Integrin-mediated mechanotransduction in vascular smooth muscle cells (VSMCs) plays an important role in the physiological control of tissue blood flow and vascular resistance. To test whether force applied to specific extracellular matrix (ECM)-integrin interactions could induce myogenic-like mechanical activity at focal adhesion sites, we used atomic force microscopy (AFM) to apply controlled forces to specific ECM adhesion sites on arteriolar VSMCs. The tip of AFM probes were fused with a borosilicate bead (2∼5 μm) coated with fibronectin (FN), collagen type I (CNI), laminin (LN), or vitronectin (VN). ECM-coated beads induced clustering of α5- and β3-integrins and actin filaments at sites of bead-cell contact indicative of focal adhesion formation. Step increases of an upward (z-axis) pulling force (800∼1,600 pN) applied to the bead-cell contact site for FN-specific focal adhesions induced a myogenic-like, force-generating response from the VSMC, resulting in a counteracting downward pull by the cell. This micromechanical event was blocked by cytochalasin D but was enhanced by jasplakinolide. Function-blocking antibodies to α5β1- and αvβ3-integrins also blocked the micromechanical cell event in a concentration-dependent manner. Similar pulling experiments with CNI, VN, or LN failed to induce myogenic-like micromechanical events. Collectively, these results demonstrate that mechanical force applied to integrin-FN adhesion sites induces an actin-dependent, myogenic-like, micromechanical event. Focal adhesions formed by different ECM proteins exhibit different mechanical characteristics, and FN appears of particular relevance in its ability to strongly attach to VSMCs and to induce myogenic-like, force-generating reactions from sites of focal adhesion in response to externally applied forces.
- myogenic mechanism
- atomic force microscopy
mechanotransduction in vascular smooth muscle cells (VSMCs) is a fundamental mechanism underlying the vascular myogenic response (9), in which vascular smooth muscle contracts in response to increased intralumenal pressure or relaxes in response to decreased pressure (11, 21).1 This VSMC behavior is a key mechanism for the establishment of vascular tone and the autoregulation of blood flow (8). It is generally assumed that, in the myogenic response, a perturbation in intravascular pressure changes vascular wall tension/stress, which then triggers the contraction of the VSMC. The VSMC then exerts its contractile force on the extracellular matrix (ECM) it is embedded within. Evidence from isolated, intact arterioles indicates that inhibition of α5β1- and αvβ3-integrins prevents myogenic constriction in response to an acute pressure elevation (23). It has also been demonstrated that ligation of α5β1-, αvβ3-, and α4β1-integrins with ECM proteins modulates Ca2+ conductance through voltage-gated Ca2+ channels (32, 36). Thus, one plausible hypothesis for the myogenic mechanism involves force transmission to sites of integrin attachment with subsequent activation of the actomyosin contractile process (9, 35, 36).
Arising from their complex function in mediating cell attachment and signal transduction, integrins have been more widely recognized to be important for cellular mechanotransduction. Supporting evidence for integrin invovlement in mechanotransduction comes from multiple cell types, for example, endothelial cells (30), fibroblasts (6), osteoblasts (26), neuronal cells (12), and VSMCs (14). Structurally, integrins are a family of transmembrane proteins interposed between ECM proteins and the cytoskeleton (29), thus providing a mechanical connection to the extracellular environment through which mechanical forces are envisioned to be bidirectionally transmitted through focal adhesion sites. Integrin association with the ECM leads to integrin clustering and subsequent association with a number of scaffolding proteins and signaling proteins that are linked as a series of connected elements with the cytoskeleton. Assembly of the scaffolding and signaling proteins and their hierarchical importance within an adhesion site are areas that are incompletely understood. Known processes include, but are not limited to, activation of FAK, recruitment of paxillin, vinculin, and Cas, activation of receptor tyrosine kinases and phosphatases, and cytoskeleton remodeling (see, e.g., Refs. 1, 5, 13, and 29). These processes of protein assembly continue as the focal contact matures into a dynamically regulated focal adhesion (4, 18, 20, 30). As focal adhesions mature, so, apparently, does the ability of the cell to transmit external physical forces and exert tensional forces to the surrounding ECM. Indeed, numerous pieces of evidence support the ability of cells to transmit forces through focal adhesions to their environment (4) and ultimately establish a mechanical balance (2, 20) that involves adaptation at the level of the focal adhesion site.
In this study, we used atomic force microscopy (AFM) to directly measure and apply nanoscale force to ECM-induced focal adhesion sites. The goal of our study was to test the hypothesis that mechanically pulling at a site of integrin-ECM adhesion would induce myogenic-like, micromechanical events from VSMCs. This study helps provide new insights that clarify the role of integrins and the ECM in VSMC mechanosensation and mechanotransduction.
MATERIALS AND METHODS
Cell isolation and cell culture.
Microvascular smooth muscle cells (mVSMCs) were isolated from the first-order feed arteriole (100–150 μm diameter) of Sprague-Dawley rat cremaster skeletal muscles using previously described methods (36). Cells were cultured in DMEM-F-12 supplemented with 10% FBS, 10 mM HEPES (Sigma, St. Louis, MO), 2 mM l-glutamine, 1 mM sodium pyruvate, 100 U/ml penicillin, 100 μg/ml streptomycin, and 0.25 μg/ml amphotericin B. Cells were maintained in 60-mm tissue culture dishes (Falcon, BD Labware, Lincoln, NJ) in a humidified incubator (Heraeus Instruments, Newtown, CT) with 5% CO2 at 37°C. For AFM experiments, cells were cultured in 35-mm tissue culture dishes with a no. 1 glass coverslip bottom (World Precision Instruments, Sarasota, FA). Low-passage cells (passages 3∼10) were used in all experiments. Except for HEPES, all reagents were purchased from Invitrogen (Carlsbad, CA).
Isolated vessel experiments.
Arterioles were isolated from the cremaster skeletal muscle of pentobarbital-anesthetized rats (60 mg/kg) as we have previously described (11). The cremaster muscle was isolated, and a distal segment of the external spermatic artery (first-order arteriole 1A; 120–190 μm inner diameter, passive) was excised and placed in a chamber containing Krebs buffer [composed of (in mM) 112 NaCl, 12 glucose, 25.5 NaHCO3, 9.1 HEPES, 1.2 MgSO4, 2.5 CaCl2, 1.2 KH2PO4
A Bioscope AFM System (model IIIa or IVa, Digital Instruments, Santa Barbara, CA) was mounted on an Axiovert 100 TV inverted microscope (Carl Zeiss, Thornwood, NY). AFM data were collected and analyzed using Nanoscope (Digital Instruments) and Matlab (MathWorks, Natick, MA) software. Immunofluorescently labeled cells were visualized using either a Meridian ULTIMA Z-Laser Confocal Microscope System or a SP2 Confocal/Multiphoton Microscope System (Leica, Bannockburn, IL).
Protein coating of beads.
AFM probes with biotin-labeled borosilicate beads were purchased form Novascan (Ames, IA) with a spring constant of 0.01 N/m. ECM proteins [fibronectin (FN), collagen type I (CNI), laminin (LN), and vitronectin (VN)] were first biotinylated using EZ-Link Sulfo-NHS-LC-Biotin (Pierce, Rockford, IL) following instructions provided by the manufacturer. Briefly, 200 μl of protein (0.5 or 1 mg/ml) were mixed with 6∼12 μl of Sulfo-NHS-LC-Biotin (10 mg/ml) in a microcentrifuge tube (Corning, Corning, NY). The mixture was incubated on ice for 2 h and filtered through a microconcentrator (Millipore, Bedford, MA). Filtered proteins were dissolved in Dulbecco's PBS (DPBS) with 0.1% NaN3 (Sigma-Aldrich) to a concentration of 0.5 mg/ml. AFM probes with biotin-labeled borosilicate beads were incubated with avidin (1 mg/ml, Sigma-Aldrich) for 5 min at room temperature. Probes were then washed five times with DPBS and incubated with biotinylated protein for 6 min at room temperature, followed by another five times of DPBS washing. Polystyrene fluorescent beads (5.46 μm diameter) were purchased from Bangs Laboratory (Fishers, IN) with dragon green (480/520-nm excitation/emission) and streptavidin coating. Beads were spun down and washed three times with DPBS. After being washed, beads were incubated with biotin-conjugated ECM proteins at room temperature for 30 min on a rolling plate. Beads were then washed three times with DPBS and resuspended in 80 μl DPBS. Borosilicate glass beads (5 μm diameter) were purchased from SPI. Beads (1 mg) were mixed with CNI or FN and incubated at room temperature for 3 h. Beads were then spun down, washed three times with DPBS, and resuspended in 80 μl HBSS.
AFM force application and measurement.
To apply pulling forces to the VSMC surface, the AFM was operated in contact mode with the scan size set to 0.1 nm. The experiment was performed at room temperature, and VSMCs were incubated in HBSS. To minimize drift, after the probe was submerged in cell bath, the whole system was thermoequilibrated for 1 h. This equilibration time was longer than that suggested by the manufacturer and was determined to effectively reduce drift experimentally. After thermal equilibration, the protein-coated beads were brought into contact with the cell surface and were kept in a static force neutral position on the cell surface for 25–30 min. The deflection set point was then manually adjusted to apply step increases of pulling force (800∼1,600 pN) to the VSMC at the site of bead contact. The pulling force was generated by the bending of the AFM cantilever and was calculated according to Hooke's law as follows: where F is force (in pN), d is cantilever deflection (in nm), and k is the cantilever spring constant (in pN/nm). Bead displacements were recorded for 4 min in the contact mode. Height data were recorded continuously and analyzed to obtain quantifiable changes in the magnitude of the micromechanical event of the VSMC following the upward pull by AFM.
To examine the role of the actin cytoskeleton, the VSMC micromechanical event was evaluated in the presence of cytochalasin D (13.3 μM, 10 min of incubation), a F-actin depolymerizing agent, or jasplakinolide (0.2 μM, 10 min of incubation), a F-actin stabilizing agent. A paired experimental design was used for these experiments. A control response to pulling with the AFM (800 pN) was obtained before treatment, and a second response was then recorded after the addition of cytochalasin D or jasplakinolide to the cell bath. Vehicle control experiments were also performed.
To evaluate the involvement of α5- and β3-integrins, specific function-blocking antibodies to α5- and β3-integrins were added into the cell bath, and AFM force application and measurements were then performed as described above. Parallel control experiments were performed without antibodies.
To determine the involvement of Src family kinases, cells were incubated with 5 μM PP2 or PP3 (EMD Biosciences, San Diego, CA) for 30 min at room temperature, and AFM force application and measurements were then performed as described above.
Cell staining for immunocytochemistry and confocal microscopy.
Cells were allowed to grow until 50% confluent on glass-bottom tissue culture dishes. FN-coated fluorescent beads were added to the culture dish, and cells were incubated with the beads for 2 h at 37°C. Cells were then washed with DPBS and fixed with 2% paraformaldehyde, followed by the addition of glycine buffer (0.1 mM glycine) for paraformaldehyde quenching. After being washed with PBS, cells were incubated with a primary antibody (1:200 dilution in labeling buffer composed of 150 mM NaCl, 15 mM Na3C6H5O7, 0.05% Triton X-100, and 2% BSA) at 4°C overnight. Cells were then washed six times with cold buffer (composed of 150 mM NaCl, 15 mM Na3C6H5O7, and 0.05% Triton X-100), followed by an incubation with Cy5-conjugated secondary antibody (1:100 dilution in labeling buffer) or Alexa 568-conjugated phalloidin for 1 h at room temperature in a dark environment. Labeled cells were washed six times with cold buffer and imaged on a confocal microscope using excitation wavelengths of 488 nm and 647 nm sequentially. A through-focus image set was collected for each cell with a z-step interval of 0.2 μm. Images were analyzed using ImagePro Plus software (Media Cybernetics, Carlsbad, CA) and Matlab (MathWorks).
Human plasma FN was purchased from Invitrogen, rat plama FN was purchased from EMD Biosciences, and human natural VN was purchased from BD Bioscience. Mouse LN and cytochalasin D were purchased from Sigma. CNI was isolated from the rat tail as previously reported (28). For confocal microscopy, rabbit anti-α5-integrin polyclonal antibody, rabbit anti-rat β3-integrin polyclonal antibody, mouse anti-FAK monoclonal antibody, and rabbit anti-paxillin monoclonal antibody (Millipore) were used as primary antibodies, and Cy5-cojugated goat anti-rabbit IgG antibody (Jackson ImmunoResearch) was used as the secondary antibody. Jasplakinolide, Alexa 647-phalloidin, and Alexa 568-phalloidin were purchased from Molecular Probes. For integrin-blocking experiments, antibodies HMα5-1 and F11 were purchased from Pharmingen.
To analyze the mechanical properties of the collective cellular structures associated with ECM-labeled beads, a mechanical model of viscoelasticity was adopted from Bausch et al. (3) to describe the viscoelastic behavior of cells. The model was modified to fit the experimental condition where the cantilever-attached bead was pulled by an upward vertical force rather than by a tangential force. Briefly, the cellular structure was modeled by a mechanical circuit, known as a Kelvin body, in which the springs represent the elastic elements and the dashpot represents the viscous elements in the cell structure (Fig. 1, A and B).
Upon the application of a step increase of force, the bead displacement could be described by the function below: where the relaxation time τ is given by where F represents the step increase of pulling force, t is time, and x is the vertical bead displacement. k0 and k1 represent the elasticity of the elastic elements of the cell structure (in pN/nm) and are measures of the ability of the solid cellular structure (i.e., the cytoskeleton) to resist elastic deformation caused by extensional stress. γ0 is the viscosity of the viscous elements (pN·nm·s) and is a measure of the ability of the amorphous cellular materials to resist deformation caused by extensional stress. The equations were used to numerically fit the experimental measurements (data acquired within the first 2 s after the application of AFM pulling force, Fig. 1C), and k1 + k0 and γ0 were calculated to represent the pulling elasticity and viscosity of the cell structure under the beads. The pulling elasticity and viscosity reflected the strength of the cell attachment to the bead. High values of elasticity and viscosity indicate a tight (or rigid) cell-bead attachment, and vice versa.
Results were compared using paired or unpaired Student's t-tests. Significance was assumed at a P value of ≤0.05.
VSMC response to forces applied through FN-coated beads.
Figure 2A shows an AFM probe applied to the surface of a VSMC. The AFM probe had a FN-coated bead fused on the tip. The force required to detach the bead from the cell surface increased as a function of contact time (Fig. 2B), suggesting a progressive increase of FN-integrin binding at the adhesion site (data previously published in Ref. 31). In subsequent experiments, the probe with the FN-coated bead was brought into contact with the surface of the VSMC for 25–30 min to allow consistent adhesion to form between the FN-conjugated bead and cell surface integrins. This was then followed with the application of a series of step increases of pulling force (∼1,600 pN) to the bead. Upon each pull, the bead was displaced upward by ∼30–100 nm but remained attached to the cell. Within a few seconds (10 ± 5 s), a cell-generated, myogenic-like force pulled the bead downward toward the cell. The pulling force was maintained constant during the cellular response (Fig. 2C). Figure 2D shows a schematic representation of this micromyogenic event. The response was observed using either human FN (as shown) or rat FN (data not shown). By comparison, in the intact first-order cremaster arteriole, the arteriolar response to a step increase in intraluminal pressure (15 mmHg) was characterized by an initial distention followed by a slowly developing myogenic constriction that reached steady state in 3–5 min (Fig. 2, E and F). Thus, the micromyogenic event observed with the AFM closely parallels observations of the myogenic response in the intact arteriole.
Involvement of integrins and actin in the VSMC response to applied force.
To determine the involvement of specific integrin subtypes and the actin cytoskeleton in the ability of the VSMC to produce this micromyogenic event with the FN-coated bead, immunofluorescence microscopy was used to examine if integrins and actin filaments were associated with FN-coated beads after placement on the cell surface. FN-coated beads were placed on VSMCs and allowed to form adhesions on the cell surface. As shown in Fig. 3, A and B, both α5- and β3-integrins formed ring-shaped clusters around the FN-coated bead, indicating the presence of these integrins and supporting the involvement of both α5β1- and αvβ3-integrins. Actin filaments were also observed to cluster around the FN-coated bead (Fig. 3C), demonstrating the close association of cytoskeletal elements beneath the FN-coated bead.
The involvement of integrins in the micromyogenic event was further tested by blocking α5- and β3-integrins with specific function-blocking antibodies (HMα5-1 and F11, respectively). The presence of either antibody did not abolish the ability of the VSMC to establish an adhesion with the FN-coated bead, but the presence of either antibody significantly inhibited the micromyogenic event (Fig. 4, A and B). The inhibitory effect of the antibodies was concentration dependent (20 or 50 μg/ml). A concentration-dependent increase in initial bead displacement was observed with both inhibitory antibodies, indicating that the pulling elasticity and viscosity of adhesion site and underlying cytoskeletal attachments were reduced in the presence of either antibody (Fig. 4, C and D).
To evaluate the role of the actin cytoskeleton, the micromyogenic event was studied in the presence of cytochalasin D (13.3 μM), an actin filament depolymerizing agent, or jasplakinolide (0.2 μM), an actin filament stabilizing agent. Cytochalasin D totally abolished the VSMC force response, with the FN-coated bead being quickly pulled away from the cell surface, indicating that the adhesive strength of the focal adhesion with the FN bead was significantly reduced (Fig. 5, A and B). In contrast, after the application of jasplakinolide (0.2 μM), the micromyogenic event was augmented, resulting in more rapidly developing tension (0.59 ± 0.11 vs. 0.34 ± 0.18 nm/s; Fig. 5, C and D). DMSO was applied as a vehicle control for jasplakinolide and cytochalasin D and had no effect on micromyogenic event (data not shown). As anticipated, cytochalasin D decreased the cell elasticity compared with jasplakinolide and the DMSO control, suggesting an effect on the intracellular cytoskeletal elements associated with the FN bead (Fig. 5E). Collectively, these results provide evidence for the involvement of the actin cytoskeleton in the micromyogenic event that occurs with the FN-coated bead.
VSMC response to force applied through CNI-, VN-, or LN-coated beads.
To determine whether other ECM proteins induced a similar micromyogenic event, beads coated with CNI, VN, or LN were applied to the cell surface. Beads coated with BSA alone were applied to the cell surface as a control for the ECM proteins. As shown in Fig. 6 a, CNI, VN, or LN did not induce the micromyogenic event from VSMCs. Force applied to sites of adhesion with CNI-coated beads exhibited strong and stable attachment that resisted the direction of pull by AFM. In comparison, beads coated with VN or LN were gradually but continuously pulled away from the cell surface using similar pulling force to that used for FN and CNI. This indicates a much weaker form of adhesion with these ECM proteins under the conditions of our experiments. In contrast to ECM-coated beads, BSA-coated beads failed to show evidence of adhesion and were immediately lifted off the cell and out of the range of measurement. A mechanical analysis of the viscoelastic properties of the cellular structures associated with the beads indicated that both the pulling elasticity and viscosity were greater with CNI- or FN-coated beads than with VN- or LN-coated bead (Fig. 6, B and C). When VSMCs with ECM-coated beads were investigated using qualitative immunocytochemistry, we observed that all four ECM proteins studied, but not BSA (data not shown), were associated with the actin cytoskeleton after 30 min of bead contact (Fig. 7). In addition, beads coated with CNI, VN, and LN were also found to induce clustering of α5- and β3-integrins (Fig. 7). BSA did not induce any observable integrin clustering (data not shown).
Since adhesion to FN and collagen exhibited similar adhesive properties as quantified by elasticity and viscosity, we further investigated the selective recruitment of FAK and/or paxillin to these adhesion sites. Selective recruitment of one or both of these focal adhesion proteins could provide insight into the ability of FN to elicit the micromyogenic event. However, both FAK and paxillin were found clustered near the FN- and CNI-coated beads (Fig. 8, E and F).
To further evaluate a possible mechanism, we examined whether tyrosine phosphorylation events mediated by Src were involved. Cells were treated with PP2, a specific inhibitor, to block Src family kinases. PP2 significantly reduced the elasticity and viscosity of cell attachment to both FN- and CNI-coated beads compared with the PP3 control group. Although the FN-coated bead continued to maintain its attachment to the cell, PP2 abolished the micromyogenic-like response to pulling through the FN-coated bead. PP2 significantly impaired the ability of the CNI-coated bead to withstand the pulling force, and the bead was gradually pulled away from the cell under constant force (Fig. 8, A–D). These results suggested that in mVSMCs, the Src-mediated signaling pathway plays an important regulatory role in the mechanisms that strengthen adhesive attachment for FN and CN, and it is important for the development of the micromyogenic event.
The goal of this study was to determine if nanoNewton scale forces applied through specific ECM-integrin focal adhesion sites could induce myogenic-like micromechanical events from single mVSMCs. Our results indicate that forces applied to an FN-induced focal adhesion site induced mVSMCs to respond with a micromyogenic event. The micromyogenic event was dependent on interactions with α5β1-integrin, αvβ3-integrin, the actin cytoskeleton, and cSrc activity. To our knowledge, this is the first study using AFM technology to quantitatively measure the mechanical responses to applied force at single focal adhesion sites formed with specific ECM proteins. In contrast to FN, force applied to focal adhesion sites induced with CNI, VN, or LN were not able to elicit the micromyogenic event, suggesting that FN could be of particular relevance to the mechanosensing and transducing pathways in mVSMCs.
In this study, we estimated the scales of time and pulling force to mimic those approximated to occur at the single cell level in an intact arteriole and that result in a myogenic response. This was done to strengthen our ability to correlate observations at the single cell level with the myogenic behavior of intact arteriole. As shown in Fig. 2E, during the myogenic response, the isolated arteriole constricted over a 3- to 5-min period following an initial distention. Similarly, the micromyogenic event in mVSMCs occurred over a similar 3- to 5-min interval. In the intact arteriole, the myogenic response to a step increase of pressure of 15 mmHg converts to an extensional stress of 2,000 pN/μm2 on the vessel wall. Assuming that mVSMCs are the major force-bearing components in the vessel wall, we estimated that mVSMC-ECM interactions could experience forces close to that magnitude. For our experiments, we selected values for pulling from 800 to 1,600 pN, with most experiments conducted at 800 pN. With the use of 2- to 5-μm-diameter beads, the applied forces experienced at a single focal adhesion site would be ∼30–200 pN/μm2. The absolute rate of tension development by the single mVSMC is ∼0.34 nm/s (rate of downward bead displacement on cell), lower than the predicted rate of cell length change in the intact vessel wall (∼41.6 nm/s). However, the estimated rates of cell deformation, obtained by normalization with their respective dimension scale, fall into the same magnitude (1.4 vs. 4.2 times 10−5·s−1). These quantitative estimates suggest that the micromyogenic behavior that we observed at single focal adhesion sites in mVSMCs closely parallel the estimated mechanical events envisioned to occur in the intact arteriole.
The observation that mVSMCs could generate a micromyogenic force in response to force applied to the adhesion with FN is consistent with similar mechanical observations in other cell types. For example, in fibroblasts, Choquet et al. (6) reported that cells generated sufficient force to pull a bead coated with FN7-10 (a major integrin-binding motif) against a trapping force (5∼60 pN) applied through laser tweezers. This suggested that the response involved integrin-mediated mechanosensing and cytoskeleton strengthening. In another study, Heidemann et al. (15) attached a LN-coated microglass needle to fibroblasts and demonstrated that the cell could contract against a force of 10∼30 nN. In contrast to these observations, mVSMCs did not respond to force applied through LN. This could be related to differences in cell type. Integrin-mediated cellular force has also been demonstrated in processes such as bending of collagen fibrils by chondrocytes (22), contraction of fibrin clots by smooth muscle (37), and contraction of collagen gels by embryo cells (17). Our results also provide further quantitative biomechanical evidence suggesting that integrins are important sites for mechanosensing and force transmission in mVSMCs. In addition, our results point to FN as an important ECM protein that is linked to micromyogenic behavior in mVSMCs.
Fluorescent imaging of mVSMCs immunolabeled for α5β1-integrin, αvβ3-integrin, and actin filaments showed accumulation of these proteins beneath the FN-coated beads (Fig. 4). In addition, FAK and paxillin were also found around the FN-coated bead (Fig. 8). The association of these proteins at the point of contact with the FN-coated bead indicated that mVSMCs formed a focal adhesion-like structure around the FN-coated bead. It was also observed in our experiments that the force required to completely detach the FN-coated bead from the VSMC increased with increasing duration of contact time. This indicated that a progressive biomechanical process was contributing to the formation of the adhesion complex between FN and the cell.
Disruption of the actin cytoskeleton by cytochalasin D abolished the cellular force response and reduced the mechanical elasticity of the focal adhesion site in response to pulling. This is consistent with the notion that the actin cytoskeleton is the major force-bearing elastic structure in the cell (19, 27, 33). Interestingly, blockade of actin depolymerization with jasplakinolide acutely enhanced the micromyogenic response. Supportive observations have been recently described by Zhang et al. (38). They showed that jasplakinolide enhanced the mechanical gating of nonselective stretch-inhibited cation channels during osmosensory transduction in neuron cells, whereas cytochalasin-D reduced it (38). In addition, Cipolla et al. (7) have shown that actin polymerization is strongly enhanced during the myogenic response in intact isolated cerebral arterioles. These authors proposed that the regulation of actin polymerization in VSMCs was an important mechanism for the regulation of the myogenic response (7). Collectively, these results support the downstream involvement of the actin cytoskeleton in the micromyogenic event observed in our study.
In our study, FN appeared to be unique in its ability to form an adhesion site on mVSMCs that was mechanically active and capable of generating force. In this regard, the application of force to attachment sites induced with CNI, VN, and LN did not result in a myogenic-like response by mVSMCs. The reason for this observation may include differences in integrins present, differences in focal adhesion scaffolding, and/or differences in cytoskeletal and/or signaling proteins present in the focal adhesion. Analysis of the viscoelastic properties of the attachment sites induced by ECM proteins used in this study (Fig. 6, B and C) revealed that attachment sites associated with CN- and FN-coated beads were stiffer and more viscous than attachment sites associated with LN- and VN-coated beads. Consistent with our observations, Hocking et al. (17) have shown that FN enhanced the contraction of smooth muscle cells embedded in collagen gels, whereas VN and LN did not. Furthermore, it has also been shown that VSMC gene expression induced by mechanical strain is different on FN substrate compared with VN and LN substrates (34), thus supporting the notion that the biomechanical composition and functional properties of mVSMC focal adhesion sites are different for different ECM proteins.
To determine if the observed differences in focal adhesion mechanical properties and the unique ability of FN to induce a micromyogenic event were due to the unique presence of α5β1- or αvβ3-integrins, fluorescence immunolabeling experiments were performed. We observed that α5β1- and αvβ3-integrins were not selective for FN but were also recruited into adhesion sites induced by CNI, VN, and LN (Fig. 7). Thus, the qualitative presence of these two integrin subtypes does not explain the differences in the mechanical properties or the molecular basis for the micromyogenic event. Additional immunlabeling experiments were performed for FAK and paxillin. In these experiments, we focused our evaluation on the focal adhesion sites with FN and CNI, since they exhibited the strongest adhesion, but only FN induced the micromyogenic event. Our results indicated that both FAK and paxillin were present at the adhesion sites for FN and CNI and thus qualitatively do not provide insight into the mechanism for the micromyogenic event. Further studies will be necessary to determine the molecular and signaling pathways that underlie the micromyogenic event and the differences in the mechanical properties of attachment. One obvious possibility is that other integrins or adhesion molecules are present at these adhesion sites or that the outside-in signaling process is very different. Recent reports have suggested that FN interactions with cell surface heparan sulfate proteoglycans or α4β1-integrin are involved in the formation of focal adhesions and stress fibers (10, 16, 24, 25). Again, further studies will need to address the possible involvement of additional receptors and/or unique recruitment of biochemical signaling molecules.
We tested the possibility that cSrc might be involved through a tyrosine phosporylation process. PP2 treatment reduced the elasticity and viscosity of both FN- and CNI-coated bead adhesions to the same extent but eliminated the micromyogenic event observed with FN. Despite the loss of the myogenic-like response, mVSMCs were able to maintain a mechanically stable adhesion with the bead during constant force loading. In contrast, CNI was not able to sustain stable adhesions with mVSMCs under constant load. Collectively, these results further indicate that signaling pathways involving cSrc are playing an important role in the micromyogenic event and are required for stabilization of the adhesion site. PP3 was used as a negative control to PP2, but it is also an inhibitor of EGF receptor (EGFR) kinase activity. We observed that it reduced the viscosity and elasticity of CNI adhesions and also attenuated the VSMC myogenic-like response to FN-coated beads, suggesting that EGFR-related signaling pathways may also be involved in VSMC mechanotransdcution. Future studies will be necessary to address the cross-talk between these signaling pathways in VSMCs and identify the relevant phosporylation targets. Work by Wu et al. (36) has clearly demonstrated that FN-induced cSrc activation is linked to phosphoryation of the L-type Ca2+ channel, providing one possibly important link to the micromyogenic event observed in our study.
In summary, our results indicate that mechanical force applied to a focal adhesion induced with FN can elicit a micromyogenic event in mVSMCs in response to the application of a constant force to the focal adhesion site. This myogenic-like response is not observed with CNI, VN, or LN, suggesting ECM specificity. The qualitative presence of α5β1-integrin, αvβ3-integrin, actin, paxillin, and FAK at the adhesion sites with these ECM proteins is not sufficient to explain the FN-induced micromyogenic effect. However, a role for cSrc appears to be important. Further work will be necessary to delineate the specific cell signaling pathways and critical focal adhesion proteins to understand the mechanism of this myogenic-like response. Understanding this micromyogenic event may provide significant new insights into the mechanism of the vascular myogenic response.
This work was supported by National Heart, Lung, and Blood Institute Grants HL-58960 and HL-062863 (to G. A. Meininger).
↵1 Supplemental material for this article can be found at the American Journal of Physiology-Cell Physiology website.
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.
- Copyright © 2008 the American Physiological Society