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EXTRACELLULAR MATRIX, CELL INTERACTIONS

Extracellular matrix-specific focal adhesions in vascular smooth muscle produce mechanically active adhesion sites

¹Dalton Cardiovascular Research Center and ²Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, Missouri

Submitted 2 November 2007 ; accepted in final form 12 May 2008

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
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 (25 µm) coated with fibronectin (FN), collagen type I (CNI), laminin (LN), or vitronectin (VN). ECM-coated beads induced clustering of ₅- 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 (8001,600 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 ₅β₁- and _vβ₃-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.

integrins; myogenic mechanism; atomic force microscopy; mechanobiology; microcirculation

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

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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 310) 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, and 4.7 KCl]. Two glass micropipettes (tip diameter: 4060 µm) filled with Krebs buffer plus 0.5% BSA were used to cannulate the arteriole. The arteriole was set to a length such that lateral bowing did not occur up to intraluminal pressures of 120 mmHg and then pressurized to an in vivo level of 70 mmHg. Vessels typically developed spontaneous tone, resulting in constriction to 50–60% of their passive diameter. Vessel dimensions were measured using a closed-circuit videomicroscopy system coupled to a video caliper (Cardiovascular Research Institute, Texas A&M University Health Science Center). The video caliper allowed lumen diameter to be stored and recorded online as a continuous trace using Powerlab software (AD Instrument). After a spontaneous tone developed in the vessel, the intraluminal pressure was lowered to 30 mmHg, and the vessel was allowed to stabilize to reach a steady state. The intraluminal pressure was then elevated by 15 mmHg in a stepwise manner, and the pressure steps were applied consecutively at 4-min intervals until the final intraluminal pressure reached 105 mmHg. The change of luminal diameter was continuously recorded by a video caliper. Data were analyzed using MS Excel and Graphpad Prism.

Instruments. 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 612 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 (8001,600 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 ₅- and β₃-integrins, specific function-blocking antibodies to ₅- 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).

Reagents. 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-₅-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 HM5-1 and F11 were purchased from Pharmingen.

Viscoelasticity analysis. 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).

View larger version (8K): [in this window] [in a new window]   Fig. 1. A: mechanical circuit that simulates the passive viscoelastic property of vascular smooth muscle cells (VSMCs). The elastic components are represented by the two springs with elasticity of K1 and K0, and the viscous components are represented by the dashpot with viscosity of 0. B: schematic representation of the instant deformation of a VSMC once pulling force (F) was applied to the bead. The envelope represents a single VSMC, and the lines inside represent the cytoskeletal elements. C: typical fitting of the experimental data () with the viscoelastic model (solid line). The dashed line represents the initial instant displacement of the bead, which was determined by the elastic element in the model (K1 + K0). x is the vertical bead displacement and t is time.

where the relaxation time 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. ₀ 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 ₀ 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.

Statistical analysis. Results were compared using paired or unpaired Student's t-tests. Significance was assumed at a P value of 0.05.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
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.

View larger version (23K): [in this window] [in a new window]   Fig. 2. A: trans-illumination image a bead-fused atomic force microscopy (AFM) probe applied to surface of a VSMC. B: force required to detach the bead from the cell surface plotted as a function of contact time. C: displacement of a fibronectin (FN)-coated bead on the VSMC surface in response to step increases of pulling force applied by AFM. The FN-coated bead fused to the AFM cantilever was brought into contact with the VSMC surface to form molecular connections. Top, step increases of pulling force (z-direction) applied onto the bead-cell connection site using AFM. Bottom, corresponding z-direction movements of the FN-coated bead. D: schematics of the VSMC response to a step increase of pulling force. E: myogenic response of an intact isolated rat cremaster skeletal muscle arteriole induced by a stepwise increase of the intraluminal pressure. Arrows depict time points of application of 15 mmHg pressure steps; vessel diameters were normalized by the vessel diameter at 70 mmHg (D/D70), and are shown in a percentage scale. F: vessel responses to a step pressure change from 45 to 60 mmHg. Data represent means ± SE.

View larger version (23K): [in this window] [in a new window]   Fig. 3. A: 5-integrin clustering around the FN-coated bead. B: β3-integrin clustering around the FN-coated bead. C: actin filament clustering around the FN-coated bead. FN-coated fluorescent beads were allowed to adhere to the cell surface for 2 h. Cells were then fixed and stained for integrins and actin filament, respectively, and were imaged using confocal microscopy. Scale bar = 10 µm.

View larger version (20K): [in this window] [in a new window]   Fig. 4. A and B: effect of function-blocking antibodies for 5β1- and vβ3 integrins (HM5-1 and F11, respectively) on the VSMC mechanoresponse. For control, n = 7; for HM5-1 (20 µg/ml), n = 7; for HM5-1 (50 µg/ml), n = 6; for F11 (20 µg/ml), n = 4; and for F11 (50 µg/ml), n = 8. C and D: pulling elasticity and viscosity of control VSMCs and cells treated with HM5-1 or F11. Arrows indicate the time points of force application. Data represent means ± SE.

View larger version (17K): [in this window] [in a new window]   Fig. 5. Effect of cytochalasin D (CytoD; 13.3 µM) and jasplakinolide (JPK; 0.2 µM) on the force-induced VSMC mechanical response and VSMC pulling elasticity. A and C: averaged displacement of FN-coated beads by VSMCs before and after treatment with CytoD (A) or JPK (C). The bead displacement shown was recorded from 10 s before the step increase of pulling force (800 pN) and until 100260 s after force application. A: before CytoD, n = 5; and after CytoD, n = 6. C: before JPK, n = 9; and after JPK, n = 13. B and D: summarized effect of CytoD (B) and JPK (D) on the total displacement of FN-coated beads by VSMCs. The total bead displacement was determined as the change of bead position over 4 min after the application of pulling force. E: effect of CytoD, JPK, and DMSO on the pulling elasticity of VSMCs. Arrows indicate the time points of force application. Data represent means ± SE. *P < 0.05.

View larger version (12K): [in this window] [in a new window]   Fig. 6. Comparison of VSMC mechanical responses, pulling elasticity, and viscosity with beads coated with collagen type I (CNI), laminin (LN), vitronectin (VN), and FN. A: displacement of CNI-, FN-, LN-, VN-, or BSA-coated beads by VSMCs. The bead displacement was recorded from 10 s before a step increase of pulling force and until 160260 s after force application. FN-coated bead, n = 25 and force = 800 pN; CNI-coated bead, n = 19 and force = 800 pN; VN-coated bead, n = 6 and force = 500 pN; LN-coated bead, n = 13 and force = 500600 pN; and BSA-coated bead, n = 2 and force = 800 pN. Arrows indicate the time points of force application. B: pulling elasticity of VSMCs with CNI-, FN-, LN-, or VN-coated beads. C: pulling viscosity of VSMCs with CNI-, FN-, LN-, or VN-coated bead. Data represent means ± SE.

View larger version (32K): [in this window] [in a new window]   Fig. 7. Immunofluorescent microscopy of actin filament structures and 5- and β3-integrin clusterings formed by CNI-coated (A), VN-coated (B), and LN-coated (C) beads. Extracellular matrix protein-coated fluorescent beads were allowed to contact the cell surface for 30 min. Cells were then fixed and either stained for actin filament using Alexa 568-conjugated phalloidin or for 5- and β3-integrins using Cy5. Cells were visualized using confocal microscopy. Arrows depict the position of actin filament clustering and integrin clustering. Scale bars = 10 µm.

View larger version (17K): [in this window] [in a new window]   Fig. 8. A and B: VSMC mechanoresponses to FN-coated (A) and CNI-coated (B) beads in the presence of PP2 or PP3. VSMCs were incubated with PP2 (5 µM) or PP3 (5 µM) for 30 min before the application of AFM pulling force and measurement of bead displacement. For each group, n = 5 and force = 800 pN. C: pulling elasticity of VSMCs with FN- and CNI-coated beads and with PP2 or PP3 treatment. D: pulling viscosity of VSMCs with FN- and CNI-coated beads and with PP2 or PP3 treatment. Data represent means ± SE. E and F: immunofluorescent microscopy of FAK and paxillin clusterings formed by CNI-coated (E) and FN-coated (F) beads. FN- and CNI-coated borosilicate beads were allowed to contact the cell surface for 1 h. Cells were then fixed and stained for FAK and paxillin with Alexa 488 and visualized using confocal microscopy. Scale bars = 10 µm. *P < 0.05.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
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 ₅β₁-integrin, _vβ₃-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 (560 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 1030 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 ₅β₁-integrin, _vβ₃-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 ₅β₁- or _vβ₃-integrins, fluorescence immunolabeling experiments were performed. We observed that ₅β₁- and _vβ₃-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 ₄β₁-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 ₅β₁-integrin, _vβ₃-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.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported by National Heart, Lung, and Blood Institute Grants HL-58960 and HL-062863 (to G. A. Meininger).

	FOOTNOTES

 

Address for reprint requests and other correspondence: G. A. Meininger, Dalton Cardiovascular Research Center, Dept. of Medical Pharmacology and Physiology, 134 Research Park Dr., Univ. of Missouri, Columbia, MO 65211 (e-mail: meiningerg{at}missouri.edu)

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.

¹ Supplemental material for this article can be found at the American Journal of Physiology-Cell Physiology website.

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