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Canadian Institutes of Health Research Group in Matrix Dynamics, Faculty of Dentistry, University of Toronto, Toronto, Ontario, Canada M5S 3E8
Submitted 25 April 2003 ; accepted in final form 25 June 2003
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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-smooth muscle actin (SMA) expression is the classical marker for myofibroblast differentiation, we examined force-induced SMA expression and regulation by specific MAPK pathways. Rat cardiac fibroblasts were separated from myocytes and smooth muscle cells, cultured, and phenotyped by using the presence of SMA, vimentin, and ED-A fibronectin and the absence of desmin as myofibroblast markers. Static tensile forces (0.65 pN/µm2) were applied to fibroblasts via collagen-coated magnetite beads. In neonatal cardiac fibroblasts cultured for 1 day, immunostaining and Western and Northern blotting showed very low basal levels of SMA. After the application of force, there were 1.5- to 2-fold increases of SMA protein and mRNA within 4 h. Force-induced SMA expression was dependent on ERK phosphorylation and on intact actin filaments. In contrast to cells cultured for 1 day, cells grown for 3 days on rigid substrates showed prominent stress fibers and high basal levels of SMA, which were reduced by 32% within 4 h after force application. ERK was not activated by force, but p38 phosphorylation was required for force-induced inhibition of SMA expression. These results indicate that mechanical force-induced regulation of SMA content is dependent on myofibroblast differentiation and by selective activation of MAPKs.
force application; integrins; cell differentiation; extracellular signal-regulated kinase; p38
The cardiac fibroblast is the most abundant cell type in the myocardium (14), and, in many instances, fibroblastic remodeling of the extracellular matrix is sensitive to changes of the mechanical and chemical properties of the tissue environment (5). Cardiac fibroblasts respond to mechanical loading by a switch to a myofibroblastic phenotype in which cells express -smooth muscle actin (SMA) (22), an actin isoform strongly associated with increased contractile force (16). SMA expression is increased in myofibroblasts found in hypertrophic and fibrotic hearts and at sites of myocardial infarction (7, 20). Furthermore, SMA is strongly upregulated in cells expressing angiotensin II receptors or after the development of hypertension (37). Currently, little is known about how mechanical overload is converted into the intracellular signals that stimulate the differentiation of fibroblasts into myofibroblasts and how mechanical forces regulate SMA gene expression in cardiac fibroblasts.
SMA expression in cultured fibroblasts is modulated by cytokines such as transforming growth factor (TGF)-
(11) and by the compliance of the substrate (2), suggesting a role for mechanical signals in regulating SMA expression. Indeed, mechanical tension is considered to be an important factor for promoting the differentiation of cultured fibroblasts into myofibroblasts and in the expression of SMA (39). The development of in vitro model systems that can apply forces of precise amplitude and duration to integrins (13) permits analysis of the effects of mechanical stimuli on cardiac fibroblast function in vitro. Some of the responses induced by mechanical forces in cardiac fibroblasts include increased proliferation, reduced collagenolytic activity, and increased collagen production (6). Several signaling pathways that mediate these mechanically induced effects have been identified in vitro and include the ERK (29, 33), JNK (21), and p38 kinase pathways (9). In rat cardiac fibroblasts subjected to passive biaxial stretching, the ERK and JNK pathways are rapidly activated, whereas the p38 kinase pathway is unaffected (25). In contrast, tensile forces applied through integrins in rat fibroblasts activate p38, whereas ERK and JNK are unaffected (23, 43).
In myofibroblasts, SMA is an important contractile protein that is required for integrin-mediated collagen remodeling (1). Because the SMA content of fibroblasts can be altered by culturing cells on surfaces of low compliance (2) and by the duration of culture, growth of cells on substrates with defined compliance and for fixed periods provides an approach for adjusting the basal levels of SMA expression at the outset of experiments. Accordingly, we examined the role of mechanical forces on SMA expression in experiments in which the basal levels of SMA were predetermined. The results indicate that tensile force-induced regulation of SMA content is dependent on baseline levels of SMA and by the selective activation of different MAPK pathways.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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-actin (clone AC-15), desmin (clone DE-U-10), and vimentin (clone VIM13.2) as well as cytochalasin D, actinomycin D, collagenase (C-5138), and BSA were purchased from Sigma (Oakville, Ontario, Canada). Goat anti-mouse IgG2a, goat anti-mouse IgG + IgM (H+L), and goat anti-mouse IgG1 were purchased from Caltag (Burlinghame, CA). PD-98059 and SB-203580 were purchased from Calbiochem (San Diego, CA). Antibodies to p38, ERK1/2, and JNK as well as the phospho-specific antibodies for each of these kinases were purchased from Cell Signaling Technology. The monoclonal antibody (clone IST-9) to cellular fibronectin (ED-A) was purchased from ABCAM. GAPDH monoclonal antibody (clone 6C5) was purchased from Advanced Immuno Chemical Soluble; type I bovine collagen (Vitrogen) was obtained from Celltrix (Palo Alto, CA). Cell cultures. Primary cultures of cardiac fibroblasts were obtained from 1-day-old, 10-g Wistar rats. Animal experiments were conducted in accordance with guidelines established by the University of Toronto Animal Care and Use Committee, who provided approval for these experiments. In brief, rats were killed, and the hearts were quickly removed under sterile conditions. Ventricular tissue was excised, minced, and digested with 0.3% collagenase containing 1.8% (wt/vol) sorbitol, 0.05% (wt/vol) DNase, 6.25 U/ml elastase, and 0.05% (wt/vol) trypsin in Krebs buffer with Zn2+. Nonadherent cells (primarily myocytes, leukocytes, and endothelial cells) were washed away. Cardiac fibroblasts attached to culture dishes within 15 min and proliferated much more rapidly than the other cardiac cell types. These properties enabled us to obtain virtually pure cultures of fibroblasts (>98% by immunophenotyping for vimentin and the absence of desmin). Cells were maintained in high-growth Dulbecco's modified Eagle's (HG-DME) medium containing 10% fetal bovine serum and a 1:10 dilution of an antibiotic solution [0.17% (wt/vol) penicillin V, 0.1% gentamycin sulfate, and 0.01 µg/ml amphotericin; Sigma] at 37°C in a humidified incubator gassed with 95% O2-5% CO2. Cells were passaged with 0.01% trypsin (GIBCO-BRL; Burlington, Ontario, Canada). Studies were performed on cells at days 1–3 in HG-DME serum-free medium.
Bead coating. As described earlier (13, 23), 0.4 g of magnetite beads (Sigma) were incubated for 1 h with 1 ml of an acidic bovine collagen solution (>95% type I collagen) at 37°C and neutralized to pH 7.4 with 100 µl of 1 N NaOH. Under these conditions, collagen polymerizes and forms fibrils around the beads within 30 min. The beads were sonicated to eliminate clumps and then dispersed. Analysis of bead size was performed by electronic particle counting (Coulter Channelyzer, Coulter Electronics; Hialeah, FL). Particles tended to exhibit a heterogeneous size distribution with a pronounced modal peak at 5 µm, although there were many particles with smaller diameters. Beads were rinsed in PBS, washed three times, and resuspended in Ca2+Mg2+-free PBS.
Force application. A ceramic permanent magnet (grade 8, 2.2 x 9.6 x 11 cm, Jobmaster; Mississauga, Ontario, Canada) was used to apply perpendicular forces to beads attached to the dorsal surface of cells. For all experiments, the pole face was parallel with and 2 cm from the cell culture dish surface. At this distance, the force on a single fibroblast with 
750-µm2 area of dorsal bead coverage was 480 pN or 0.65 pN/µm2, force levels that would be expected to be generated in myocardium (18). As the surface area of the magnet was larger than the culture dishes, and as the bead covering was relatively uniform for all cells, the forces applied to cells across the width of the culture dish were relatively uniform (43). Constant forces of varying duration were used for all experiments. Before incubation with cells, beads were rinsed in PBS, washed three times, resuspended in calcium-free buffer, and added to attached cells in complete medium for 10 min. Cells were washed three times to remove unbound beads before exposure to force.
Immunofluorescence and immunoblotting. We assessed SMA content in early passage cultures by immunostaining for SMA with 1A4 antibody, followed by FITC-conjugated goat anti-mouse IgG. Cells were examined in an epifluorescence microscope and photographed. For immunoblots, protein from beads or cell lysates prepared from cell cultures (60-mm dishes) that had been subjected to an applied force for specific time intervals were analyzed. Cells were rinsed with PBS, lysed by adding 200 µl of SDS sample buffer [62.5 mM Tris · HCl (pH 6.8), 2% SDS, 10% glycerol, 50 mM dithiothreitol, 0.1% (wt/vol) bromophenol blue], and transferred to a microfuge tube. The samples were kept on ice and then boiled for 5 min. Protein concentration was assessed by a Bio-Rad assay, and equal amounts of protein were loaded in each lane. Isolated proteins were separated by SDS-PAGE (10% acrylamide) and transferred to nitrocellulose membranes. Blots were blocked for 1 h with 5% skim milk in PBS and incubated with the indicated antibody (SMA, -actin, GAPDH, JNK, ERK1/2, and p38 diluted 1:1,000 in 0.5% Tween-PBS) for 1 h at room temperature. Blots were washed with 0.5% Tween-PBS for 10 min, incubated with appropriate second antibodies for 1 h, washed four times in Tween-PBS, and developed by enhanced chemiluminescence (Amersham). X-OMAT Kodak films were exposed to the blots, and the density of the bands was analysis by IP Lab Gel Scientific Image Processing (Signal Analytics; Vienna, VA).
Northern blot analysis. Total RNA was isolated from cells with the QIAGEN RNAeasy Total RNA kit according to the manufacturer's instructions and quantified by spectrophotometry (Ultrospec 3000, Pharmacia Biotech; Montreal, Quebec, Canada). RNA samples (10 µg) were separated in 1.2% denaturing agarose gels containing 2.2 M formaldehyde in MOPS running buffer, transferred to a nitrocellulose membrane (OPTITRAN, Schleicher & Schuell), cross-linked by ultraviolet light, and hybridized with 32P-labeled oligonucleotide probes. These probes were designed from portions of the sequences of the rat -SMA mRNA 5'-untranslated region (5'-GAAAAGAACTGAAGGCGCTGATCCACAAAACATTCACAGTTG-3') and from the rat -actin mRNA 3'-untranslated region (5'-CGCCTTCACCGTTCCAGTTTTTAAATCCTTGAGTCAAAAGCGCCA-3').
The oligonucleotides were synthesized by the Biotechnology Service Centre (Hospital for Sick Children, Toronto, Ontario, Canada). Probes were labeled with [32P]ATP (Du-Pont-NEN; Oakville, Ontario, Canada) using 3' end labeling. The blots were washed four times with 0.5% SSC + 0.5% SDS at room temperature for 10 min and twice for 40 min at 50°C and then exposed to Kodak X-OMAT films at -70°C.
Statistical analysis. For all assays, three or more separate experiments were performed. Means ± SE were calculated for continuous variables and, when appropriate, comparisons between two groups were analyzed by unpaired t-tests, or when multiple comparisons were made, ANOVA was performed by Tukey's test. In all instances, statistical significance was set at P < 0.05.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Culture of myofibroblasts. On the basis of previous data of human gingival fibroblasts, which normally synthesize minimal SMA in vivo but can be induced to express SMA when cultured on rigid substrates (2), we examined SMA content in primary cultures of fibroblasts obtained from 1-day-old rat myocardium. Cells were plated on rigid tissue culture plastic, incubated in 10% fetal bovine serum for up to 3 days, and evaluated by immunofluorescence. The composition of the cultures was >98% desmin negative, vimentin positive, and weakly SMA positive (Fig. 2), and the cells consistently showed fibroblastic morphology by phase contrast microscopy. As cardiac myocytes do not express SMA (38), and as vascular smooth muscle cells express desmin, culture of adherent cells for 1–3 days showed that these cells were indeed fibroblasts (desmin negative, SMA positive, and vimentin positive). Under these conditions, staining for SMA and ED-A fibronectin (another myofibroblast marker) was very low on the first day of culture (Fig. 2), but within 2 days of plating there was abundant SMA and increased ED-A fibronectin. At 3 days, cells were brightly stained for SMA (predominantly in stress fibers) and for ED-A fibronectin.
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Immunoblots of SMA, fibronectin ED-A, and GAPDH (as a loading control) in cells cultured with 10% serum showed large increases of SMA and fibronectin ED-A between 1 and 3 days in culture, whereas the relative abundance of GAPDH did not change appreciably. We calculated the ratio of SMA to GAPDH and found significant (2.2-fold, n = 4, P < 0.001 by ANOVA) increases of SMA between 1 and 3 days in culture (Fig. 3A). In cells cultured without serum, there was no change of SMA between 1 and 3 days in culture (Fig. 3B). These data are consistent with earlier work showing that induction of SMA expression is mediated by serum response factor binding to the CArG-B box in the SMA promoter (44).
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Culture conditions for determination of SMA content. Myofibroblast differentiation is dependent in part on the mechanical stiffness of the extracellular matrix to which cells are attached (39). In addition to conventional, rigid tissue culture plastic, we used collagencoated agar gels (11a) (Young's modulus = 50,000 N/m2) to more closely model the rheological properties of the myocardium. As shown earlier (2), more compliant substrates reduce the mechanical tension generated by cultured fibroblasts and allowed us to adjust basal SMA expression. Neonatal cardiac fibroblasts were cultured for 1 and 2 days on collagen-coated agar or collagen-coated tissue culture plastic with equivalent amounts of collagen coating. Cells on collagencoated agar showed no rhodamine phalloidin-stained stress fibers, whereas cells on collagen-coated plastic showed abundant stress fibers (Fig. 3C), indicating that cells on the soft gels exhibited relatively less tension. Cells were lysed, and equal amounts of cellular proteins were evaluated by immunoblotting. There was abundant SMA in cells cultured for 1 or 2 days on rigid tissue culture plastic, whereas cells plated on collagen-coated agar exhibited no detectable SMA (Fig. 3C). The cellular content of -actin was only very slightly reduced by plating cells on the softer substrate.
Effect of exogenous mechanical tension on SMA expression. We next determined whether exogenously applied tensile forces can regulate SMA expression in cardiac fibroblasts. Cardiac fibroblasts from 1-day-old rats were subjected to static tensile forces (0.65 pN/µm2) delivered via collagen-coated magnetite beads (43). As shown by immunostaining (Fig. 4A), force application induced different effects on SMA and -actin content. There was only weak staining for SMA in cells loaded with beads but without force application. After 4 h of force, there was strong staining for SMA, which was enhanced near the bead locus. In contrast, application of force caused no significant change of -actin staining. In cells cultured in serum, immunoblotting showed that SMA was increased after 2–4 h of constant force application, whereas the relative abundance of GAPDH did not change (Fig. 4B). We quantified the change of SMA by calculating the ratio of SMA to GAPDH and found a 1.6-fold increase of this ratio between 0 and 4 h after the application of force (n = 4, P < 0.05 by ANOVA). There was no effect of force on SMA content if cells were cultured without serum. The force-induced increases of SMA did not affect the total GAPDH content. These results indicated that in neonatal cardiac fibroblasts with low basal levels of SMA, exogenously applied tensile forces increase SMA content.
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We examined the effect of force on SMA mRNA in neonatal cardiac fibroblasts cultured for 1 day. After force application, SMA and -actin mRNA content were measured by Northern blotting using oligonucleotide probes specific for these isoforms. The relative abundance of SMA mRNA normalized to -actin mRNA was increased 1.7-fold after force application (n = 3, P < 0.05 by t-test; Fig. 5). The increase of SMA mRNA after force was reduced by treatment with actinomycin D (5 µg/ml, P > 0.2 compared with controls without force). The force-induced increase of SMA required intact actin filaments because cells preincubated with cytochalasin D (1 µM, 20 min) and subjected to applied tensile force showed no increase of the ratio of SMA to -actin (n = 4, P < 0.05 by ANOVA; Fig. 6).
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Role of MAPKs and basal levels of SMA in forceinduced myofibroblast differentiation. Force-induced gene expression in cardiac cells is thought to be mediated in part by MAPKs (25, 43). We determined whether mechanical force-induced regulation of SMA content is dependent on selective activation of MAPKs. Phosphorylated and nonphosphorylated forms of p38, JNK, and ERK were examined after the application of force in cells from 1-day-old rats that were cultured for 1 day (i.e., low levels of basal SMA). Positive controls used PMA or ultraviolet light to activate ERK or JNK, respectively. Force induced a time-dependent increase of phosphorylated ERK (Fig. 7A). We quantified the blot densities of phosphorylated and total ERK and expressed these densities as a ratio. Within 0.5 h after force application, there was a nearly 2.8-fold increase of force, which was reduced to 2.1- and 2.4-fold at 2 and 4 h after continuous force application (n = 4, P < 0.01 above 0-h control by ANOVA). If cells were pretreated with the ERK inhibitor PD-98059 (50 µM) and then with applied force, the force-induced increase of SMA was blocked, as was ERK phosphorylation (Fig. 7A). Force did not affect p38 and JNK activation, although ultraviolet light (as a positive control) activated p38 and JNK (Fig. 7B).
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We then determined whether SMA expression and MAPK activation in response to applied forces are dependent on the basal levels of SMA. When cardiac fibroblasts from 1-day-old rats were cultured on rigid substrates for 3 days, there was a spontaneous development of high levels of SMA (Fig. 2). After force application for 2 or 4 h to cells with high basal levels of SMA, immunoblots of SMA and GAPDH showed reduced SMA content but no effect on GAPDH content (n = 4, P < 0.05 by ANOVA; Fig. 8A). We determined whether the reduction of SMA content may be due to selective activation of MAPKs. Immunoblots of phosphorylated and nonphosphorylated forms of p38, JNK, and ERK1/2 were prepared from cells that were previously stimulated with force. There was a time-dependent increase of phosphorylated p38, which was totally blocked by the p38 kinase inhibitor SB-203580 (10 µM; Fig. 8B). Quantification of the ratio of phosphorylated p38 to total p38 showed that force increased the ratio by greater than twofold within 0.5 h (n = 4, P < 0.01 by ANOVA). Treatment with SB-203580 completely blocked the force-induced reduction of SMA (Fig. 8A) and, as expected, phosphorylation of p38. Positive control cultures treated with ultraviolet light also showed greatly increased phosphorylated p38 kinase. Force did not promote ERK1/2 activation, although cells treated with PMA as a positive control exhibited abundant phosphorylated ERK1/2 (Fig. 8B). Similarly, force had no effect on JNK, whereas stimulation with ultraviolet light activated JNK (Fig. 8B).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Cardiac fibroblast model systems. The mechanisms that regulate the differentiation and persistence of myofibroblasts in cardiac lesions are not defined, as are the mechanisms that regulate deletion of myofibroblasts in physiological situations (10). We have used neonatal cardiac fibroblasts subjected to applied tensile forces as a model for studying the promotion of cardiac myofibroblast differentiation by hemodynamic stress independent of humoral or inflammatory factors (28). Fibroblasts comprise approximately two-thirds of myocardial cells (14) but are not the only cells in the myocardium capable of expressing SMA. Whereas postnatal cardiomyocytes express negligible or very low levels of SMA (38, 47), smooth muscle cells contain very large amounts of SMA. Accordingly, to establish the validity of these in vitro studies, it was important that analyses were restricted to fibroblasts [i.e., cells with fibroblastic morphology and that were vimentin positive and desmin negative but retained the capacity of expressing SMA (34)]. After single cell suspensions were prepared from ventricular tissues, cardiomyocytes and smooth muscle cells failed to attach after short-term (i.e., 15 min) adherence to culture dishes; as a result, fibroblasts were enriched on the basis of attachment (>98% of cells fulfilled the phenotypic criteria for fibroblasts). Consequently, we were confident that analyses of the myofibroblast differentiation program described here were restricted to fibroblasts.
We used neonatal cardiac fibroblasts as we found in preliminary experiments that these cells initially express very low levels of SMA in culture but respond rapidly to tensile force with increased SMA expression. In this context, neonatal cardiac fibroblasts provide a permissive model system in which force-induced expression of SMA rises rapidly from very low baseline levels. Similar, rapid responses to tensile forces have been observed in rat osteosarcoma cells, which also have extremely low levels of SMA in unstimulated, basal conditions (44). The increase of SMA in cardiac fibroblasts appeared to be selective for SMA because there was no evidence of a force-induced change of -actin or GAPDH. In a previous study (43), we demonstrated that fibroblasts from the adult rat heart can differentiate into myofibroblasts when cultured on rigid substrates, a culture method that facilitates generation of high levels of intracellular tension (2) and very high levels of SMA. Thus the in vitro behavior of neonatal fibroblasts when subjected to tensile forces shown here is similar to adult cardiac fibroblasts and helps to rationalize the model described herein.
Cardiac fibroblasts adhere to extracellular matrixes through integrins, which provide sites for transfer of tensile forces to the actin cytoskeleton (25). Expression of collagen receptors by cardiac fibroblasts (5) provides the attachment for collagen-coated beads and the linkages to the actin cytoskeleton that, as we showed here in experiments with cytochalasin D, are essential for force-induced regulation of SMA. Furthermore, we (42, 43) have shown previously that cardiac fibroblasts exhibit specific changes of SMA content to forces applied through collagen but not BSA or poly-L-lysine-coated beads, supporting the view that the force effect was indeed mediated by specific collagen receptors, which may include integrins or discoidin receptors. We should emphasize that the collagen receptor is not itself the mechanotransducer and almost certainly involves other processes including, for example, stretchactivated channels and/or voltage-sensitive calcium channels.
SMA is regulated by mechanical force. In chronic cardiac volume or pressure overload, exogenous physical forces are converted into intracellular signals that stimulate the differentiation of cardiac fibroblasts into myofibroblasts (22). This differentiation process is characterized by expression of SMA in fibroblasts (34). Our results show that increased expression of SMA in cultured cardiac fibroblasts can be induced either by the application of exogenous force via integrins (i.e., collagen-coated magnetite beads) or by the culture of cells on rigid substrates to facilitate the endogenous generation of tension. Indeed, tensile force is hypothesized to be a crucial determinant of SMA expression by fibroblasts (39). As SMA is an important contractile protein in fibroblasts (16) as well as a phenotypic marker for myofibroblast (34), the regulation of SMA by mechanical force in the myocardium is of medical importance because of the central role of myofibroblasts in synthesizing extracellular matrix proteins (8) and their contribution to cardiac fibrosis and hypertrophy (12).
We (43) have previously shown that when fibroblasts from adult rat cardiac tissues are cultured on rigid substrates, they express very high levels of SMA, and, after application of tensile forces, SMA expression is reduced. Similarly, as shown in the present study, when fibroblasts from the neonatal rat ventricle for 3 days were cultured on rigid substrates, SMA was increased but, thereafter, the SMA content was reduced after application of tensile forces. Therefore, the basal SMA content and the relative abundance of polymerized actin may be important regulators of SMA synthesis in response to applied forces (3, 30).
Our results showed that exogenously applied mechanical forces could increase SMA protein and mRNA expression in neonatal cardiac fibroblasts cultured for 1 day and with very low basal levels of SMA. The force-induced increase of SMA mRNA was largely blocked by treatment with actinomycin D, indicating that at least part of the force effect was due to increased production of nascent SMA transcripts. Accordingly, SMA may be a mechanotranscriptionally regulated gene (44).
Force-induced activation of MAPKs is dependent on myofibroblast differentiation. The influence of mechanical force on gene expression in many cell types is thought to be mediated in part through the MAPK pathway (17, 19, 21, 32). As MAPKs have been implicated in force-induced gene expression in rat cardiac fibroblasts (25, 43), we measured the activation of MAPKs in response to mechanical force in cells with different basal SMA content and differentiation status. In cells obtained from 1-day-old rats and cultured for 1 day, force induced a time-dependent increase of phosphorylated ERK, similar to results from a previous report (25) using biaxial stretching of cardiac fibroblasts. The force-induced activation of ERK was apparently important for the increase of SMA, as cells that were treated with force and the ERK inhibitor PD-98059 showed no change of SMA content. Notably, force did not activate p38 or JNK. In quiescent cardiac fibroblasts, ERK and JNK can be stimulated by static equibiaxial stretching forces (24), and ERKs may be involved in mechanical force-induced cardiac hypertrophy (31).
In contrast to cardiac fibroblasts from neonatal rats grown for 1 day, cells cultured on rigid substrates for 3 days showed spontaneous development of high expression levels of SMA. This effect apparently required generation of endogenous tension because culture of the same cells on a highly compliant substrate or incubation with cytochalasin D blocked this increase. In cells with high basal levels of SMA, force induced a time-dependent increase of phosphorylated p38, which was completely blocked by the p38 kinase inhibitor SB-203580. Force did not activate ERK1/2 or JNK. The p38 inhibitor SB-203580 also blocked the force-induced reduction of SMA. Notably, in cardiac myocytes, the p38 -isoform is involved in regulating apoptosis (45), in stretch-induced inhibition of SMA in adult cardiac fibroblasts (43), and of the -skeletal actin gene when transfected in fibroblasts (23).
In summary, our findings suggest that force-induced SMA expression was dependent on ERK phosphorylation and activation of myofibroblast differentiation. In well-differentiated myofibroblasts with abundant SMA content, force-induced activation of p38 may induce an inhibitory pathway (e.g., protein kinase R; unpublished observations) that blocks SMA transcription. Collectively, these results indicate that tensile force-induced activation of MAPKs is dependent on the myofibroblast differentiation status.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
-smooth muscle actin expression by fibroblasts. J Cell Physiol 159: 161–175, 1994.[Web of Science][Medline]
induction of -smooth muscle actin in fibroblasts. Am J Pathol 154: 871–882, 1999.1 in cardiac fibroblasts and myofibroblasts. J Mol Cell Cardiol 29: 1947–1958, 1997.[Web of Science][Medline]
1 integrin-mediated transcriptional circuit that regulates the actin-binding protein filamin-A. J Biol Chem 277: 47541–47550, 2002.1 induces -smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J Cell Biol 122: 103–111, 1993. upregulates angiotensin II type 1 receptors on cardiac fibroblasts. Circ Res 85: 272–279, 1999.-Smooth muscle actin expression upregulates fibroblast contractile activity. Mol Biol Cell 12: 2730–2741, 2001.1 integrins and tyrosine kinases. Circ Res 79: 310–316, 1996.-actin gene transcription by applied mechanical forces through integrins and actin. Biochem J 341: 647–653, 1999.[Medline]
-Actin isoform distribution in normal and failing human heart: a morphological, morphometric, and biochemical study. J Pathol 199: 387–397, 2003.[Web of Science][Medline]
1 promotes the morphological and functional differentiation of the myofibroblast. Exp Cell Res 257: 180–189, 2000.[Web of Science][Medline]
-smooth muscle actin. Tissue Cell 33: 86–96, 2001.[Web of Science][Medline]
-Smooth muscle actin is transiently expressed in embryonic rat cardiac and skeletal muscles. Differentiation 39: 161–166, 1988.[Web of Science][Medline]This article has been cited by other articles:
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L. Chilton, S. Ohya, D. Freed, E. George, V. Drobic, Y. Shibukawa, K. A. MacCannell, Y. Imaizumi, R. B. Clark, I. M. C. Dixon, et al. K+ currents regulate the resting membrane potential, proliferation, and contractile responses in ventricular fibroblasts and myofibroblasts Am J Physiol Heart Circ Physiol, June 1, 2005; 288(6): H2931 - H2939. [Abstract] [Full Text] [PDF]
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 H. Shiratsuchi and M. D. Basson Activation of p38 MAPK{alpha} by extracellular pressure mediates the stimulation of macrophage phagocytosis by pressure Am J Physiol Cell Physiol, May 1, 2005; 288(5): C1083 - C1093. [Abstract] [Full Text] [PDF]
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H. Shiratsuchi and M. D. Basson Extracellular pressure stimulates macrophage phagocytosis by inhibiting a pathway involving FAK and ERK Am J Physiol Cell Physiol, June 1, 2004; 286(6): C1358 - C1366. [Abstract] [Full Text] [PDF]
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