Role for stress fiber contraction in surface tension development and stretch-activated channel...

ROLE FOR STRESS FIBER CONTRACTION ON SURFACE TENSION DEVELOPMENT

AND STRETCH-ACTIVATED CHANNEL (SAC) REGULATION IN C2C12 MYOBLASTS

Francesca Sbrana1, Chiara Sassoli2, Elisabetta Meacci3, Daniele Nosi2, Roberta Squecco4,

Ferdinando Paternostro2, Bruno Tiribilli5, Sandra Zecchi-Orlandini2, Fabio Francini4, Lucia

Formigli2.

1C.S.D.C. - Dept. of Physics, University of Florence, Florence, Italy

2 Dept. Anatomy, Histology, Forensic Medicine, University of Florence, Florence, Italy

3Dept. Biochemical Sciences, University of Florence, Florence, Italy

4Dept. Physiological Sciences, University of Florence, Florence, Italy

5ISC-CNR, Institute for Complex Systems, Sesto Fiorentino, Italy

The first two authors contributed equally to this work.

Corresponding author:

Prof. Lucia Formigli

Dept. of Anatomy, Histology, Forensic Medicine

University of Florence

Viale Morgagni 85,

50134 Florence,

Italy

Phone number +39-0554271809

Fax number +39-0554379500

e-mail: [email protected]

Running head: cytoskeleton mediated SAC activation.

Articles in PresS. Am J Physiol Cell Physiol (May 14, 2008). doi:10.1152/ajpcell.00014.2008

Copyright © 2008 by the American Physiological Society.

Cytoskeleton mediated SAC activation

1

ABSTRACT

Membrane-cytoskeleton interaction regulates trans-membrane 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 which, tensed by myosin II

motor, activate SACs by modifying the topography, the viscoelastic (Young’s modulus and

hysteresis) and electrical passive (membrane capacitance, Cm) properties of the cell surface.

Stimulation with sphingosine 1-posphate (S1P) 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 cytoskeletal-mediated corrugations in the cell surface

topography, suggesting a contribution of SF contraction to plasma membrane stretching. Moreover,

membrane capacitance, 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.

Keywords actin remodeling, atomic force microscopy (AFM), Young’s modulus, membrane

capacitance, hysteresis.


2

INTRODUCTION

Stretch activated channels (SACs) are voltage-independent non selective 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, muscle cell contraction to regulation of gene

expression and cell differentiation (8, 9, 27). 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 supports 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 complicate 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


3

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 might also exert tension forces

at the plasma membrane and provoke SAC opening.

On the basis of the aforementioned, in this 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 non-invasive 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.


4

MATERIALS AND METHODS

Cell cultures and treatments.

Murine C2C12 skeletal myoblasts obtained from American Type Culture Collection (Manassas,

VA, USA), were cultured in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with

10% fetal calf serum (FCS), 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, USA). In parallel experiments, the cells were

treated for 30 min with the following reagents in the presence of S1P: Y-27632 (10

μΜ, Calbiochem) used to inhibit endogenous Rho kinase activation and SF formation, blebbistatin

(10 μM, Sigma) to inhibit myosin II ATPase activity.

Confocal immunofluorescence.

Cells grown on glass coverslips were fixed in 0.5% buffered paraformaldehyde (PFA) for 10 min at

room temperature, permeabilized with cold acetone for 3 min, washed in phosphate buffered saline

(PBS) and incubated with 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

albumine (BSA; Sigma) and 3% glycerol in PBS for 30 min, and incubated with either 1:100

dilution of rabbit polyclonal anti-FAK (Focal Adhesion Kinase, 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, USA). Negative controls were carried out

by replacing the primary antibodies with non immune 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


5

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, USA) and observed under a Bio-Rad MRC 1024 ES Confocal Laser Scanning

Microscope (CLSM) (Bio-Rad, Hercules, CA, USA) equipped with a Krypton/Argon (Kr/Ar) 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 60X/1.4NA 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 (NIH). Densitometric analysis of radial and longitudinal SFs

was performed measuring the average fluorescence intensity on regions of interest (ROI, 6 µm2

area) in single focal plane images of 3D confocal stacks, using ImageJ (NIH) software. At least 4

ROIs in 50 different cells were analyzed in each experiment (three preparations/experiment), and

the mean ± SE optical density (OD) was then calculated. In the co-localization 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

63X/1.40NA oil immersion objective. To minimize crosstalk between fluorochromes, the

fluorescent signals were acquired sequentially at excitation wavelengths of 488 nm and 543 nm.

Series of optical sections (1024 X 1024 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 co-localization

between TRPC1 and WGA fluorescence signals was performed by calculating the overlap

coefficient (ranging from 0, minimum co-localization degree, to 1, maximum co-localization

degree), using the Leica Application Suite Software. At least 50 different cells were analyzed in

each experiment (three preparations/experiment), and the mean ± SE overlap coefficient was then


6

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, USA).

Atomic Force Microscopy (AFM) measurements and imaging.

Force-indentation measurements. A PicoSPM-scanning probe microscope (Molecular Imaging Inc.

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 micro-indenters in order 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, 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 20nN), and then

retracted. The corresponding plots, so called force-distance curves, were collected. Deflection

signal was converted in nm; 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

about 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. In order 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-δ) curves. The 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(δ)=4R1/2Eδ3/2/3(1-

ν2), where F was the loading force, E the Young’s modulus, ν the Poisson ratio and δ the

indentation. The Poisson ratio was assumed to be 0.5, as previously reported for living cells (35)


7

and the indentation was calculated as difference between the scanner position and the cantilever

deflection. The 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 sub-

cortical cytoskeleton accordingly 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

curves for extension (Aext) and retraction (Aret) were calculated integrating over the indentation

depth and the difference between these areas (hysteresis) normalized respect to the input energy

Aext. The measurements were made at a velocity of about 1 μm/sec. Since it has been reported that

starting from a velocity of 0.25 μm/sec the viscous energy accounts for more than 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 were

expressed as mean ± S.E. Statistical analysis was performed by using ANOVA test (P<0.05).

AFM imaging. AFM imaging were performed on living cells, using the PicoSPM-scanning probe

microscope equipped with a Pulsed Force Mode (PFM) external unit (WITec, Wissenshaftliche

Instrumente und Technologie GmbH, 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 sub-cortical

(cytoskeletal) structures. A sinusoidal z modulation was applied by PFM to the z-axis scanner, and

the system worked in a non-resonant 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

vs time curve, acquired point by point. Rectangular Ultrasharp silicon cantilevers (NT-MDT & Co.

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


8

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 in order 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, 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,

trans-membrane 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 (τ) were used to calculate the cell

linear capacitance (Cm) and Ra from τ =RaCm. The measurement of membrane resistance (Rm) were

corrected for Ra and calculated from the steady-state membrane current (Im) using the relation: Rm

=(ΔV-ImRa)/Im, where ΔV 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=ΔQ(Rm+Ra)/RmΔV, where ΔQ is the sum of the time

integral of the current transient and Imτ 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 (Gm versus Cm relation). To

ensure that the registered trans-membrane currents occurred through putative SACs, parallel


9

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), Cm and normalized hysteresis, were evaluated by linear

and non-linear best fit. Data fitting used a nonlinear curve fitting procedure based on the Marquardt-

Levenberg algorithm (Sigmaplot 4 and Table Curve 3.10 by Jandel Scientific, CA, USA, and

Clampfit 6.02 by Axon Instruments, CA, USA). 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, LRS, with the same formalism previously

reported (12). The improvement of the fit was evaluated by χ2 statistics. The number of degrees of

freedom by using linear or one or two exponential terms was two or three or six. We used χ2

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 χ2

statistics exceeded 7.8. The data and errors followed a Gaussian distribution and were expressed as

mean ± SE. One-way ANOVA with repeated measures was utilized for multiple comparisons and α

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 centre of the cell and used for electrophysiological measurements. The other microelectrode,

sealed to 10-20 μm far, 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.


10

The extent of stretch was expressed as the percentage change in cell length (L) relative to the

original length: L = (Lstretch - Loriginal)/Loriginal × 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 [ΔF/Fb: ratio of fluorescence

difference, peak-basal (Fp-Fb), to basal (Fb) values], as previously reported (41).


11

RESULTS

Effects of stress fibers (SFs) 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, trans-membrane 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 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 Focal

Adhesion Kinase (FAK) (Fig. 1B), allowed us to show that these filaments terminated in focal

adhesion FAK-containing plaques which 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).

The actin network has been reported to be the major molecular constituent in determining

cell stiffness (28). 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.2A, and B) obtained using modified AFM spherical tips as a micro-

indenters. Young’s modulus is indicative of the extension to which myoblasts deform after


12

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 in

order to evaluate the sub-cortical and radial cytoskeleton (see 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. 1E, and G) in coincidence

with a drastic decrease (P<0.01) of the 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. 1F, 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.

To correlate sub-cortical actin cytoskeleton with local variations of the topographic (heights)

and stiffness features of the cell surface, simultaneous topographic and stiffness images were

acquired by Pulsed Force Mode-AFM. As observed by comparing the pseudo-color scales of the

topographic and elastic maps (Fig. 3A, 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


13

note, as shown in the line profiles (Fig. 3D) (evaluated along the white lines depicted in panel 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. 3, E and F) 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.

2H), were reduced in S1P-stimulated cells compared to unstimulated cells (from 0.24 ± 0.03 of

controls to 0.17 ± 0.03 μm of S1P stimulated cells; mean ± SD, P<0.05), suggesting that formation

of robust SFs could contribute to mechanically stretch and flatten the plasma membrane.

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 cytoskeletal-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.

Effects of stress fibers (SFs) 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 (SFs) on membrane capacitance (Cm) of C2C12 myoblasts.


14

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 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 (approximately 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) = -0.22Cm + 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

for 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.

Effects of stress fibers (SFs) on SAC activity of C2C12 cells.

On the basis of the above reported findings, we next examined whether cytoskeletal-mediated

effects (stiffness, surface topography and Cm) were associated with parallel modifications in SAC

current using patch clamp analyses. SACs are, in fact, trans-membrane 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 trans-membrane ion current (Im) (Fig.6, A and B) and

conductance (Gm) (Fig.6, F and G) as compared to 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


15

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 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 (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 the 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%).

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 about the 10% determined an increase of 1.5 ± 0.3 and 3.7 ± 0.4 folds of Gm in control

(n=5) and S1P stimulated (n=6) cells respectively, compared to 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 [ΔF/Fb] was about two fold higher in the S1P-stimulated cells


16

compared to control ones. 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.

Effects of stress fibers (SFs) 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 co-localization 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 of 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.


17

DISCUSSION

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, 28). 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 which form an essential part of the motile machinery of SFs generating the forces

to move actin filaments relative to each other (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


18

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 to untreated control cells. The experiments performed in cells treated

with blebbistatin, a specific inhibitor of myosin II ATPase activity which 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 cytoskeletal-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 cytoskeletal-

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 sub-membrane cytoskeleton may generate tensional pre-stress

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 to 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


19

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 sub-membrane 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

depended 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


20

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.


21

GRANTS. This paper was supported by grants from the University of Florence (ex 60%) to L.F.,

S.Z.O., F.F; from Ente Cassa di Risparmio di Pistoia e Pescia to S.Z.O.


22

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FIGURE LEGENDS

FIGURE 1. Confocal fluorescence analysis of cytoskeleton organization of skeletal myoblasts.

C2C12 cells were grown on glass coverslips, fixed with PFA 0.5% in PBS and stained with TRITC

labeled phalloidin to detect F-actin (red); in some experiments the cells were further incubated with

rabbit polyclonal anti-FAK antibody followed by Alexa488-conjugated IgG to reveal FA-adhesion

plaques (green). (A, B) Control, unstimulated C2C12 cells; (C, D) cells stimulated with 1μM S1P

for 30 min; cells treated with either (E) 10μM Y-27632 or (F) with 10μM blebbistatin in presence

of S1P. The fluorescence distribution and intensity of F-actin are shown in the reported pseudo-

color scale. In panels A and C two types of SFs, radial (underscored by continuous arrows) and

longitudinal (underscored by dotted arrows) aligned prevalently underneath the plasma membrane

are visible. (A-D) S1P elicits formation of dense SFs and FA plaques, whereas (E) Y-27632

prevents S1P-induced SF accumulation. Interestingly, (F) the treatment with blebbistatin 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. Mean ± SE from three separate experiments. *

and ° indicate parameters that are significantly different at P < 0.05 compared with those of control

cells.

FIGURE 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 Figure 1 and examined with a modified

spherical 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 difference between the scanner position and

cantilever deflection. (B) Apparent Young’s elastic modulus in the indicated treatments affecting


29

actin cytoskeletal status. Young’s modulus was obtained from force-indentation curves using the

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.

FIGURE 3. AFM imaging and analysis of living skeletal myoblasts.

C2C12 cells were grown on glass coverslips and examined prior (A-D) and after (E-H) stimulation

with 1μM S1P for 30 min. In particular, C2C12 cells were first imaged by Pulse Force Mode; then

the same cells were stimulated with S1P and imaged again after 30 min of stimulation.

Representative (A, E) topography, (B, F) deflection and (C, G) stiffness images acquired

simultaneously. (A, E) In the topography images, the higher zones are represented in brighter

colors. (B, F) In the deflection images, the local variations of heights appear more evident. (C, G) In

the stiffness maps, the brighter colors correspond to stiffer areas. (D, 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 unit (a.u.) with a maximal value of 1 a.u. 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, due the presence of fibrous structures underneath, are less

pronounced and more densely packed in S1P-stimulated cells compared to 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.

FIGURE 4. AFM imaging and analysis of living skeletal myoblasts with altered cytoskeletal

status.


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C2C12 cells cultured on glass coverslips were treated (A-D) with 10μM Y-27632 or (E-H) with

10μM blebbistatin, stimulated with 1μM S1P for 30 min and then examined with Pulse Force

Mode. Representative (A, E) topography, (B, F) deflection and (C, G) stiffness images acquired

simultaneously (D, 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.

FIGURE 5 Effects of cytoskeleton on membrane capacitance (Cm) of living skeletal myoblasts.

C2C12 cells were cultured and treated as reported in Figure 1 and examined by patch clamp

analysis to measure membrane capacitance (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. The 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.

FIGURE 6. SAC-current in living skeletal myoblasts with different cytoskeleton organization.

(A-E) Representative current traces recorded by whole cell patch clamp in single C2C12 cells (A)

in control condition, (B) after 1 μM S1P stimulation for 30 min, (C) treated with 10 μM Y-27632

and stimulated with S1P, (D) treated with 10 μM blebbistatin and stimulated with S1P, and (E)

stimulated with S1P and recorded in the presence of GdCl3. 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


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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.

FIGURE 7. SAC-current after mechanical stretch of control and S1P stimulated skeletal

myoblasts. (A-B) Representative images obtained when (A) two patch pipettes were attached on

the cell at resting length and (B) after 10 % cell stretch induced by the movement of the lower

pipette in the longitudinal direction. (C-F) Time course of SAC-mediated currents (Im) in control

and S1P stimulated cells (C, E) before and (D, F) after the application of the cell stretch.

FIGURE 8. Visualization of Ca2+ entry in skeletal myoblasts after AFM mechanical

stimulation. (A-D) Control and (E-H) S1P-stimulated C2C12 cells were pre-loaded with Fluo3-

AM and mechanically stimulated with the rectangular tip of an AFM probe (grey lines).

Fluorescence images were acquired at a rate of 1 image/sec. The pseudo-coloring represents the

global Ca2+ increase as indicated by the color bar. (I) Histogram of relative fluorescence increase

(ΔF/Fb) in control and S1P-stimulated cells (mean ± SE, * P<0.05).

FIGURE 9. Effects of stress fiber formation on plasma membrane localization of TRPC1 in

skeletal myoblasts.

Cellular localization of TRPC1 in (A-D) control and (E-H) S1P-stimulated C2C12 myoblasts

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, E) Plasma membrane staining is colored in red; (B, F) TRPC1 staining

is colored in green; (C, G) merged red and green images showing co-localization (yellow); (D, 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) The histogram shows the overlap coefficient between


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TRPC1 and plasma membrane in control and S1P-stimulated cells. The images are representative of

5 independent experiments with similar results.

Role for stress fiber contraction in surface tension development and stretch-activated channel...

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