttcgttgcctgagc-3' and Bam Rho3' 5'-caaggatcccacaagacaaggcaac-3', which amplifies residues 127 to 733 of RhoA (Genbank accession #L25080). To facilitate cloning into the GFP fusion vector pEGFP-N1 (Clontech, Palo Alto, CA) the primer for the 5' end contains an Xho1 site and the 3' primer a BamHI site. Nucleotide 773 is changed from A to G, creating a tryptophan codon in place of the stop, allowing the coding sequence of RhoA to be fused in frame with GFP. The 611-bp amplicon was digested with BamHI and XhoI and cloned into the corresponding sites of pEGFP-N1, and the structure of the clones was verified by restriction digestion. ASM cells were transfected as above when 50% confluent with 2 µg DNA per 10 cm2 well in serum-free media. GFP expression was determined by direct visualization of the living cells in situ under fluorescent light before initiating experiments. Experiments were conducted by subjecting the cells to strain by the above protocol for short periods of time from 5 min to 2 h. Cells were fixed on the membranes and images of GFP-positive cells were captured and analyzed as described.
Cell Imaging and Image Analysis
To determine the influence of the above conditions on strain-induced cell morphology and F-actin staining, sections of the culture membranes with cells intact were fixed with 4% paraformaldehyde/0.1% glutaraldehyde in phosphate-buffered saline (PBS) for 10 min and rinsed with PBS, followed by 0.1% sodium borohydrate for 5 min. Cells were permeabilized with 0.5% Triton-X 100 in PBS for 5 min. FITC-tagged DNase (0.3 µM; Molecular Probes, Eugene, OR) was used to label globular actin, and rhodamine (RITC)-phalloidin (0.165 µM; Molecular Probes) was used to label filamentous actin. The membranes were mounted on glass slides with Crystal/Gel mount (Biomed, Hayward, CA.) and cells viewed on an inverted epifluorescence microscope (Axiovert 35; Zeisse; Thornwood, NY). For FITC visualization, fluorescence was excited at 495 nm and emission filtered at 540 nm. For rhodamine, fluorescence was excited at 535 nm and emission filtered at 560 nm. To measure actin staining, cells were visualized through a x40 objective. For studies examining localization of green fluorescent protein, fluorescence was excited at 475 nm, and light reflected by a 505 nm dichroic mirror and emitted light filtered through a 535 nm filter. Images of GFP-labeled cells were visualized through a x63 objective. Images were captured by a cooled CCD camera (Model CH 250; Photometric, Ltd, Tucson, AZ) to a host computer and analyzed with IP Lab software (Scanalytics Inc., Fairfax, VA). Exposure times and aperture settings were held constant from field to field. Images from multiple conditions within one experiment were all captured on the same day under identical conditions to avoid differences in staining intensity from photobleaching or minor variations in reagents. Computer software determined organization of actin into filaments by calculating the ratios of the intensity of phalloidin to DNase staining for individual cells.
Several cell types subjected to cyclic strain migrate so that their long axis are perpendicular to the direction of strain (14). The apparatus used in the present study creates a vacuum beneath circular wells containing the silastic membranes on which cells are grown. The major direction of deformation is from the edge of the circular well to the center, thus alignment of the cells' long axes is parallel to the well circumference so that the cells appear to form a ring around the edge of the well (11). To quantify the cells' tendency to realign perpendicular to the direction of strain, the outer edge of the well was placed horizontally in the microscope field of view. Computer software identified the long axis of the cell and measured the angle between this axis and the edge of the well. Cells with their long axis parallel to the edge of the well were assigned an angle value of 0°, and cells with their long axis oriented toward the center of the well were assigned a value of 90° (11). Accordingly, a population of cells with random alignment would be expected to have a mean angle of 45°.
When RhoA is in the active state, it migrates to the cell membrane (13). To study whether strain causes RhoA localization to the cell membrane, cells were transfected with a plasmid expressing RhoA-GFP, subjected to strain, and the ratio of staining intensity between the membrane and the cytoplasm was measured. A line was drawn manually with the computer cursor along the cell membrane, including at least 25% of the circumference of the cell. A second line of equal length was drawn inside the cell. When drawing the lines, artifacts from nuclear staining were avoided by staying near the periphery of the cell. The computer software summed the total intensity of the line pixels and calculated the mean intensity of staining per pixel within the lines. These mean intensities were used to calculate a ratio of membrane to cytoplasmic staining.
Rhotekin Binding Assay for RhoA-GTP
RhoA is bound to GDP in its inactive state, and is activated when GDP is exchanged for GTP. To determine if strain increased RhoA activation, i.e., RhoA-GTP, we used the Rhotekin binding domain affinity precipitation assay for RhoA-GTP described by Ren and coworkers (15) and O'Connor and colleagues (16). ASM cells were grown to confluence and then changed to serum-free media for 24 h. After application of cyclic strain for periods of time from 5 min to 2 h, cells were lysed in RIPA buffer (50 mM Tris, pH 7.2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 500 mM NaCl, 10 mM MgCl, 10 µg/ml each of leupeptin and aprotinin, and 1 mM PMSF) on ice, and the lysates centrifuged at 13,000 x g and 4°C for 3 min. One aliquot of lysate was analyzed for total protein content and another aliquot was analyzed by Western blotting (normalized for total protein), to compare total RhoA content between samples. The remainders of the cell lysates (500 µl; 0.5–1.0 µg/µl protein) were added to 80 µl of the glutathione bead slurry (Amersham Pharmacia Biotech, Piscataway, NJ; Rho-binding domain of Rhotekin kindly provided by Martin Schwartz, Scripps Research Institute, La Jolla, CA, or purchased from Upstate Biotechnology, Lake Placid, NY) at 4°C for 45 min. The beads were washed thoroughly with buffer and retained proteins were separated by SDS-polyacrylamide gel electrophoresis (PAGE) in 12% acrylamide gels (100 V for 2 h) and then transferred (100 V for 1 h) to Immobilon-P membranes (Millipore, Bedford, MA). RhoA was detected with anti-RhoA primary antibodies (Santa Cruz Biotechnology, Inc. Santa Cruz, CA) and horseradish peroxidase–conjugated goat anti-mouse IgG (Sigma). Bands were detected with enhanced chemiluminescence (ECL-Super Signal; Pierce, Rockford, IL) and images on radiographic film were quantified using two-dimensional laser densitometry (USB Sciscan 5000; US Biochemical, Cleveland, OH).
Strain-Induced Inhibition of Myosin Phosphatase
We have previously reported strain-induced decreases in the activity of myosin phosphatase (12), and increases in calcium sensitivity of force production of cultured cells (4). We wished to determine if these effects could also be related to RhoA activation. It has been shown that myosin phosphatase inhibition can result from phosphorylation of the myosin-binding subunit (MBS) of myosin phosphatase. This occurs through activation of Rho kinase, which in turn requires RhoA activation (9). We wished to determine if strain increased the phosphorylation of the MBS of myosin phosphatase, and if this required Rho kinase activation. Cells were grown to confluence and then placed in serum-free media overnight. Cells were then subjected to strain for 1 h and compared with cells subjected to strain in the presence of the Rho kinase inhibitor Y-27632 and control cells (no strain). Cells were lysed in Laemmli sample buffer, and proteins separated by SDS-PAGE on 4–20% minigels (Biorad Laboratories, Hercules, CA) at 100 V for 1 h. After electrophoretic transfer to nitrocellulose paper (125 V for 50 min) in Tris-glycine buffer, the membranes were blocked with 5% milk. Bands were detected using antibody that recognizes phosphorylation at the Ser-854 site of MBS and does not recognize nonphosphorylated MBS (a kind gift from Kojo Kaibuchi, Nara Institute, Nara, Japan) (9). The blots were developed using ECL as described.
Statistics
Comparison of results between different experimental conditions was performed by one-way ANOVA with Bonferroni's correction. Significance was set at P < 0.05. Data are expressed as mean ± SD.
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Results
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F-actin Staining and Cell Alignment Correlate with RhoA Activation
To determine if strain-induced increases in cell organization involve RhoA activation, cells were subjected to strain in the presence of agents known to increase (serum, LPA) or decrease (C3) RhoA activation or inhibit Rho kinase activity (Y-27632). In serum-free conditions, strain caused significant increases F-actin:G-actin staining, and increased cell alignment perpendicular to the direction of strain compared with identical cells not subjected to strain (Figures 1 and 2). F-actin staining and cell realignment was enhanced when cells were subjected to strain in the presence of LPA or serum. Conversely, use of either C3 to inhibit RhoA or the Rho Kinase inhibitor Y-27632 caused marked loss of F-actin staining, and cells more randomly oriented relative to strain direction (Figures 1 and 2).
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Figure 1. Strain-induced changes in cell organization affected by RhoA activation. (A) Photomicrographs of ASM cells subjected to strain (48 h; serum-free conditions) and stained with rhodamine phalloidin demonstrate strain-induced increases in F-actin staining. Compared with no-strain, serum-free conditions used as control (B), mechanical strain also caused cells to orient perpendicular to direction of strain (arrow) in serum free conditions. Addition of serum (C) or LPA (10 µM) (D) to strain cells increased F-actin staining, whereas pretreatment of cells with C3 (E) or the Rho kinase inhibitor Y27632 (F) inhibited F-actin staining and cell alignment relative to the direction of strain.
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Figure 2. Quantitation of F-actin staining and cell alignment from recorded images. Ratios of staining intensity for F-actin (phalloidin) and G-actin (DNase) showed increased F-actin staining (A) and cell alignment (B) with strain compared with control (no-strain) in serum-free (-) conditions. Serum (+) or LPA increased F-actin staining in both control or strain cells and enhanced strain effects. C3 and the Rho kinase inhibitor Y27632 reduced strain-induced F-actin staining (C) and cell realignment (D). (n = 200–300 cells/per condition in each of three experiments; *P < 0.05 = compared with control, ANOVA.)
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Expression of RhoA Isoforms Alter Cell Response to Strain
To further determine the role of RhoA in strain-induced cell morphologic changes, cells were transfected with the vectors expressing either wild-type RhoA or the dominant-negative isoform N17-RhoA, co-expressed with the marker protein AU5. After expression was noted in sample wells, other wells expressing these vectors were subjected to strain for 48 h, fixed, and dual stained for F-actin and AU5 as described. In the cells that expressed wild-type RhoA, strain-induced elongation and F-actin staining was similar to normal cells not expressing this protein (Figure 3A). However, expression of N17-RhoA caused cells to lose F-actin staining and prevented cell re-alignment and elongation relative to the direction of strain (Figure 3B).
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Figure 3. RhoA isoform expression alters F-actin staining. Cells co-expressing the marker protein AU5 with either N19-RhoA (dominant-negative isoform) or wild-type RhoA were subjected to strain for 48 h and then stained for AU5 (green arrows) or F-actin (red). In cells that expressed wild-type RhoA, strain-induced elongation and F-actin staining was similar to normal cells not expressing this protein (A). However, expression of N17-RhoA caused disorganization of stress fibers and prevented cell realignment and elongation relative to the direction of strain (B).
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Strain Increases RhoA Localization to Membranes
As an index of RhoA activation, cells were subjected to strain after expression of the RhoA-GFP fusion protein to determine distribution of RhoA in the cell. Increases in membrane localization of RhoA-GFP were noted with strain within 5 min, and were maintained with continuous strain of up to 24 h (Figure 4). This staining intensity was not uniform around the entire membrane circumference, but rather seemed to concentrate at the ends of the cells at points where maximal strain was sensed by focal adhesions. Furthermore, there was often linear distribution of the stain that correlated to the distribution of F-actin staining.
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Figure 4. RhoA localizes to the cell membrane during mechanical strain. To determine the effects of strain on RhoA localization, a RhoA-GFP fusion protein was expressed in cultured ASM cells. Cells not subjected to strain showed diffuse staining of the cytoplasm (A). Cells subjected to strain demonstrated increased GFP staining at the cell membrane (B; arrows). One line was traced along the cell membrane (arrows) and one in the cell body (arrowhead) (C). Software computed the mean pixel intensity for the two lines (D). Quantification of membrane to cytoplasm staining for control and strain cells demonstrates early and sustained localization in strain cells (n = 20 cells per time point from three experiments; *P < 0.05 = compared with control, ANOVA).
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Strain Increases RhoA Activation
To study activation of RhoA, relative proportions of RhoA-GTP to total RhoA, were compared between control cells (no strain) and strain cells with the Rhotekin binding domain affinity assay. Strain increased the cell content of RhoA-GTP relative to total Rho within 5 min. These increases were maximal at 1–2 h, and were sustained through 24 h though at lower levels than the 1-h samples (Figure 5).
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Figure 5. Strain increases cell content of RhoA-GTP. (A) ASM cells were subjected to cyclic strain for various periods and content of RhoA-GTP relative to total RhoA was assessed by RBD affinity assay. In cell subjected to strain, RhoA-GTP increased rapidly compared with control. RhoA-GTP levels continued to be increased relative control during 24 h of strain. (B) Densitometry data from ECL preparations expressed in integrated optical density units (I.O.D.) RhoA-GTP/total RhoA as a percent of control for each experiment. (P < 0.05, ANOVA, n = 3.)
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Strain Increases MBS Phosphorylation
To demonstrate the potential for RhoA-induced myosin phosphatase inhibition with strain, we examined whether there was increased phosphorylation of the MBS of myosin phosphatase, and if such increases were dependent on the RhoA effector Rho kinase. Western blots demonstrated increased phosphorylation of MBS occurring within 30 min of cyclic deformational strain (Figure 6). Preincubation of the cells subjected to strain with the Rho kinase inhibitor Y27632 prevented strain-induced phosphorylation of MBS.
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Figure 6. Strain increases myosin phosphatase phosphorylation. (A) Strain cells (S) were compared with control cells (C) by Western blotting to determine if strain-induced RhoA activation was accompanied by myosin phosphatase MBS phosphorylation. To demonstrate involvement of Rho kinase downstream of RhoA, separate cells were treated with Y27632 while undergoing strain. Strain increased MBS phosphorylation (MBS-P) at 30 min, an effect that was inhibited by Y27632. (B) Densitometry data from ECL preparations expressed in I.O.D. units of phosphorylated to total MBS (*P < 0.05, n = 4).
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Discussion
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We have reported strain-induced increases in the cytoskeletal organization, contractile protein production, and contractility in cultured ASM cells (4, 12, 17). Because increased smooth muscle contractility is seen in tissue chronically exposed to abnormal stress, the cultured cells may help determine the processes underlying the hypercontractile state. This study extends our previous work by demonstrating a critical role for RhoA in the transduction of mechanical signals into cellular responses that might increase contractile potential. RhoA activation in this system occurred rapidly and was independent of humoral factors in serum such as LPA. The increase in activation continued, whereas the mechanical stress was sustained but diminished with prolonged strain. In this cell culture system, there was clear enhancement of the morphologic responses to mechanical strain when RhoA activity was increased either pharmacologically or by expression vectors. In contrast, there was clear inhibition of strain-induced effects with pharmacologic inhibition of RhoA. Involvement of the downstream effector Rho kinase was demonstrated with inhibition of strain effects by the specific inhibitor Y27632 and increased phosphorylation of the MBS of myosin phosphatase, a known consequence of Rho kinase activation. Finally, the use of expression vectors demonstrated dependency of RhoA on the cell response to strain in that expression of the dominant-negative isoform prevented this response.
The relevance of these studies is in demonstrating cellular mechanisms whereby mechanical stress might contribute to increased smooth muscle contractility in certain diseases. Several parameters of contractility were increased in the in vitro system, such as force production, calcium sensitivity, and shortening velocity (4, 17). The cellular processes involved could be accounted for by increased production of contractile proteins, increased activation or decreased inactivation of myosin light chain and cytoskeletal organization. Accordingly, RhoA activation has been shown to play a role in each of these processes, making RhoA particularly relevant to the function of smooth muscle (6, 8–10). Strain-induced RhoA activation could result in increased contractility through increased production of contractile proteins and force-generating units (6). Calcium sensitization appeared to be in part secondary to decreases in myosin phosphatase activity mediated by the downstream effector of RhoA, Rho kinase (9). Inhibited phosphatase activity would also result in higher levels of light chain activation, increased velocity of shortening and force generation. In fact, we have previously shown a decrease in myosin phosphatase activity in cells subjected to mechanical strain (12).
We also reported strain-induced increases in stress fiber formation, as have others (11, 18, 19). Such cytoskeletal organization is a well-described consequence of RhoA activation, and is used as an index of RhoA activation (7). Although, to our knowledge, increased cytoskeletal element organization has not been reported from pathologic specimens of smooth muscle, increased organization of cytoskeletal elements has been proposed to account for increased reactivity of ASM and impairment of relaxation in obstructive lung diseases (20). It is reasonable to speculate that RhoA-induced increases in cytoskeletal organization would increase the contractile potential of smooth muscle cells. By orienting contractile forces along a uniform vector, shortening could become more efficient. An increase in the actin scaffolding would increase the number of sites for myosin cross-bridge cycling, and contraction would be enhanced (21). In fact, we reported increased force generation in the cultured cells that could not be entirely explained by increased activation potential or enzyme activity (4). This would indicate that RhoA activation could enhance cell contractility though cytoskeletal formation and organization.
The time course of RhoA activation in this study is of interest, showing rapid increases that are sustained with continued strain. There appears to be a decline in RhoA activation with strain of long duration, though levels do not return to baseline. This was evident both in the studies of localization of RhoA to the membrane and the Rhotekin affinity assay. Because RhoA plays a role in cell migration, focal adhesion formation, and stress fiber formation, we speculate that after cells reorient perpendicular to the direction of strain, elements have rearranged to minimize internal stress and less membrane tension is generated. Stretching the width rather than the length of the cells would lessen cell distortion (19). We have reported increased phosphorylation of focal adhesion related proteins such as paxillin and focal adhesion kinase with mechanical strain in this system that demonstrate a similar time course and thus support this speculation (11). Tyrosine phosphorylation of these proteins in turn may participate in cell organizational changes and smooth muscle contractility (22, 23). In fact, RhoA participates in increasing paxillin and FAK tyrosine phosphorylation (24), and through this pathway increases cytoskeletal organization, similarly increasing contractile potential (22). An alternative explanation for this time course may be that focal adhesions are undergoing more rapid turnover at the 24-h time point than the early time points, and RhoA activation is suppressed during integrin engagement (25)
This study demonstrates mechanical stress activation of RhoA in cultured ASM cells that controls the cell reorganization responses to stress. Furthermore, an enhanced organization was seen when RhoA activation was increased pharmacologically. Conversely, inactivation of RhoA inhibited strain-induced cell elongation and organization of cytoskeletal elements. The response apparently involved the downstream effector of RhoA, Rho kinase as use of a specific inhibitor prevented strain-induced responses. The events upstream of RhoA activation are not explored in these studies, but could elucidate means of modifying cellular responses to abnormal mechanical stress. Potential mechanisms include mechanosensitive ion channels, integrin mediated-signaling, or cross-talk between G protein–coupled receptors and receptor tyrosine kinases. We also have not directly correlated strain-induced activation of RhoA with contractile properties of the individual cell, but these data would suggest that changes in cell compliance and force generation should be consequences of cytoskeletal reorganization.
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Acknowledgments
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The authors thank Mitch Drumm, Ph.D. and Frank Mularo for construction of GFP-RhoA expression vector, Dr. Xu Ren and Martin Schwartz for provision of RBD constructs and advice, and Mitsou Ikebe, Ph.D. for Rho expression vectors. They also thank Frank Brozovich, M.D. and Kathy O'Connor, Ph.D. for constructive comments. This work was supported by National Heart, Lung and Blood Institute grant HL-03409-2 (P.G.S.).
Received in original form October 29, 2001
Received in final form September 27, 2002
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Abstract
Introduction