Abl is a nonreceptor tyrosine kinase that is required for smooth muscle contraction. However, the mechanism by which Abl regulates smooth muscle contraction is not completely understood. In the present study, Abl underwent phosphorylation at Tyr412 (an index of Abl activation) in smooth muscle in response to contractile activation. Treatment with a cell-permeable decoy peptide, but not the control peptide, attenuated Abl phosphorylation during contractile stimulation. Treatment with the decoy peptide did not affect the association of Abl with the cytoskeletal protein vinculin and the spatial location of vinculin in smooth muscle. Inhibition of Abl phosphorylation by the decoy peptide attenuated the agonist-induced phosphorylation of Crk-associated substrate (CAS), an adapter protein participating in the signaling processes that regulate force development in smooth muscle. Additionally, previous studies have shown that contractile stimulation triggers the dissociation of CAS from the vimentin network, which is important for cytoskeletal signaling and contraction in smooth muscle. In this report, the decrease in the amount of CAS in cytoskeletal vimentin in response to contractile activation was reversed by the Abl inhibition with the decoy peptide. Moreover, force development and the enhancement of F-actin-to-G-actin ratios (an indication of actin polymerization) upon contractile activation were also attenuated by the Abl inhibition. However, myosin phosphorylation induced by contractile activation was not affected by the inhibition of Abl. These results suggest that Abl regulates the dissociation of CAS from the vimentin network, actin polymerization, and contraction by modulating CAS phosphorylation in smooth muscle.
- tyrosine kinase
- Crk-associated substrate
smooth muscle contraction plays a fundamental role in regulating the functions of the hollow organs in the body such as blood vessels and the airways. Abnormal smooth muscle contraction may contribute to the development of many diseases such as hypertension and asthma. Despite its important role, the mechanisms that regulate smooth muscle contraction are not completely understood.
Cross-bridge cycling regulated by myosin regulatory light chain (MRLC) phosphorylation has been thought to be the solely fundamental paradigm for the regulation of vascular smooth muscle contraction (12, 23). However, recent studies suggest that actin polymerization transpires in smooth muscle in response to contractile activation. The inhibition of actin polymerization by such inhibitors as cytochalasin D or latrunculin A prevents force development during activation with contractile stimuli without affecting MRLC phosphorylation (1, 16, 26, 27). These studies suggest that actin polymerization and myosin phosphorylation are independently regulated, and that both the dynamic change in the actin cytoskeleton and myosin activation are required for force development during contractile stimulation of smooth muscle (2, 13, 16, 20, 26, 27).
Abl (Abelson tyrosine kinase, c-Abl) is a nonreceptor tyrosine kinase that has recently been identified as a key regulator of smooth muscle cytoskeletal signaling and contraction (1, 4). Latest studies have shown that Abl has a multifaceted role in cells. Besides kinase activity, Abl may also serve as a scaffold protein to interact with structural proteins (23, 33). We have previously shown that contractile activation of smooth muscle induces Abl phosphorylation at Tyr412 (an indication of Abl activation). Moreover, silencing of Abl by RNA interference attenuates force development in smooth muscle (1). These studies suggest that Abl is required for smooth muscle contraction. Nevertheless, these studies cannot distinguish whether the effect of Abl knockdown is due to its phosphorylation or scaffold function. Answer to this question is important because this may improve our understanding regarding how Abl regulates cytoskeletal signaling in smooth muscle.
Abl affects smooth muscle contraction in part by regulating phosphorylation of Crk-associated substrate (CAS) as evidenced by in vitro biochemical analysis and studies on smooth muscle tissues (1). CAS is an actin-regulatory protein that has been shown to regulate smooth muscle contraction. The downregulation of CAS by antisense oligonucleotides dramatically attenuates force development and actin polymerization in smooth muscle in response to contractile stimulation (27). Moreover, CAS is able to bind to the vimentin intermediate filament network in smooth muscle, which may inhibit its role in cytoskeletal signaling and force development (32). Contractile stimulation induces the dissociation of CAS from cytoskeletal vimentin, which has been implicated in the regulation of smooth muscle contraction. For example, the dissociation of CAS from cytoskeletal vimentin may facilitate the translocation of CAS to the membrane promoting cortical actin polymerization and smooth muscle contraction (15, 22, 27, 32). However, the mechanism that regulates the dissociation of CAS from the vimentin network is not well elucidated.
In this study, we have utilized cellular and molecular approaches to evaluate the role of Abl phosphorylation in regulating CAS phosphorylation, the interaction of CAS with the vimentin cytoskeleton, actin filament assembly, and force development in smooth muscle.
MATERIALS AND METHODS
All animal protocols were approved by the Institutional Animal Care and Use Committee of Albany Medical College. Male Sprague-Dawley rats (225–250 g) were euthanized in a CO2 chamber and carotid arteries were removed and placed at room temperature in physiological saline solution (PSS) containing (in mM) 110 NaCl, 3.4 KCl, 2.4 CaCl2, 0.8 MgSO4, 25.8 NaHCO3, 1.2 KH2PO4, and 5.6 glucose. The solution was aerated with 95% O2-5% CO2 to maintain a pH of 7.4. After removal of the endothelium and connective tissue layer, carotid smooth muscle rings (5 mm long) were placed in PSS at 37°C in a 25-ml organ bath and attached to a Grass force transducer that had been connected to a computer with A/D converter (Grass). Passive tension (0.5–0.75 g) was applied to smooth muscle at the beginning of each experiment. After 10–20 min equilibrium they were stimulated with 80 mM KCl repeatedly until stable contractile responses were reached. For biochemical analysis, smooth muscle rings were frozen using liquid N2-cooled tongs and then pulverized under liquid N2 with a mortar and pestle. Unstimulated smooth muscle tissues were treated with 10−6 M phentolamine for 5 min to prevent the effects of potential vasoconstrictor release from smooth muscle tissues.
Production of a cell-permeable decoy peptide.
A cell-permeable decoy peptide (Abl-I) was produced using the pGEX-4T system (15). The sequence of Abl-I peptide was taken from the kinase domain of Abl (residues 350-463) (National Center for Biotechnology Information accession no. DQ294402). The peptide was conjugated with TAT sequence for cell permeability. TAT peptide is a short polybasic sequence derived from the human immunodeficiency virus TAT protein; it has been shown to successfully deliver a large variety of cargos (from small particles to peptides and proteins) into intact cells/tissues (3). Briefly, cDNA sequence encoding TAT and Abl-I peptides was subcloned into pGEX-4T followed by transformation into Escherichia coli BL21. The glutathione S-transferase (GST)-tagged peptide was harvested and purified according to Amersham's manual of GST gene fusion system. Using a similar approach, a TAT-GST chimera peptide was also produced as a control.
Pulverized tissues were mixed with 40 μl SDS sample buffer [1.5% dithiothreitol (DTT), 2% SDS, 80 mM Tris·HCl (pH 6.8), 10% glycerol, and 0.01% bromophenol blue] for 5 min and separated by SDS-PAGE. Proteins were transferred to a nitrocellulose membrane, after which the membrane was blocked with 2% bovine serum albumin (BSA) for 1 h and probed with site-specific, state-dependent antibody for phospho-CAS (Tyr410, Cell Signaling) or phospho-Abl (Tyr412, Cell Signaling) followed by horseradish peroxidase (HRP)-conjugated anti-rabbit Ig (ICN Biomedicals, Irvine, CA)(24). Proteins were visualized by enhanced chemiluminescence (SuperSignal, Pierce) using the Fuji Image System LAS-4000. The membranes were stripped and reprobed with monoclonal antibodies against total CAS (clone 24, BD Biosciences) or total Abl (clone 8E9, BD Biosciences) followed by the incubation with HRP-conjugated anti-mouse Ig (Amersham Life Sciences) (24). The levels of phosphoprotein and total protein were quantified by scanning densitometry of immunoblots (Fuji Multigauge software). Changes in protein phosphorylation are expressed as a magnitude increase over levels of phosphorylation in unstimulated tissues. The luminescent signals from all immunoblots were within the linear range. To determine Src phosphorylation, phospho-Src (Tyr416) antibody (Cell Signaling) and pan-Src antibody (Santa Cruz Biotechnology, Santa Cruz, CA) were used.
Smooth muscle tissues were placed in frozen tissue-embedding medium (Neg 52, Richard-Allen Scientific) and cryosectioned using Cryostats (Richard-Allen Scientific). For cultured cell experiments, cells were plated in dishes containing coverslips and incubated in serum-free media for 1 day. Tissues or cells were fixed for 15 min in 4% paraformaldehyde and were then washed three times in Tris-buffered saline (TBS) containing 50 mM Tris, 150 mM NaCl, and 0.1% NaN3 followed by permeabilization with 0.2% Triton X-100 dissolved in TBS for 5 min. These cells were incubated with GST antibody (Neomarkers, Fremont, CA) or vinculin antibody (Santa Cruz Biotechnology), or smooth muscle α-actin antibody (Sigma) followed by appropriate secondary antibody conjugated to Alexa-488 or Alexa-543 (Molecular Probes, Eugene, OR). The cellular localization of fluorescently labeled proteins was viewed under a high-resolution digital fluorescent microscope.
Protein interaction was assessed by coimmunoprecipitation analysis. One-two smooth muscle segments were pooled for each measurement of coimmunoprecipitation. Pulverized tissues were mixed with Triton buffer, and protein extracts were collected and precleared. The extracts were incubated overnight with vinculin antibody and then incubated for 2–3 h with a 10% suspension of protein A-Sepharose beads. Immunocomplexes were washed four times in buffer containing 50 mM Tris·HCl (pH 7.6), 150 mM NaCl, and 0.1% Triton X-100. The immunoprecipitates were separated by SDS-PAGE followed by transfer to nitrocellulose membranes. Blots of the precipitates were probed with use of vinculin antibody and Abl antibody.
Analysis of CAS dissociation from cytoskeletal vimentin.
Insoluble vimentin was collected from smooth muscle tissues by using a previously described method (15, 32). The equal amount of cytoskeletal vimentin was separated by 10% SDS-PAGE and was transferred to nitrocellulose membranes. The membranes were cut into two parts; the upper part was probed with monoclonal CAS antibody (clone 24, BD Biosciences). The lower part of the membrane was blotted with vimentin antibody (clone RV202, BD Biosciences). The ratio of CAS to vimentin was calculated after densitometrical analysis of immunoblots.
Analysis of F-actin-to-G-actin ratios.
The content of F-actin and G-actin in smooth muscle was measured using a previously described method (1, 25, 28, 34). Briefly, smooth muscle was homogenized in 150 μl F-actin stabilization buffer (50 mM PIPES, pH 6.9, 50 mM NaCl, 5 mM MgCl2, 5 mM EGTA, 5% glycerol, 0.1% Triton X-100, 0.1% Nonidet P40, 0.1% Tween 20, 0.1% β-mercaptoethanol, 1 mM ATP, 1 μg/ml pepstatin, 1 μg/ml leupeptin, and 10 μg/ml benzamidine). The supernatants of protein extracts were collected after centrifugation at 151,000 g for 60 min at 37°C. The pellets were resuspended in ice-cold H2O plus 1 μM cytochalasin D and then incubated on ice for 1 h to dissociate F-actin. The resuspended pellets were gently mixed every 15 min. The supernatant of the resuspended pellets was collected after centrifugation at 16,100 g for 2 min at 4°C. Equal volume of the first supernatant (G-actin) or second supernatant (F-actin) was subjected to immunoblot analysis using α-actin antibody. The amount of F-actin and G-actin was determined by scanning densitometry.
Analysis of MRLC phosphorylation.
Smooth muscle was rapidly liquid nitrogen frozen and immersed in precooled acetone containing 10% (wt/vol) trichloroacetic acid and 10 mM dithiothreitol (acetone-TCA-DTT). Tissues were thawed in acetone-TCA-DTT at room temperature and then washed four times with acetone-DTT mixture. Proteins were extracted for 4 h in 8 M urea, 20 mM Tris base, 22 mM glycine, and 10 mM DTT. MRLC was separated by glycerol-urea PAGE and transferred to nitrocellulose membranes. The membranes were blocked with 5% milk and incubated with myosin light chain 20 antibody. The primary antibody was reacted with HRP-conjugated anti-rabbit IgG (ICN Biomedicals). Unphosphorylated and phosphorylated bands of MRLC were visualized by enhanced chemiluminescence and quantified by scanning densitometry. MRLC phosphorylation was calculated as the ratio of phosphorylated MRLC to total MRLC.
Primary vascular smooth muscle cells were isolated from rat aorta. Two aortic segments (10 mm long) were incubated for 10 min with 5 ml dissociation solution [130 mM NaCl, 5 mM KCl, 1.0 mM CaCl2, 1.0 mM MgCl2, 10 mM HEPES, 0.25 mM EDTA, 10 mM d-glucose, 10 mM taurine, pH 7, 4.5 mg collagenase (type I), 10 mg papain (type IV), 2 mg elastase, 1 mg tyrosine inhibitor, 1 mg/ml BSA, and 1 mM DTT]. All enzymes were obtained from Sigma. The tissues were then washed three times with a HEPES-buffered saline solution (composition in mM: 10 HEPES, 130 NaCl, 5 KCl, 10 glucose, 1 CaCl2, 1 MgCl2, 0.25 EDTA, and 10 taurine, pH 7), and they were triturated with a pipette to liberate individual smooth muscle cells from the tissue. The cell suspension was mixed with Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% (vol/vol) fetal bovine serum (FBS) and antibiotics (100 U/ml penicillin, 100 μg/ml streptomycin). Cells were cultured in 100-mm dishes at 37°C in the presence of 5% CO2 in the same DMEM. The medium was changed every 3–4 days until the cells reached confluence, and confluent cells were passaged with trypsin-EDTA solution. Cells within passage 2 were then serum deprived 24 h before immunofluorescence experiments. These smooth muscle cells in culture express high levels of smooth-muscle-specific α-actin, as determined by immunoblot analysis.
All statistical analysis was performed using Prism 4 software (GraphPad Software, San Diego, CA). Comparison among multiple groups was performed by one-way analysis of variance followed by Tukey's multiple-comparison test. Differences between pairs of groups were analyzed by Student-Newman-Keuls test or Dunn's method. Values of n refer to the number of experiments used to obtain each value. P < 0.05 was considered to be significant.
Characterization of a cell-permeable decoy peptide (Abl-I).
We assessed whether the decoy peptide Abl-I is able to enter the smooth muscle tissue. Endothelium-free carotid smooth muscle preparation was treated with 2 μg/ml GST-tagged Abl-I or the control peptide for 60 min. Cryosection of the preparation was immunostained for GST to visualize the tagged peptides. The tissue sections were also immunostained for smooth muscle α-actin to identify the smooth muscle layer. Immunostaining images showed that the GST-tagged peptides were homogenously distributed in the medial smooth muscle layer, suggesting an effective penetration of the peptides into smooth muscle tissues (Fig. 1).
In subsequent studies, we have determined Abl phosphorylation (an index of Abl activation) in smooth muscle tissues treated with the peptides. Phenylephrine (PE) is an agonist of α-receptor that plays an important role in regulating vascular smooth muscle contractility and blood pressure (1, 4, 13). In addition, our previous studies showed that contractile force reaches a maximal level in vascular smooth muscle in response to stimulation with PE for 5 min (1). Stimulation with PE (10 μM, 5 min) induced Abl phosphorylation in untreated smooth muscle tissues and in tissues treated with the control peptide (2 μg/ml, 60 min). However, the agonist-induced Abl phosphorylation was reduced in tissues treated with 2 μg/ml Abl-I peptide for 60 min (Fig. 2A). The phosphorylation level of Abl in response to PE stimulation was significantly lower in Abl-I-treated tissues than in untreated smooth muscle and in tissues treated with the control peptide (Fig. 2B, n = 5–6, P < 0.05). Treatment with the decoy peptide did not affect Abl phosphorylation in unstimulated tissues. The results suggest that the decoy peptide effectively inhibits Abl activation in smooth muscle in response to contractile activation.
Abl phosphorylation is mediated by Src tyrosine kinase in mammalian cells (6). To evaluate whether the inhibition of Abl phosphorylation by the decoy peptide is due to the inhibition of Src, we determined the effects of Abl-I on Src phosphorylation at Tyr416 (an indication of Src activation) during contractile activation. Treatment with Abl-I did not attenuate PE-induced Src phosphorylation (Fig. 2, C and D). The results suggest that the decoy peptide selectively inhibits Abl phosphorylation, but not Src activation. It is possible that Abl-I may compete with endogenous Abl for Src, therefore inhibiting Abl phosphorylation during contractile activation.
In addition to the kinase activity, Abl has also been proposed to serve as a scaffold protein interacting with other proteins such as vinculin, a critical component of focal adhesions that link actin filaments to the membrane (7, 18). Thus, we also assessed whether Abl-I peptide affects the scaffold function of Abl. Smooth muscle tissues were treated with Abl-I or the control peptide (2 μg/ml, 60 min). Coimmunoprecipitation analysis was used to evaluate the association of Abl with vinculin. As shown in Fig. 3B, Abl was coimmunoprecipitated with vinculin and metavinculin (muscle isoform of vinculin) in smooth muscle tissues not treated with peptides. Treatment with the peptides did not affect the amount of Abl precipitated with vinculin. Furthermore, treatment with the peptides did not alter the spatial localization of vinculin in smooth muscle cells, indicating that Abl inhibition does not disrupt the structure of focal adhesion (Fig. 3C). These results demonstrate that Abl-I peptide specifically inhibits Abl activation without affecting its scaffold function.
PE-induced CAS phosphorylation is reduced by Abl inhibition.
The adapter protein CAS has been implicated in smooth muscle cytoskeletal signaling and contraction. CAS undergoes phosphorylation at Tyr-410 in smooth muscle during contractile activation (1, 17). To assess whether CAS is regulated by Abl phosphorylation, immunoblotting analysis was utilized to evaluate CAS phosphorylation in smooth muscle.
In untreated smooth muscle, exposure to PE (10 μM, 5 min) induced CAS phosphorylation at Tyr-410, which is consistent with previous studies (1, 17). Moreover, pretreatment with the control peptide did not affect the PE-induced CAS phosphorylation. However, the phosphorylation of CAS upon contractile activation was attenuated in smooth muscle tissues treated with Abl-I peptide (Fig. 4A). The levels of CAS phosphorylation during contractile activation were significantly lower in tissues treated with Abl-I peptide than in untreated tissues and tissues treated with the control peptide (Fig. 4B, n = 4–6, P < 0.05). Treatment with the peptides did not affect CAS phosphorylation before stimulation.
We also assessed the effects of the Abl pharmacological inhibitor imatinib (Gleevec, Novartis) on CAS phosphorylation induced by PE. Treatment with imatinib (20 μM, 60 min) inhibited PE-induced CAS phosphorylation in smooth muscle (Fig. 4C).
Effects of Abl inhibition on the dissociation of CAS from cytoskeletal vimentin.
Previous studies have shown that CAS is able to bind to the vimentin cytoskeleton in smooth muscle (32). Contractile stimulation induces the dissociation of CAS from cytoskeletal vimentin, which is an important cellular process during contractile activation of smooth muscle (15, 22, 27, 32). To assess how much of the total cellular CAS pool is associated with cytoskeletal vimentin, soluble vimentin and pellet cytoskeletal vimentin were separated by the fractionation assay and were probed for CAS and vimentin (Fig. 5A). Quantification analysis showed that about 20% of total CAS was associated with cytoskeletal vimentin (Fig. 5B).
To determine the role of Abl in the protein dissociation, cytoskeletal vimentin from unstimulated and stimulated smooth muscle that had been treated with Abl-I peptide or the control peptide was separated by SDS-PAGE. Blots of the fraction were probed with antibodies against CAS and vimentin. In untreated smooth muscle and the control peptide-treated tissues, treatment with PE led to a decrease in the amount of CAS associated with cytoskeletal vimentin (Fig. 5C). Quantification analysis showed that ∼50% of CAS dissociated from cytoskeletal vimentin (Fig. 5D). In contrast, the amount of CAS in cytoskeletal vimentin upon contractile activation was higher in tissues treated with Abl-I or imatinib than in untreated smooth muscle and tissues treated with the control peptide. The ratios of CAS to vimentin in response to PE stimulation were statistically higher in smooth muscle tissues treated with Abl-I peptide or imatinib than in untreated tissues and smooth muscle treated with the control peptide (Fig. 5D, n = 4–5, P < 0.01). Treatment with the peptides did not alter the ratios of CAS over vimentin in tissues before PE stimulation.
Abl inhibition diminishes an increase in F-action-to-G-actin ratios (an index of actin polymerization) in smooth muscle during contractile activation.
To evaluate the role of Abl activation in actin dynamics, we determined the effects of Abl-I peptide on F-actin-to-G-actin (F/G-actin) ratios by the fractionation assay. As shown in Fig. 6, stimulation with PE led to the increase in F/G-actin ratios, suggesting that the actin cytoskeleton is undergoing polymerization. However, the enhancement of F/G-actin ratios in response to contractile activation was significantly suppressed in tissues treated with Abl-I peptide, but not the control peptide (P < 0.05, n = 3–4). The results indicate that Abl kinase activity is necessary to regulate actin filament polymerization in smooth muscle.
Inhibition of Abl phosphorylation by Abl-I peptide attenuates force development in smooth muscle in response to PE stimulation.
To evaluate whether Abl phosphorylation is required for smooth muscle contraction, we assessed the effects of Abl-I peptide on force development in smooth muscle. Contractile response of carotid smooth muscle tissues to PE was determined. The contractile agonist was rinsed out after reaching maximal contraction. Smooth muscle tissues were then treated with 2 μg/ml Abl-I peptide or the control peptide for 60 min. Dose-response curves were then determined.
As shown in Fig. 7, treatment with PE induced smooth muscle contraction in a dose-dependent manner. The PE dose-response curve in tissues treated with the control peptide was similar to that of untreated tissues. In contrast, the dose-response curve was right shifted with reduced maximal contraction in smooth muscle rings treated with the decoy peptide (n = 4–7, P < 0.05).
Blockage of Abl phosphorylation does not inhibit MRLC phosphorylation.
MRLC phosphorylation has been thought to be a major biochemical change during contractile activation of smooth muscle (12). To determine whether the reduced contraction in Abl-I-treated tissues was due to changes in myosin activation, we determined MRLC phosphorylation in smooth muscle tissues treated with the inhibitory peptide. Increases in MRLC phosphorylation in untreated smooth muscle and tissues treated with the control peptide were similar after PE stimulation. Moreover, although treatment with Abl-I peptide attenuated contractile response, the agonist-induced MRLC phosphorylation was not reduced in smooth muscle treated with Abl-I peptide (Fig. 8). The results suggest that Abl is not involved in the regulation of MRLC phosphorylation, which validates our previous findings (1).
Abl is a multifaceted protein that is required for smooth muscle contraction. The mechanism by which Abl regulates force development in smooth muscle is not completely elucidated. In the present study, Abl undergoes phosphorylation at Tyr412 in smooth muscle upon contractile activation. Treatment with the cell-permeable decoy peptide inhibits Abl phosphorylation without affecting its scaffold function. Abl inhibition by the decoy peptide impairs CAS phosphorylation, the dissociation of CAS from cytoskeletal vimentin, actin filament polymerization, and force development in smooth muscle during contractile activation. The results suggest that Abl phosphorylation regulates the release of CAS from the vimentin cytoskeleton, actin polymerization, and contraction by modulating CAS phosphorylation.
In addition to kinase activity, Abl may also serve as a scaffold protein to interact with structural proteins (23, 33). In the present study, Abl binds to the focal adhesion-associated protein vinculin as evidenced by coimmunoprecipitation analysis, consistent with this hypothesis. To further dissect the functional role of Abl, i.e., activation vs. scaffold protein, we characterized the cell-permeable decoy peptide in smooth muscle. Treatment with the decoy peptide inhibited Abl phosphorylation at Tyr412, a key phosphorylation site for Abl activation (1, 30). More importantly, treatment with the decoy peptide did not affect the scaffold function of Abl and the spatial localization of vinculin in smooth muscle cells. Thus, the decoy peptide approach enables us to evaluate the role of Abl phosphorylation in smooth muscle without interfering with its scaffold function. The decoy peptide may compete with endogenous Abl and therefore attenuate the phosphorylation of Abl by members of the Src family and other kinases (6, 29).
The adapter protein CAS is a major member of the CAS family of proteins, which have been thought to serve as docking sites for other proteins in integrin-mediated signaling transduction (5, 21, 32). Recent studies have shown that CAS is an important signaling molecule that participates in the regulation of cytoskeletal signaling and active force development in smooth muscle. CAS undergoes tyrosine phosphorylation in arterial smooth muscle in response to activation with various contractile stimuli (1, 4, 17). Furthermore, the knockdown of CAS by antisense oligonucleotides in smooth muscle depresses active force development by inhibiting actin dynamics, a cellular process that is important for contractile force development in smooth muscle (see below).
CAS is able to bind to the intermediate filament protein vimentin in in vitro biochemical analysis and studies on smooth muscle tissue. Far Western analysis shows that purified vimentin protein interacts with CAS immobilized on the nitrocellulose membrane, suggesting the association of vimentin with CAS in vitro. The fractionation assay also demonstrates the interaction of CAS with cytoskeletal vimentin in smooth muscle tissues. The association of CAS with the vimentin network may hinder its role in cytoskeletal signaling and force development in smooth muscle (15, 21, 22, 32).
There is evidence that contractile activation results in the dissociation of CAS from cytoskeletal vimentin in smooth muscle. Contractile activation leads to the decrease in the ratios of CAS to cytoskeletal vimentin in smooth muscle cells and tissues. Moreover, the ratios of insoluble to soluble CAS are decreased in smooth muscle cells in response to contractile stimulation (15, 32). These results suggest that CAS is dissociated from the vimentin cytoskeleton during contractile activation. Additionally, recent studies demonstrate that the dissociation of CAS from the vimentin cytoskeleton is mediated in part by p21-activated kinase in smooth muscle (21, 22, 31).
Results from the in vitro kinase assay show that CAS is a substrate of Abl tyrosine kinase (1). In this report, Abl inhibition attenuates CAS phosphorylation in smooth muscle during contractile activation. More importantly, the dissociation of CAS from cytoskeletal vimentin in smooth muscle is reduced by Abl inhibition. Thus, it is highly probable that Abl activation coordinates the release of CAS from the vimentin network by modulating CAS phosphorylation. To the best of our knowledge, this is the first evidence to suggest that the dissociation of CAS from the vimentin cytoskeleton is mediated by the Abl-CAS pathway.
Immunofluorescence microscopic analysis reveals that CAS is distributed in the myoplasm of unstimulated smooth muscle cells. CAS relocalizes to the cell periphery in smooth muscle cells in response to contractile activation (32). Because actin assembly transpires in the cortical region of cells (19, 34), it is probable that CAS dissociated from the vimentin cytoskeleton may translocate to the cell border, enhancing cortical actin polymerization and force development in smooth muscle (15, 22, 27, 32). CAS may interact with the adapter protein CrkII, which may facilitate the formation of the multiprotein complex including CrkII, neuronal Wiskott-Aldrich syndrome protein (N-WASP) and the Arp2/3 (actin-related protein) complex and initiate actin polymerization and branching mediated by the Arp2/3 complex (19, 25, 28, 34). In addition, CAS may also regulate the activity of the G-actin-binding protein profilin to promote actin filament polymerization in smooth muscle (21, 27). Finally, Abl inhibition does not attenuate increases in MRLC phosphorylation during contractile stimulation, suggesting that Abl is not involved in the regulation of myosin activation in smooth muscle.
Actin polymerization may enhance smooth muscle force development by the following mechanisms. First, the actin filaments of smooth muscle cells attach to the transmembrane β-integrins, promoting mechanical transduction between the contractile apparatus and extracellular matrix. Cortical actin assembly may strengthen the linkage of actin filaments to integrins and enhance the transmission of contractile force (7, 14, 19, 25, 28, 34). Second, actin assembly has been shown to increase the number of contractile units and the length of actin filaments, providing more and efficient contractile elements for force development (10, 11). Third, newly polymerized filaments may be a part of reorganization processes that allow for rapid adjustment of stiffness and tension (7–9, 25, 28, 34). Fourth, actin filament assembly may participate in the “latch” formation of contractile elements, supporting force maintenance under the condition of lower cross-bridge phosphorylation (9, 20) (Fig. 9).
No conflicts of interest, financial or otherwise, are declared by the authors.
The authors thank Qing-fen Li, Ruping Wang, and Amy M. Spinelli for technical assistance.
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