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Am J Physiol Cell Physiol 295: C160-C172, 2008. First published May 14, 2008; doi:10.1152/ajpcell.00014.2008
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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS

Role for stress fiber contraction in surface tension development and stretch-activated channel regulation in C2C12 myoblasts

Francesca Sbrana,1,* Chiara Sassoli,2,* Elisabetta Meacci,3 Daniele Nosi,2 Roberta Squecco,4 Ferdinando Paternostro,2 Bruno Tiribilli,5 Sandra Zecchi-Orlandini,2 Fabio Francini,4 and Lucia Formigli2

1Department of Physics, Centro Studi Dinamiche Complesse, University of Florence, Florence; 2Department of Anatomy, Histology, Forensic Medicine, University of Florence, Florence; 3Department of Biochemical Sciences, University of Florence, Florence; 4Department of Physiological Sciences, University of Florence, Florence; and 5ISC-CNR, Institute for Complex Systems, Sesto Fiorentino, Florence, Italy

Submitted 11 January 2008 ; accepted in final form 11 May 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Membrane-cytoskeleton interaction regulates transmembrane currents through stretch-activated channels (SACs); however, the mechanisms involved have not been tested in living cells. We combined atomic force microscopy, confocal immunofluorescence, and patch-clamp analysis to show that stress fibers (SFs) in C2C12 myoblasts behave as cables that, tensed by myosin II motor, activate SACs by modifying the topography and the viscoelastic (Young's modulus and hysteresis) and electrical passive (membrane capacitance, Cm) properties of the cell surface. Stimulation with sphingosine 1-phosphate to elicit SF formation, the inhibition of Rho-dependent SF formation by Y-27632 and of myosin II-driven SF contraction by blebbistatin, showed that not SF polymerization alone but the generation of tensional forces by SF contraction were involved in the stiffness response of the cell surface. Notably, this event was associated with a significant reduction in the amplitude of the cytoskeleton-mediated corrugations in the cell surface topography, suggesting a contribution of SF contraction to plasma membrane stretching. Moreover, Cm, used as an index of cell surface area, showed a linear inverse relationship with cell stiffness, indicating participation of the actin cytoskeleton in plasma membrane remodeling and the ability of SF formation to cause internalization of plasma membrane patches to reduce Cm and increase membrane tension. SF contraction also increased hysteresis. Together, these data provide the first experimental evidence for a crucial role of SF contraction in SAC activation. The related changes in cell viscosity may prevent SAC from abnormal activation.

actin remodeling; atomic force microscopy, Young's modulus; membrane capacitance; hysteresis

STRETCH-ACTIVATED CHANNELS (SACs) are voltage-independent nonselective ion channels localized on the plasma membrane, where they play an important role as mechanoreceptors (33). These channels, in fact, are gated by tension developed within the lipid bilayer (25) and transduce the mechanical stimulus into increased cation current and Ca2+ influx, thus converting electrical signals into biochemical events involved in the coordination of numerous cellular processes, ranging from cell volume regulation, membrane potential control, and muscle cell contraction to regulation of gene expression and cell differentiation (8, 9, 28). The analysis of mechanotransduction has been focused on the identification of critical molecules and cellular components, such as integrins, focal adhesions, cadherins, gap junctions, and cytoskeleton, which, modulating cell-cell and cell-matrix interactions, contribute to the mechanosensitivity and promote SAC opening (18). However, there is increasing evidence suggesting that actin cytoskeleton reorganization may also represent an important mechanism of SAC regulation in the structural context of the single cell. This assumption gains support from recent studies suggesting a close structural interaction between cytoskeleton and SACs. Indeed, single molecule force spectroscopy studies have shown that individual peptide domains within proteins found in the actin cytoskeleton unfold when SACs are mechanically extended (19, 31), and recent evidence has demonstrated that members of the transient receptor potential (TRP) family of cation channels, likely molecular candidate for SACs (25, 34, 38), are physically linked with ankyirin and dystrophin (30, 40). However, the functional impact of actin on SAC activity is rather complex and seems to depend mostly on the different status of the microfilament system in specialized cells. In fact, actin cytoskeletal remodeling after the application of mechanical stimuli or the loss of microfilaments by treatment with actin disrupter agents, such as cytochalasins or latrunculin, could modulate the activity of SACs and increase stretch-induced calcium transients (23, 44). On the other hand, actin cytoskeletal disassembly has been shown to cause a decrease in single current and conductance of SACs in myeloid leukemia cells and ventricular myocytes (20, 37), suggesting that the organization of the cortical actin may be determinant in positively modulating channel function in these cells. To further complicate this scenario, we have demonstrated that actin polymerization and stress fiber (SF) formation in response to stimulation with sphingosine 1-phosphate (S1P), a bioactive lipid, is also able to modify the intrinsic conductive properties of SACs and increase the channel ion current (9, 10), thus contributing to generate the idea that a well-organized cytoskeleton may also exert tension forces at the plasma membrane and provoke SAC opening.

On the basis of the aforementioned, in the present study we combined atomic force microscopy (AFM) and electrophysiological patch-clamp analysis to gain further insights into the relationship between actin cytoskeletal features and mechanosensitivity (SAC activity) in living C2C12 myoblasts, with the potential to give information in a noninvasive way on the role of cytoskeleton in the regulation of cell surface topography and plasma membrane stretching (15). To further stress this point, we examined the effects of SF formation and myosin II-driven SF contraction on cell surface area and plasma membrane remodeling. The identification of the basic mechanisms regulating SAC activation in myoblastic cells may have considerable implications in the understanding of the role played by these channels in skeletal muscle biology and disease.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell Cultures and Treatments Murine C2C12 skeletal myoblasts obtained from American Type Culture Collection (Manassas, VA) were cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum, penicillin (100 U/ml), and streptomycin (100 µg/ml) (Sigma, Milan, Italy) and maintained at 37°C in a humidified atmosphere of 5% CO2. For AFM investigation, cells were grown on glass coverslip mounted on pierced culture 35-mm dishes, which were clamped to a petri dish holder on the stage of the microscope. To induce SF formation, cells were stimulated with S1P for 30 min (1 µM, Calbiochem, San Diego, CA). In parallel experiments, the cells were treated for 30 min with the following reagents in the presence of S1P: Y-27632 (10 µM, Calbiochem), used to inhibit endogenous Rho kinase activation and SF formation; and blebbistatin (10 µM, Sigma), to inhibit myosin II ATPase activity.

Confocal Immunofluorescence Cells grown on glass coverslips were fixed in 0.5% buffered paraformaldehyde for 10 min at room temperature, permeabilized with cold acetone for 3 min, washed in phosphate-buffered saline (PBS) and incubated with tetramethylrhodamine isothiocyanate (TRITC)-labeled phalloidin (1:100; Sigma) to detect F-actin filaments. In some experiments, the cells were also immunostained to reveal focal adhesions (FAs) or TRPC1, a structural component of SACs. To this purpose, the cells were blocked with 0.5% bovine serum albumin (Sigma) and 3% glycerol in PBS for 30 min and were incubated with either 1:100 dilution of rabbit polyclonal anti-focal adhesion kinase (FAK; Santa Cruz, Milan, Italy) or 1:80 dilution of rabbit polyclonal anti-TRPC1 (Santa Cruz) antibodies, followed by Alexa488-conjugated IgG (1:100; Molecular Probes, Eugene, OR). Negative controls were carried out by replacing the primary antibodies with nonimmune mouse serum; cross-reactivity of the secondary antibodies was tested in control experiments in which primary antibodies were omitted. Plasma membrane localization of TRPC1 was also determined. To this purpose, living cells were incubated with TRITC-conjugated wheat germ agglutinin (TRITC-WGA) (1:250, Molecular Probes) for 10 min at room temperature to label the plasma membrane before being fixed and immunostained for the expression of TRPC1. The coverslips containing the labeled cells were then mounted with an antifade mounting medium (Biomeda gel mount, Electron Microscopy Sciences, Foster City, CA) and observed under a confocal laser-scanning microscope (MRC 1024 ES, Bio-Rad, Hercules, CA) equipped with a krypton/argon laser source. The argon (488 nm) and krypton (568 nm) laser lines were used to excite the cells, and the emitted fluorescence signals were collected with a Nikon Plan Apo x60/1.4 numerical aperture (NA) oil immersion objective. Series of optical sections (512 x 512 pixels each; pixel size 200 nm x 200 nm or 150 nm x 150 nm) were then taken through the depth of the cells at intervals of 0.4 µm. Images were then z-projected onto a single max intensity projection image. Confocal images were deconvolved using ImageJ 3D deconvolution software [National Institutes of Health (NIH)]. Densitometric analysis of radial and longitudinal SFs was performed measuring the average fluorescence intensity on regions of interest (ROIs; 6-µm2 area) in single focal plane images of three-dimensional confocal stacks, using ImageJ (NIH) software. At least 4 ROIs in 50 different cells were analyzed in each experiment (3 preparations/experiment), and optical density (means ± SE) was then calculated. In the colocalization experiments, a confocal Leica TCS SP5 microscope (Leica Microsystems, Mannheim, Germany) equipped with a HeNe/Ar laser source was used, and the observations were performed using a Leica Plan Apo x63/1.40 NA oil immersion objective. To minimize cross talk between fluorochromes, the fluorescent signals were acquired sequentially at excitation wavelengths of 488 nm and 543 nm. Series of optical sections (1,024 x 1,024 pixels each; pixel size 200 nm x 200 nm) were taken through the depth of the cells at intervals of 0.4 µm. Quantitative assessment of colocalization between TRPC1 and WGA fluorescence signals was performed by calculating the overlap coefficient (ranging from 0, minimum colocalization degree, to 1, maximum colocalization degree), using the Leica Application Suite software. At least 50 different cells were analyzed in each experiment (three preparations/experiment), and the overlap coefficient (means ± SE) was then calculated. Statistical analysis of differences was performed using one-way ANOVA test. P < 0.05 was considered significant. Calculation were performed using GraphPad Prism software program (GraphPad, San Diego, CA).

Atomic Force Microscopy Measurements and Imaging Force-indentation measurements. A PicoSPM-scanning probe microscope (Molecular Imaging, Tempe, AZ) was used to evaluate the elastic and viscous properties of single living cells. To this purpose, modified spherical AFM tips were used as microindenters to minimize the damaging effects of tips and to perform quantitative measurements of the elasticity. These tips were prepared by gluing a microglass sphere of 22 µm in diameter (Whitehouse-Scientific, Chester, United Kingdom) at the end of a silicon triangular cantilever (MikroMasch, Tallinn, Estonia) with a calibrated spring constant of 0.109 N/m. Load/unloading cycles were performed, where the AFM spherical tip was approached to the cell, pushed to indent (maximal applied force 20 nN), and then retracted. The corresponding plots, so-called force-distance curves, were collected. Deflection signal was expressed as nanometers; calibration was obtained for each experiment from a force-distance curve on a glass coverslip with cell culture medium. The time to record a force curve cycle was ~0.6 s. Multiple force-distance curves were systematically collected in six different areas of each cell, in five different cells for each treatment. The measurements were accomplished by avoiding epinuclear regions, to exclude the contribution of the nuclear stiffness, as well as near the edge of the cells to avoid the possible contribution of the glass. To calculate elasticity (Young's modulus) and viscosity (normalized hysteresis), only the portion after contact of the approaching curve was considered and converted into force-indentation (F-{delta}) curves. Young's modulus (E) was estimated using the Hertz model extended by Sneddon (36) that describes the indentation of a rigid tip onto a semi-infinite, isotropic homogeneous elastic surface. For hard spherical tips with radius R, the force-indentation relation (42) was given by F({delta}) = 4R1/2E{delta}3/2/3(1 – {nu}2), where F is the loading force, E is Young's modulus, {nu} is the Poisson ratio, and {delta} is the indentation. The Poisson ratio was assumed to be 0.5, as previously reported for living cells (35), and the indentation was calculated as the difference between the scanner position and the cantilever deflection. Young's modulus was estimated in the first 400 nm on the approaching indentation curve; this depth allowed us to analyze the mechanical properties of the plasma membrane and subcortical cytoskeleton according to previous reports (7, 21). Hysteresis was evaluated by estimating the energy dissipated into the cell from the indentation of the AFM tip (45). The areas under the curve for extension (Aext) and retraction were calculated integrating over the indentation depth and the difference between these areas (hysteresis) normalized with respect to the input energy Aext. The measurements were made at a velocity of about 1 µm/s. Since it has been reported that starting from a velocity of 0.25 µm/s the viscous energy accounts for >70% of the hysteresis, we assumed that the data obtained were indicative of cell viscosity (26). The data and errors followed a Gaussian distribution. Young's modulus and normalized hysteresis data are expressed as means ± SE. Statistical analysis was performed by using ANOVA test (P < 0.05).

AFM imaging. AFM imaging was performed on living cells, using the PicoSPM-scanning probe microscope equipped with a Pulsed Force Mode (PFM) external unit (WITec, Wissenshaftliche Instrumente und Technologie, Ulm, Germany). Simultaneous images of the topography (heights), deflection, and local stiffness of cells were acquired. The deflection images were related to the gradient of the height and showed the fine details of the plasma membrane and subcortical (cytoskeletal) structures. A sinusoidal z modulation was applied by PFM to the z-axis scanner, and the system worked in a nonresonant intermediate-contact mode. The measurements were performed at a frequency of 104 Hz, while the modulation amplitude was adjusted in such a way that the AFM tip, briefly but periodically, touched the sample surface. The stiffness map was recorded as the difference between a chosen trigger point and the maximum amplitude in the force versus time curve, acquired point by point. Rectangular Ultrasharp silicon cantilevers (NT-MDT; Moscow, Russia) with a nominal spring constant of K = 0.10 N/m were used. Images (256 x 256 pixels) were acquired using a 30-µm scanner at a scan rate of 0.2 line/s. The AFM was combined with an inverted optical microscope (TE300 Eclipse, Nikon) to guide the movement of AFM probes directly above the designated cells. All experiments were performed in cell culture medium at room temperature. In the stiffness maps, the values were represented by pseudo-color scales. The height and stiffness profiles were evaluated along selected lines depicted in the images using WSxM Nanotech software (16). The lines were selected in peripheral regions of the cells to avoid epinuclear areas.

Electrophysiological Analyses Whole cell path clamp. The electrophysiological properties of C2C12 cells were analyzed by single microelectrode whole cell patch clamp in voltage-clamp conditions, as previously described (9). Briefly, the patch pipettes were filled with a solution containing 150 mM CsBr, 5 mM MgCl2, 10 mM EGTA, and 10 mM HEPES, which was filtered through 0.22-µm pores. pH was titrated to 7.4 with NaOH and to 7.2 with TEA-OH for bath and pipette solution, respectively. To block K+ channels, transmembrane currents were recorded in K+-free bath solution (14) containing 122.5 mM NaCl, 2 mM CaCl2, 20 mM TEA-OH, and 10 mM HEPES.

The cell was held at –60 mV, and step pulses of 10 mV and 100 ms of duration, from –80 to 0 mV, were applied every 10 s. Electrode capacitance was compensated before disrupting the patch. Access resistance (Ra) was not compensated for monitoring membrane area. The area beneath the capacitive transient and the time constant of the transient's decay ({tau}) were used to calculate the cell linear capacitance (Cm) and Ra from {tau} = RaCm. The measurement of membrane resistance (Rm) was corrected for Ra and calculated from the steady-state membrane current (Im) using the following relation: Rm = ({Delta}V – ImRa)/Im, where {Delta}V is the command voltage step amplitude. Since the exponential rise of the voltage step causes underestimation of Cm, particularly when leak conductance increases greatly, Cm was corrected using the relation Cm = {Delta}Q(Rm + Ra)/Rm{Delta}V, where {Delta}Q is the sum of the time integral of the current transient and Im{tau} elicited by each voltage step (10). Cm, as an index of the cell surface area (assuming that membrane-specific capacitance is constant at 1 µF/cm2), was used to compare the SAC currents in cells of different shape and surface area [transmembrane conductance (Gm) vs. Cm relation]. To ensure that the registered transmembrane currents occurred through putative SACs, parallel experiments were performed using gadolinium chloride (GdCl3; 50 µM; Sigma), a commonly used SAC blocker, which was added to control and S1P-stimulated myoblasts 3 min prior to the electrophysiological analysis. The SAC conductance was evaluated by subtracting from Gm the leak conductance (Gm,leak) evaluated as the residual current in the presence of GdCl3. All experiments were performed at room temperature (20–23°C). The relations between Gm and Cm, Gm and Young's modulus (E), Cm and Young's modulus (E), and Cm and normalized hysteresis were evaluated by linear and nonlinear best fit. Data fitting used a nonlinear curve fitting procedure based on the Marquardt-Levenberg algorithm (Sigmaplot 4 and Table Curve 3.10, Jandel Scientific; and Clampfit 6.02, Axon Instruments). Linear or a sum of exponential functions consisting of a different number of terms was fitted to the data. The best fit was chosen by means of a test based on the value of the likelihood ratio statistic, with the same formalism previously reported (12). The improvement of the fit was evaluated by {chi}2-statistics. The number of degrees of freedom by using linear or one or two exponential terms was two or three or six. We used {chi}2-statistics with one degree of freedom (equal to the difference of number of parameters when one exponential term or a linear term was used, and three degrees of freedom in the case of two and one exponential terms used). The improvement of the fit was statistically significant (P < 0.05) if the {chi}2-statistics exceeded 7.8. The data and errors followed a Gaussian distribution and are expressed as means ± SE. One-way ANOVA with repeated measures was used for multiple comparisons, and {alpha}-value at P < 0.05 was considered significant.

SAC currents after mechanical stretching. To further demonstrate the presence of SACs, we stretched the cell membrane using two patch microelectrodes, according to the method described by Zhang et al. (47). Briefly, one microelectrode was positioned, by a hydraulic micromanipulator, at the center of the cell and was used for electrophysiological measurements. The other microelectrode, placed 10–20 µm far away from the former, was moved by another hydraulic micromanipulator to apply a stretch to the cell via longitudinal displacement. The stretching was applied in a region far from the nuclear zone. The extent of stretch was expressed as the percentage change in cell length (L) relative to the original length: L = (Lstretch – Loriginal)/Loriginal x 100. The used stretch extent was 10%.

Ca2+ Imaging After Mechanical Stimulation Ca2+ entry through SACs was estimated from the changes in total fluorescence that occurred when SACs opened and Ca2+ bound to Fluo3-AM, used as a fluorescent Ca2+ indicator (Molecular Probes). The channels were mechanically activated using the rectangular tip of an AFM (PicoSPM-scanning probe microscope), as previously reported (10). For the observations, the AFM was mounted on top of an inverted optical microscope (Nikon) equipped with a digital camera (Nikon D100). The observations were performed during and soon after the mechanical stimulation. Images were filtered with fast Fourier transform-based band-pass filter to remove structures down to 80 µm and up to 2 µm, corresponding to diffused light halos and digital camera noise, respectively. The intracellular Ca2+ levels were expressed as relative fluorescence [{Delta}F/Fb: ratio of fluorescence difference, peak-basal (Fp – Fb), to basal (Fb) values], as previously reported (41).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effects of Stress Fibers on C2C12 Cell Surface Stiffening and Topography Since previous reports and data from our laboratory have suggested the existence of a structural and functional relationship between cytoskeleton and SAC activation (10, 19, 31), we first investigated whether SF formation was capable of modifying the mechanical and topographic properties of the cell surface and then correlated these features with the development of cell surface tension and SAC activation. SACs are, in fact, transmembrane channels that are activated by membrane stretch (18, 25). To this purpose, living C2C12 myoblasts were stimulated with S1P to obtain formation of SFs and were examined by confocal microscopy to visualize actin cytoskeleton and by AFM microscopy to characterize the surface morphology and the mechanical properties (i.e., stiffness and viscosity) of the cells.

Confocal immunofluorescence of unstimulated control C2C12 cells showed the presence of two types of actin distribution (Fig. 1A) : radial actin bundles, connecting the perinuclear zone with the cell surface (radial SFs) and cortical actin bundles (longitudinal SFs) running parallel underneath the plasma membrane. Double staining of these cells with specific antibodies against FAK (Fig. 1B) allowed us to show that these filaments terminated in focal adhesion FAK-containing plaques that connected radial and longitudinal SFs to the plasma membrane, further stressing the existence of a structural coupling between actin filaments and the cell surface in myoblastic C2C12 cells. Stimulation with 1 µM S1P for 30 min greatly increased radial and longitudinal SF and FA densities (Fig. 1, C, D, and G).


Figure 1
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  		Fig. 1. Confocal fluorescence analysis of cytoskeleton organization of skeletal myoblasts. C2C12 cells were grown on glass coverslips, fixed with 0.5% paraformaldehyde in PBS, and stained with tetramethylrhodamine isothiocyanate (TRITC)-labeled phalloidin to detect F-actin (red); in some experiments, the cells were further incubated with rabbit polyclonal anti-focal adhesion kinase antibody followed by Alexa488-conjugated IgG to reveal focal adhesion (FA) plaques (green). A and B: control, unstimulated C2C12 cells. C and D: cells stimulated with 1 µM sphingosine 1-phosphate (S1P) for 30 min. E and F: cells treated with either 10 µM Y-27632 (E) or 10 µM blebbistatin (F) in the presence of S1P. The fluorescence distribution and intensity of F-actin are shown in the reported pseudo-color scale. In A and C, two types of stress fibers (SFs), radial (underscored by continuous arrows) and longitudinal (underscored by dotted arrows), aligned prevalently underneath the plasma membrane are visible. S1P elicits formation of dense SFs and FA plaques (A–D), whereas Y-27632 prevents S1P-induced SF accumulation (E). Interestingly, the treatment with blebbistatin (F) does not interfere with SF assembly. The images are representative of 5–6 independent experiments with similar results. G: quantitative analysis of radial and longitudinal SF fluorescence in C2C12 myoblasts in the indicated experimental conditions. AU, arbitrary units. Values are means ± SE from 3 separate experiments. *,{circ}Parameters that are significantly different at P < 0.05 compared with those of control cells.

 
The actin network has been reported to be the major molecular constituent in determining cell stiffness (27). Therefore, cell stiffness in C2C12 cells undergoing cytoskeletal remodeling was quantified by estimating the apparent elastic Young's modulus from the statistical analysis of force-indentation curves (Fig. 2, A and B) obtained using modified AFM spherical tips as microindenters. Young's modulus is indicative of the extension to which myoblasts deform after application of a stress, and its increase indicates reduced passive deformation after application of a load (5). Young's modulus was estimated in the peripheral regions to exclude the nuclear zone and was estimated taking into consideration the first 400 nm on the approaching indentation curve to evaluate the subcortical and radial cytoskeleton (see MATERIALS AND METHODS and Fig. 2A). As expected, this parameter increased significantly (P < 0.001) after SF formation by S1P (Fig. 2B). Since SFs are considered as temporary cytoskeletal structures composed by filamentous actin (F-actin) cross-linked by myosin IIs to form tensile bundles (4), we evaluated whether alterations of SF formation and/or contraction could affect C2C12 cell microelasticity. To this end, the myoblasts were first stimulated with S1P in the presence of 10 µM Y-27632, a specific inhibitor of Rho kinase, which has been shown to regulate S1P-induced SF assembly in these cells (10, 11). In these experimental conditions, C2C12 myoblasts underwent disorganization of SFs (Fig. 1, E and G) in coincidence with a drastic decrease (P < 0.01) of Young's modulus (Fig. 2B). To inhibit SF contraction, the cells were then stimulated with S1P in the presence of 10 µM blebbistatin, a specific inhibitor of myosin II ATPase activity (1). Short-term treatment (30 min) with blebbistatin also significantly (P < 0.01) reduced Young's modulus (Fig. 2B) without perturbing much actin filament architecture and SF formation, (Fig. 1, F and G). All these data indicated that not SF formation alone but the generation of tensional forces within SFs contributed to the stiffening response of the myoblastic cells.


Figure 2
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  		Fig. 2. Effects of cytoskeleton on the elastic (Young's modulus) and viscous (hysteresis) properties of living skeletal myoblasts. C2C12 cells were cultured and treated as reported in Fig. 1 and examined with a modified spherical atomic force microscopy (AFM) tip to perform force-indentation curves and evaluate the viscoelastic properties of the cell. A: representative AFM force-indentation curves performed in C2C12 cells stimulated with S1P (continuous lines) and in C2C12 cells treated with blebbistatin and stimulated with S1P (dashed lines). The indentation was calculated as the difference between the scanner position and cantilever deflection. B: apparent Young's elastic modulus in the indicated treatments affecting actin cytoskeletal status. Young's modulus was obtained from force-indentation curves using Hertz's modified model and was estimated taking into consideration the first 400 nm on the approaching indentation curve. C: normalized hysteresis in the indicated treatments affecting actin cytoskeletal status. Hysteresis was calculated as the difference between the areas under the extension curve and that of the retraction curve normalized by the input energy. In each experimental condition, data are from 4–6 cells from three different experiments. Statistical analysis was performed by using ANOVA test. *P < 0.05, **P < 0.01 and *** P < 0.001.

 
To correlate subcortical actin cytoskeleton with local variations of the topographic (heights) and stiffness features of the cell surface, simultaneous topographic and stiffness images were acquired by PFM-AFM. As observed by comparing the pseudo-color scales of the topographic and elastic maps (Fig. 3, A and C), the peripheral regions of the cells appeared lower (darker areas, Fig. 3A) and stiffer (brighter areas, Fig. 3C) as compared with the nuclear zones. Accordingly, the deflection AFM images revealed in the peripheral areas the presence of well-defined filamentous structures overhanging the plasma membrane (Fig. 3B); the cytoskeletal bundles ran along the entire cells leading to the formation of prominent cell surface corrugations. Of note, as shown in the line profiles (Fig. 3D) (evaluated along the white lines depicted in Fig. 3, A and C, and selected in the peripheral regions of the cells to avoid the nucleus), the local higher stiffness values corresponded to the higher topography values, presumably due to the presence of SFs underneath the plasma membrane. Cell surface topography (Fig. 3E) and stiffness (Fig. 3G) of the same cell appeared greatly modified after 30 min of S1P stimulation. In particular, as shown in the deflection image (Fig. 3F), the surface corrugations in the peripheral regions of the cells appeared less pronounced, and the sizes (depths) of depressions, in the topographic line profile (Fig. 3H), were reduced in S1P-stimulated cells compared with unstimulated cells (from 0.24 ± 0.03 of controls to 0.17 ± 0.03 µm of S1P stimulated cells; means ± SD, P < 0.05), suggesting that formation of robust SFs could contribute to mechanically stretch and flatten the plasma membrane.


Figure 3
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  		Fig. 3. AFM imaging and analysis of living skeletal myoblasts. C2C12 cells were grown on glass coverslips and examined before (A–D) and after (E–H) stimulation with 1 µM S1P for 30 min. In particular, C2C12 cells were first imaged by Pulsed Force Mode; then the same cells were stimulated with S1P and imaged again after 30 min of stimulation. Representative topography (A and E), deflection (B and F), and stiffness (C and G) images were acquired simultaneously. In the topography images (A and E), the higher zones are represented in brighter colors. In the deflection images (B and F), the local variations of heights appear more evident. In the stiffness maps (C and G), the brighter colors correspond to stiffer areas. D and H: AFM height and stiffness profiles evaluated along the horizontal white lines depicted in the images. The lines were selected in peripheral regions of the cells where the measurements were not influenced by the nucleus. Stiffness values are reported in arbitrary units with a maximal value of 1 AU corresponding to the glass. A close correlation (arrows) between the higher fluctuations of stiffness and the corrugation peaks in the topography profile was observed. Of note, the topographic line profiles show that the surface corrugations, because of the presence of fibrous structures underneath, are less pronounced and more densely packed in S1P-stimulated cells compared with those of unstimulated ones, suggesting that SF assembly could mechanically stretch (flatten) the cell surface. Images are representative of 4–5 independent experiments with similar results.

 
As expected, the treatment with Y-27632 of S1P-stimulated myoblasts caused smoothening (Fig. 4, A, B, and D) and softening (Fig. 4, C and D) of the cell surface, in coincidence with the disappearance of typical local variations. Of interest, stimulation of the cells in the presence of blebbistatin abolished not only the normal development of cytoskeleton-mediated cell surface stiffness (Fig. 4, G and H) but also the presence of the peripheral corrugations in the AFM deflection images (Fig. 4, E, F, and H). All these data taken together suggested that the assembly of contractile SF could generate tension in the plasma membrane of skeletal myoblasts.


Figure 4
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  		Fig. 4. AFM imaging and analysis of living skeletal myoblasts with altered cytoskeletal status. C2C12 cells cultured on glass coverslips were treated with 10 µM Y-27632 (A–D) or with 10 µM blebbistatin (E–H), stimulated with 1 µM S1P for 30 min, and then examined with Pulsed Force Mode. Representative topography (A and E), deflection (B and F), and stiffness (C and G) images were acquired simultaneously. D and H: AFM height and stiffness profiles evaluated along the white lines depicted in the images. The lines were selected in peripheral regions of the cells where the measurements were not influenced by the nucleus. Note that both disruption of SFs and inhibition of SF contraction elicit a remarkable smoothening of the cell surface associated with substantial loss of the surface rigidity. The images are representative of 4–5 independent experiments with similar results.

 
Effects of Stress Fibers on the Viscous Properties of C2C12 Cells Since the true behavior of biological systems is viscoelastic, the effects of actin cytoskeletal remodeling on hysteresis were also evaluated. This parameter is indicative of the viscous energy dissipation into the cell during AFM indentation (5, 26, 46). Interestingly, either formation of SFs by S1P (P < 0.05), or SF disassembling by treatment with Y-27632 (P < 0.05), or SF contractility inhibition by blebbistatin (P < 0.01) resulted in a small but significant increase in hysteresis relative to control cells (Fig. 2C), suggesting a role for actin dynamics in the regulation not only of the elastic but also of viscous properties of C2C12 myoblasts.

Effects of Stress Fibers on Membrane Capacitance of C2C12 Myoblasts To further correlate the stiffening response related to SF contraction with the possible development of plasma membrane tension, membrane capacitance (Cm) from the capacitative current transients defined in voltage-clamp experiments was measured and used as an index of the electrically accessible membrane area. This was done in view of the previous reported findings showing that tension in the cytoskeleton normally balances plasma membrane bleb formation and creates a tension in the bilayer (6). Of note, stimulation with S1P caused a small (10.5%) but significant (P < 0.05) decrease of Cm (Fig. 5A), whereas treatments with either Y-27632 or blebbistatin produced opposite effects, increasing significantly Cm (~26%, P < 0.05, and 34%, P < 0.01, respectively) (Fig. 5A). Notably, an apparent negative linear correlation was found between Cm and Young's modulus (E) [E (kPa) = –0.22 Cm + 5.8, where Cm is in pF; R2 = 0.93] (Fig. 5B). These data indicated that SF contraction was associated with reduced plasma membrane area, stressing the role of cytoskeletal contraction in the regulation of plasma membrane remodeling and stretching. Of note, the plot of Cm and normalized hysteresis showed a U-shaped relation (Fig. 5C), suggesting that any deformation (either increase or decrease) of the cell surface area was able to increase cell viscosity.


Figure 5
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  		Fig. 5. Effects of cytoskeleton on membrane capacitance (Cm) of living skeletal myoblasts. C2C12 cells were cultured and treated as reported in Fig. 1 and examined by patch-clamp analysis to measure Cm. Cm was used as an index of cell surface area. A: Cm in the indicated experimental conditions. Statistical analysis was performed by using ANOVA test; *P < 0.05. B: relationship between Cm and Young's modulus. Young's modulus shows a linear inverse relation with Cm. C: relationship between Cm and normalized hysteresis. Normalized hysteresis shows a U-shaped relation with Cm. For each experimental condition, electrophysiological (Cm) and AFM data (Young's modulus and hysteresis) are from different cells (15–19 for the electrophysiology and 4–6 for the AFM analyses) of parallel experiments.

 
Effects of Stress Fibers on SAC Activity of C2C12 Cells On the basis of the above reported findings, we next examined whether cytoskeleton-mediated effects (stiffness, surface topography, and Cm) were associated with parallel modifications in SAC current using patch-clamp analyses. SACs are, in fact, transmembrane channels that are activated by tension in the bilayer (25). It was found that S1P had marked effects on SAC activity and gating (Fig. 6); the addition of the sphingolipid to the culture medium resulted, in fact, in an evident increase (P < 0.005) of the amplitude of transmembrane ion current (Im) (Fig. 6, A and B) and conductance (Gm) (Fig. 6, F and G) as compared with those of controls (unstimulated cells). The treatments with either Y-27632 or blebbistatin resulted in a significant (P < 0.001) reduction of Im and Gm in the stimulated cells, (Fig. 6, C, D, F, and G). As expected, the current passing through these channels was efficiently inhibited in S1P-stimulated cells by 50 µM GdCl3, a commonly used SAC inhibitor (Fig. 6, E and F). Moreover, the addition of GdCl3 induced an 8% (P < 0.05) increase of Cm in S1P-stimulated cells (Fig. 6F), implying a possible effect of calcium passing through SACs in SF contraction and cell surface regulation. By contrast, the inhibitor did not cause any significant changes of both Gm and Cm in S1P-stimulated cells in the presence of either Y-27632 or blebbistatin. In these latter experimental conditions, only the residual current, which represents the current leak, could be evaluated, denoting that membrane tension was under threshold for SAC activation. Of interest, there was a negative exponential correlation between Gm and Cm {Gm (in nS) = 6 exp[–(Cm – 18.5)/0.9] + 0.57, where Cm is in pF} and a positive exponential correlation between and Gm and Young's modulus [Gm (nS) = 0.0044 exp(E/0.28) + 0.56, where E indicates Young's modulus in kPa] (Fig. 6, F and G), indicating that the increase of Young's modulus (190%) and the little decrease (10.5%) of cell surface due to S1P-induced SF contraction were both associated with dramatic activation of SAC-mediated current (530%).


Figure 6
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  		Fig. 6. Stretch-activated channel (SAC) current in living skeletal myoblasts with different cytoskeleton organization. Im, membrane current. A–E: representative current traces recorded by whole cell patch clamp in single C2C12 cells in control condition (A), after 1 µM S1P stimulation for 30 min (B), treated with 10 µM Y-27632 and stimulated with S1P (C), treated with 10 µM blebbistatin and stimulated with S1P (D), and stimulated with S1P and recorded in the presence of GdCl3 (E). Voltage-clamp step pulses between –80 and 0 mV in 10-mV steps from a holding potential of –60 mV were applied to the cells for 100 ms. F: relationship between plasma membrane conductance (Gm) and Cm. The best fit for Gm versus Cm was obtained by a single negative exponential function plus a constant. The plots refer to myoblasts in the indicated (filled symbols) experimental conditions. Data recorded in the presence of GdCl3 (open symbols) are also reported. G: relationship between Gm and Young's modulus. Gm versus Young's modulus plot was obtained by a single positive exponential function, plus a constant. For each experimental condition, electrophysiological (Gm and Cm) and mechanical data (Young's modulus) are from different cells (15–19 for the electrophysiology and 4–6 for the AFM analyses) of parallel experiments.

 
To stress the correlation between cytoskeletal contraction and SAC activation, we tested whether SAC sensitivity to mechanical stimulation also changed in concomitance to actin cytoskeletal remodeling. To this purpose, unstimulated and S1P-stimulated C2C12 cells were mechanically stretched using two microelectrodes (Fig. 7, A and B), as previously reported (47). A stretch of ~10% determined an increase of 1.5 ± 0.3 and 3.7 ± 0.4 folds of Gm in control cells (n = 5) and S1P-stimulated cells (n = 6), respectively, compared with cell at resting length (Fig. 7, C–F), indicating that S1P almost doubled SAC sensitivity to stretch. Additional experiments were performed in cells loaded with the fluorescent Ca2+ indicator, Fluo3-AM and then stretched using the tip of an AFM probe. After mechanical stretching, the intracellular Ca2+ fluorescence increased close to the tip and then spread to adjacent regions of the cell (Fig. 8), both in control and S1P-stimulated cells. However, the general amplitude of the fluorescence changes was quite different between the two cell preparations. In fact, consistent with electrophysiological measurements, the relative fluorescence increase [{Delta}F/Fb] was about twofold higher in the S1P-stimulated cells compared with control cells. All these data taken together pointed to the role played by SF formation elicited by S1P stimulation in the regulation of SAC sensitivity to stretch stimulation.


Figure 7
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  		Fig. 7. SAC current after mechanical stretch of control and S1P stimulated skeletal myoblasts. A and B: representative images obtained when two patch pipettes were attached on the cell at resting length (A) and after 10% cell stretch induced by the movement of the lower pipette in the longitudinal direction (B). C–F: time course of SAC-mediated currents (Im) in control and S1P-stimulated cells before (C and E) and after (D and F) the application of the cell stretch.

 

Figure 8
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  		Fig. 8. Visualization of Ca2+ entry in skeletal myoblasts after AFM mechanical stimulation. Control cells (A–D) and S1P-stimulated C2C12 cells (E–H) were preloaded with Fluo3-AM and mechanically stimulated with the rectangular tip of an AFM probe (gray lines). Fluorescence images were acquired at a rate of 1 image/s. The pseudo-coloring represents the global Ca2+ increase as indicated by the color bar. I: histogram of relative fluorescence increase ({Delta}F/Fb) in control and S1P-stimulated cells (means ± SE, *P < 0.05).

 
Effects of Stress Fibers on SAC Expression It has been recently reported that mechanisms involved in cytoskeletal remodeling may have a critical role in controlling the plasma membrane insertion of ion channel proteins (2). On these bases, we evaluated whether S1P-induced SF formation was associated with changes in the surface expression of TRPC1, which has been recently identified as a component of SAC in several cell types including C2C12 cells (25, 34, 38). By confocal microscopy, it was observed that TRPC1 protein was expressed in a vesicular manner in the cytoplasm and at the plasma membrane, as assayed by colocalization with TRITC-WGA lectin (Fig. 9), in both control and S1P-stimulated cells. Moreover, the plasma membrane insertion of this protein did not vary in S1P-treated cells compared with control cells. In fact, the overlap coefficient between TRPC1 and the plasma membrane-specific signals (approximately 0.60–0.65) was similar in the two cell preparations, leading us to exclude the involvement of increased TRPC1 membrane association in cells undergoing SF contraction and further suggesting that this latter event was the main contributor to SAC activation in S1P-stimulated cells.


Figure 9
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  		Fig. 9. Effects of stress fiber formation on plasma membrane localization of TRPC1 in skeletal myoblasts. Cellular localization of TRPC1 in control myoblasts (A–D) and S1P-stimulated C2C12 myoblasts (E–H) immunostained with polyclonal anti-TRPC1 followed by Alexa488-conjugated IgG and counterstained with TRITC-WGA lectin to visualize plasma membrane. The images refer to single confocal optical section. A and E: plasma membrane staining is colored in red. B and F: TRPC1 staining is colored in green. C and G: merged red and green images showing colocalization (yellow). D and H: scatterplots showing the distribution of sampled pixels plotted as a function of the red (y-axis) and green (x-axis) emission intensity; colocalized pixels in the image are included in the yellow orange region between the two white lines. I: histogram showing the overlap coefficient between TRPC1 and plasma membrane in control and S1P-stimulated cells. The images are representative of 5 independent experiments with similar results.

 

   DISCUSSION
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 MATERIALS AND METHODS
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There is substantial evidence that actin cytoskeletal remodeling is a crucial regulator of SAC activity in numerous cell types (9, 19, 20, 23, 37, 44). However, the mechanisms involved in this process are incompletely understood. In the current study, we provided the first experimental evidence that SFs regulates gating behavior of SACs modulating plasma membrane stretching and tension development. In particular, we have shown that formation of a well-structured cytoskeleton coupled with the plasma membrane via focal adhesions is associated with an increase in cell stiffness in C2C12 myoblasts stimulated with S1P. These results agree with previous reports obtained in other cell systems and in models of cell envelopes showing that the plasma membrane microelasticity decreases with the increase of actin surface density (21, 27). Here, this phenomenon was associated with modifications of cell surface topography suggestive of membrane deformations and flattening and with reduction of cell surface area.

The use of specific pharmacological inhibitors of both SF formation and contraction allowed us to clarify the mechanisms underlying the effects of SF on cell stiffness. In fact, SFs are considered as temporary cytoskeletal structures composed by G-actin polymerized into filamentous actin (F-actin) cross-linked by myosin IIs to form tensile bundles. Myosin IIs are ATP-driven molecular motors that form an essential part of the motile machinery of SFs generating the forces to move actin filaments relative to one another (4). We have previously demonstrated that formation of SFs by S1P in C2C12 cells is mediated by the activation of Rho/Rho kinase (ROK) pathway, on the basis that the ectopic expression of RhoGDI (a physiological inhibitor of GDP dissociation from Rho proteins) or with a selective ROK inhibitor, Y-27632, is accompanied by a drastic disorganization of actin microfilaments (9–11). Several mechanisms have been envisioned to explain the role of Rho pathway in the cytoskeletal response. One possible scheme is that ROK inhibits the myosin phosphatase activity through the phosphorylation of the myosin-binding subunit of myosin phosphatase and increases the phosphorylation levels of myosin light chain, resulting in the Ca2+-free contraction of the acto-myosin system and in the following bundling of F-actin into SFs (11). Additional mechanisms involve the activation of other Rho-effectors, such as mDia and ERM proteins, that contribute to the cytoskeletal response, favoring accumulation of polymerized actin (10). Accordingly, we have demonstrated that stimulation with S1P of C2C12 cells in the presence of the specific Rho kinase inhibitor, Y-27632, resulted in a profound disorganization of SFs, thus emphasizing the importance of the Rho kinase activity for maintaining the integrity of SFs in these cells. This cytoskeletal response was associated with a significant decrease in C2C12 cellular stiffness as compared with untreated control cells. The experiments performed in cells treated with blebbistatin, a specific inhibitor of myosin II ATPase activity that reportedly prevents acto-myosin dissociation and the transition of filament interaction into force-producing state (1), allowed us to better clarify the mechanisms underlying the driving force of the cytoskeleton-induced cell stiffness and to dissect the roles of F-actin bundling from that of myosin II-driven SF contraction in this event. In fact, blebbistatin was able to virtually abolish the normal development of cytoskeleton-mediated cell stiffness, without substantially modifying SF formation in S1P-stimulated cells, leading to the suggestion that not SF polymerization alone but the generation of tensional stresses within SFs are responsible for the observed stiffening response to S1P.

Taking into consideration the tensegrity model of cell architecture proposed by Ingber (18), the organization and contraction of the submembrane cytoskeleton may generate tensional prestress against the plasma membrane. In line with this, we have shown that the surface corrugations in AFM topography were less pronounced in S1P-stimulated cells compared with unstimulated cells, suggesting that formation of robust SFs may contribute to mechanically stretch the plasma membrane. More importantly, we have also shown that SF formation and contraction were able to reduce the myoblastic cell surface area. Since regulation of cell surface area depends on plasma membrane remodeling (13, 39), it is likely that SF contraction may reduce plasma membrane bleb formation by inducing membrane internalization and vacuole formation and contribute to tension development in the bilayer, as also suggested previously (6). On the basis of these considerations, it is also possible that the increase in cell surface area observed in cells with altered SF formation and contraction (i.e., treated with Y-27632 and blebbistatin) may be due to the translocation and fusion of submembrane vacuoles to the cell membrane as a consequence of the reduced plasma membrane tension (6). The negative exponential correlation between Gm and Cm and the positive exponential correlation between and Gm and Young's modulus may point out that the increase of Young's modulus (190%) and the slight decrease (10.5%) of cell surface area caused by cytoskeletal contraction may act as an amplifier of the stress tension generated in the plasma membrane. The amplified stress tension may in turn determine the remarkable increase of SAC activation (530%). These assumptions were further strengthened by the results showing that SAC activity was strictly dependent on actin cytoskeletal remodeling; in fact, the amplitude of SAC-mediated current and stretch-activated Ca2+ influx reached the highest values in coincidence with maximal SF formation and contraction, whereas SAC opening was hampered and virtually abolished after cytoskeletal disassembly and inhibition of acto-myosin interaction.

Previous observations have shown that actin remodeling may affect cell surface viscosity. In particular, experiments performed in mouse dystrophic myotubes (39) and skeletal myoblasts (46) have shown that the loss of dystrophin and the accompanying alterations of other cytoskeletal components increase membrane hysteresis, probably due to an increase in surface viscosity resulting from altered cytoskeletal organization and coupling to plasma membrane. In such a view, the data reported here fit well with this idea, showing that both the treatments with Y-27632 and blebbistatin result in a small but significant increase in the myoblastic hysteresis. Of note, we have found that also formation of SFs increases hysteresis, underscoring the possible involvement of cytoskeleton and its interaction with specific membrane receptors in the modulation of viscous properties of cells. Taking into consideration the recent findings suggesting that membrane viscosity may influence inherent channel sensitivity (39), it is tempting to speculate that increased hysteresis following S1P-induced SF formation may be determinant in filtering the ions influx through SACs, thus enabling the cells to resist passive deformation and preventing them from excessive SAC activation.

In conclusion, these findings support our hypothesis that SFs are the major component responsible for the modulation of plasma membrane tension generation and SAC activation in skeletal myoblasts. However, formation of SFs alone does not provide the molecular basis for SAC activation, and SAC inhibition induced by blebbistatin, a specific inhibitor of myosin ATPase, indicates that the sustained contraction of SFs may represent the force-bearing event responsible for cell surface stretching and SAC opening. We hypothesize that the increased viscosity observed in S1P-stimulated cells may produce a proportional increase in plasma membrane stability against passive deformations, and prevent cells from abnormal SAC function.

Taking into consideration the emerging evidence of the roles played by SACs (9, 43) and cytoskeletal remodeling in skeletal muscle differentiation (9, 29), this mechanism of SAC activation may have profound implications in muscle development, regeneration, and diseases, underlying the need to understand the functions of the subplasmalemmal acto-myosin network and its ability to transmit tensional forces to the cell membrane.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
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REFERENCES
 
This study was supported by grants from the University of Florence (ex 60%) to L. Formigli, S. Zecchi-Orlandini, and F. Francini, and from Ente Cassa di Risparmio di Pistoia e Pescia to S. Zecchi-Orlandini.


   FOOTNOTES
 

Address for reprint requests and other correspondence: L. Formigli, Dept. of Anatomy, Histology, Forensic Medicine, Univ. of Florence, Viale Morgagni 85, 50134 Florence, Italy (e-mail: formigli{at}unifi.it)

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.

* F. Sbrana and C. Sassoli contributed equally to this work. Back


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L. Formigli, C. Sassoli, R. Squecco, F. Bini, M. Martinesi, F. Chellini, G. Luciani, F. Sbrana, S. Zecchi-Orlandini, F. Francini, et al.
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00014.2008v1
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