Stretch-activated cation channels in skeletal muscle myotubes from sarcoglycan-deficient hamsters

Tomoe Y. Nakamura1,3, Yuko Iwata1, Maurilio Sampaolesi1, Hironori Hanada1, Naohiro Saito2, Michael Artman3,4, William A. Coetzee3,4, and Munekazu Shigekawa1

1 Department of Molecular Physiology, National Cardiovascular Center Research Institute, Suita, Osaka 565-8565; 2 Research Institute, Kowa Company, Higashimurayama, Tokyo 189-0022, Japan; and 3 Pediatric Cardiology and 4 Physiology and Neurosciences, New York University School of Medicine, New York, New York 10016


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Deficiency of delta -sarcoglycan (delta -SG), a component of the dystrophin-glycoprotein complex, causes cardiomyopathy and skeletal muscle dystrophy in Bio14.6 hamsters. Using cultured myotubes prepared from skeletal muscle of normal and Bio14.6 hamsters (J2N-k strain), we investigated the possibility that the delta -SG deficiency may lead to alterations in ionic conductances, which may ultimately lead to myocyte damage. In cell-attached patches (with Ba2+ as the charge carrier), an ~20-pS channel was observed in both control and Bio14.6 myotubes. This channel is also permeable to K+ and Na+ but not to Cl-. Channel activity was increased by pressure-induced stretch and was reduced by GdCl3 (>5 µM). The basal open probability of this channel was fourfold higher in Bio14.6 myotubes, with longer open and shorter closed times. This was mimicked by depolymerization of the actin cytoskeleton. In intact Bio14.6 myotubes, the unidirectional basal Ca2+ influx was enhanced compared with control. This Ca2+ influx was sensitive to GdCl3, signifying that stretch-activated cation channels may have been responsible for Ca2+ influx in Bio14.6 hamster myotubes. These results suggest a possible mechanism by which cell damage might occur in this animal model of muscular dystrophy.

muscular dystrophy; calcium; cell membrane


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

MUSCULAR DYSTROPHY is a heterogeneous genetic disease that affects striated muscle (skeletal as well as cardiac muscle). The genetic defects associated with muscular dystrophy often include mutations in one of the components of the dystrophin-glycoprotein complex (DGC), such as dystrophin or sarcoglycan (SG).

The DGC is a multisubunit complex (4, 36) that spans the sarcolemma to form a structural link between the extracellular matrix and the actin cytoskeleton (7). The proteins of the DGC are structurally organized into distinct subcomplexes. In skeletal and cardiac muscle, beta -dystroglycan forms the membrane-spanning linkage between alpha -dystroglycan (receptor for laminin-2) and cytosolic dystrophin, which directly binds to the actin cytoskeleton. This complex is associated with four transmembrane glycoproteins, the SGs (alpha -, beta -, gamma -, and delta -SG). The SGs are combined in a larger complex together with sarcospan, the SG-sarcospan subcomplex (5, 27). The SG-sarcospan subcomplex can presumably strengthen the binding of alpha -dystroglycan to the sarcolemma, thereby stabilizing the transmembrane DGC. Disruption of DGC could, therefore, affect membrane integrity and/or stability and maintenance during muscle contraction and relaxation.

Mutations in dystrophin, laminin-2, and SGs each give rise to different forms of muscular dystrophy in humans and in animal models (4, 27). For example, Duchenne and Becker muscular dystrophies are caused by a variety of mutations in the dystrophin gene. The mdx mouse, a model for this form of dystrophy, also carries a loss-of-function mutation in dystrophin (3, 17). In this animal model, dystrophin-associated proteins including SG-sarcospan subcomplex and dystroglycans are also absent or greatly reduced (4, 26, 27). Skeletal fibers from mdx mice as well as cultured myotubes from skeletal muscle of Duchenne patients were reported to have chronically elevated levels of intracellular Ca2+ (9, 38), although this has not been confirmed in some other studies (2, 13). This could be partly caused by a high basal activity of Ca2+-permeable channels (Ca2+-leak channels) (9) or mechanosensitive Ca2+-permeable channels (11, 12), as has been reported in cultured myotubes from mdx mice. In contrast, in dy/dy mice, an animal model for a classic form of human congenital muscular dystrophy caused by a defect in the laminin-2 gene, the activity of the mechanosensitive Ca2+-permeable channels is not elevated (12), although these animals were reported to have elevated levels of intracellular Ca2+ in skeletal muscle (6, 39).

Mutation of SG genes causes an autosomal recessive limb girdle type of muscular dystrophy in humans (4, 27). The Bio14.6 hamster, which has a defective delta -SG gene (25, 32), has long been used as an animal model for autosomal recessive cardiomyopathy, although it exhibits extensive fiber damage in both skeletal and cardiac muscles. Deficiency of delta -SG in Bio14.6 hamsters causes disruption of the DGC and almost complete loss of other SGs as well as reduction of alpha -dystroglycan (16, 20, 30, 34). However, dystrophin and beta -dystroglycan are still retained at approximately one-half of the normal levels in the myopathic hamster myocytes (19, 20, 30), suggesting that the selective loss of the SG complex may be sufficient to cause muscular dystrophy (1, 24).

At present, little is known about normal and pathological functions of SGs and the ion-handling properties of skeletal myocytes from delta -SG-deficient Bio14.6 hamsters. In this study, we used cultured myotubes isolated from skeletal muscle to investigate the possibility that ionic conductances are altered in Bio14.6 hamsters and that these altered ionic conductances may lead to abnormal intracellular ionic homeostasis, which might contribute to the myocellular damage observed in these animals.


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Animals. Male Bio14.6 hamsters (J2N-k strain) between 60 and 70 days of age and age-matched normal controls (J2N-n) maintained at the animal facility of Kowa Pharmaceutical Company were studied. The J2N-n had the same genetic background as the J2N-k, except for the difference of a genetic locus for cardiomyopathy. Hamsters were anesthetized with ether, and their skeletal muscles were then excised according to guidelines for animal experimentation at the National Cardiovascular Center.

Cell culture. Satellite cells were isolated from extensor digitorum longus muscles by enzymatic dissociation. Minced muscles (0.3 g) were incubated in 1 ml of an enzyme cocktail that contained 0.25% trypsin/1 mM EDTA (GIBCO BRL, Tokyo, Japan), 0.1% collagenase (Worthington), and 10 U/ml DNase I (GIBCO BRL) at 37°C for 30 min. Undigested muscles were removed by centrifugation (700 rpm, 5 min). The supernatant was diluted with 9 ml of DMEM and then centrifuged (1,400 rpm, 5 min). Isolated cells were resuspended in 1 ml of DMEM supplemented with 20% fetal bovine serum (GIBCO BRL) and 1% chick embryo extract (GIBCO BRL). This cell isolation procedure was repeated four times using undigested muscles. Dissociated cells were combined and collected by centrifugation. Cell suspension was filtered through a fine mesh nylon filter (100 µm) and subjected to preplating at 37°C for 30 min to remove fibroblasts (differential adhesion). Nonadhering cells were plated onto collagen-coated (100 µg/ml collagen type I; Sigma, St. Louis, MO) culture dishes at a density of 5,000 cells/cm2. When myoblasts were grown to 80% confluency, they were trypsinized and then plated on collagen-coated glass coverslips for electrophysiological experiments. After 2-3 days, medium was changed to DMEM containing 2% horse serum (Hyclone) to initiate differentiation. Myoblasts began to fuse and form myotubes in culture within 48 h. Recordings were made from myotubes 3-5 days after the first myotubes formed.

Single-channel recording. Standard patch-clamp techniques were used to obtain single-channel recordings using an Axopatch 200A amplifier and pCLAMP software (Axon Instruments). Cell-attached patch configurations were used for the experiments. Patch electrodes were prepared from thick-walled glass capillaries (1.5-mm outside diameter, 1.12-mm internal diameter) and heat-polished. When filled with a pipette solution, electrode resistance ranged between 2 and 4 MOmega , which corresponds to tip diameters of 1-2 µm. For most experiments, the pipette solution contained 110 mM BaCl2 in 3 mM HEPES (pH 7.4) solution. For some experiments, 165 mM KCl, 165 mM NaCl, or 165 mM sodium glutamate in 3 mM HEPES (pH 7.4) was used as the pipette solution. The bath solution contained 150 mM potassium aspartate, 5 mM MgCl2, 5 mM EGTA, 10 mM glucose, and 10 mM HEPES (pH 7.2 adjusted with LiOH). Thus the resting potential is expected to be close to 0 mV. The bath solution was continuously perfused at a constant flow (1 ml/min) at room temperature. Currents were filtered through an eight-pole Bessel low-pass filter 9002 (Frequency Devices) at 5 kHz and acquired at 20 kHz (pCLAMP).

Measurement of mechanosensitivity. To investigate the mechanosensitivity of channels in cell-attached patches, we applied positive or negative pressures to the pipette interiors. After successful seal formation, the side part of the pipette holder was connected to a commercial device (X-caliber, Viggo-Spectramed) that allows fine control of applied pressure/vacuum. This device also incorporates a pressure sensor and a digital display for monitoring developed pressure/vacuum. After manually increasing the pressure, a stable reading was obtained within 1 s, and this reading remained stable throughout the duration of the recording (usually ~10 s). Although not necessary in the majority of recordings, small deviations from the desired value were corrected iteratively throughout the recording. It has been reported that mechanical overstimulation of the patch, during or after tight seal formation, may result in an ion channel with altered mechanosensitivity (15). In contrast, seal formation with gentle suction (<5 mmHg for 10 s or less) best preserves mechanosensitivity when using standard (2-µm tip diameter) patch pipettes (14, 33). We therefore used a gentle sealing protocol to avoid changes in mechanosensitivity. Pipettes were manufactured using a programmable vertical puller (DMZ-Universal puller; Zeitz-instrumente, Zeitz, Germany) to reproducibly obtain the pipettes with similar tip diameter. Mechanosensitivity of channels was studied using graded pressure applied to the pipette interior for <10 s. Under these conditions, channels responded to repetitive stimulations; sudden or huge change of mechanosensitivity (that may result from overstimulation) was not observed.

Data analysis. Data were analyzed using the pCLAMP suite of software (Axon Instruments) and Origin for Windows software (Microcal Software, Northampton, MA). The unitary current amplitude was measured using one of two methods. When traces were sufficiently stable and transition levels were well defined, we directly constructed all-point histograms or events-list histograms from the recorded data. These histograms were fitted to a sum of Gaussian distributions to determine the amplitude, the mean, and the dispersion of each peak. The mean unitary currents was calculated as the difference between the means of adjacent peaks. It was not always possible to obtain reliable computer-constructed histograms because of the fast kinetics of the channels. Under these conditions, the amplitudes of detectable transitions between current levels were measured manually. Distributions of open and closed times were obtained by performing events-list analysis from idealized records, using records in which only a single open level was observed. The idealized records were obtained by setting a threshold at one-half of the amplitude of the open channel current and considering an opening event to occur when at least two consecutive sample points crossed this threshold. The open and closed time histograms were fitted by the sum of two exponential functions.

Channel open probabilities (Po) were measured by integrating idealized records of channel opening and closing transitions and dividing this by the time integral of the single-channel current. The measured Po is the Po of each individual channel (Po) multiplied by the number of open levels (N). Data are expressed as means ± SE. Comparisons between data groups were performed using a Student's paired or unpaired t-test. Differences at P < 0.05 were considered statistically significant.

Fluorescence cytochemical analysis. Cultured myotubes prepared from skeletal muscle of normal or Bio14.6 hamsters were treated with cytochalasin D (10 µM) or DMSO (0.1%) for 15 min, fixed in 4% paraformaldehyde in PBS, and processed for fluorescence cytochemical analysis as described previously (40). After being permeabilized and blocked, samples were incubated with rhodamine-phalloidin (Molecular Probes, 1:1,000 dilution) to stain filamentous actin for 1 h at room temperature. Confocal images of myotubes were obtained using an MRC-1024 confocal microscope (Bio-Rad) mounted on an Olympus BX50WI epifluorescence microscope with a plan-apochromat ×60 water-immersion objective lens (Olympus).

Actin analysis. Normal or Bio14.6 myotubes were treated with cytochalasin D (10 µM) or DMSO (0.1%) for 15 min and homogenized in a buffer that contained 10 mM NaHCO3 and protease inhibitors. They were centrifuged at 5,500 g for 15 min at 4°C. The low-spin supernatants were centrifuged at 480,000 g for 40 min at 4°C. The high-spin pellets were then dissolved in RIPA buffer [150 mM NaCl, 15 mM HEPES-NaOH (pH7.5), 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS, and protease inhibitors] for 1 h at 4°C. The low-spin pellets contained materials such as nuclei, contractile proteins, and mitochondria that are insoluble in a low-salt solution. The high-spin supernatants contained cytosolic fractions with depolymerized G-actin, whereas the high-spin pellets contained the membranes with associated polymerized F-actin. Equal amounts of protein from these fractions were subjected to SDS-PAGE on 8.5% gel. Western blot analysis was performed using an anti-actin antibody at 1:1,000 dilution (Zymed) as described previously (20).

Measurement of 45Ca2+ uptake. Normal and Bio14.6 myotubes were preincubated at 37°C for 3 min in balanced salt solution (146 mM NaCl, 4 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 10 mM glucose, 0.1% bovine serum albumin, and 10 mM HEPES-Tris, pH 7.4) that contained 0 or 0.5 mM GdCl3. 45Ca2+ uptake into cells was initiated by switching to BSS that contained 45CaCl (10 µCi/ml) and was terminated after 5 min by washing cells four times with ice-cold 5 mM LaCl3, 146 mM choline chloride, and 10 mM HEPES-Tris, pH 7.4. Cells were lysed in 1% SDS plus 0.1 N NaOH, and aliquots were taken for determination of protein and radioactivity. 45Ca2+ uptake in the presence of Gd3+, which corresponded to ~20% of total uptake in normal cells at 5 min, was considered to be nonspecific binding, and the Gd3+-inhibitable fraction of 45Ca2+ uptake was calculated by subtraction from the uptake in the absence of Gd3+.

Materials. Nifedipine, gadolinium chloride hexahydrate, cytochalasin D, and DMSO were purchased from Sigma Chemical. alpha -Bungarotoxin was from Calbiochem (La Jolla, CA). Cytochalasin D was prepared as a stock at a concentration of 10 mM in DMSO and kept at -20°C.


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We recorded single-channel activity from cell-attached patches using myotubes from normal and Bio14.6 hamsters. As described in MATERIALS AND METHODS, a gentle suctioning protocol was used to form high-resistance seals, which occurred with equal success in normal and Bio14.6 myotubes. Figure 1A shows continuous recordings (~10 s) from normal (left) and Bio14.6 (right) hamster myotubes recorded at a constant holding potential of -60 mV (pipette potential of +60mV) using 110 mM BaCl2 in the pipette. In normal myotubes, most patches exhibited little channel activity under these conditions (Fig. 1A, left). Compared with normal myotubes, Bio14.6 myotubes had higher basal channel activity in most of the patches recorded (Fig. 1A, right). The channel, which occurred in irregular bursts, had a unitary current amplitude of ~1.2 pA. Figure 1B shows the summary of mean open probabilities (NPo) for the channel obtained from normal and Bio14.6 hamster myotubes. The mean values of NPo were 0.03 ± 0.01 in normal and 0.13 ± 0.04 in Bio14.6 myotubes, respectively.


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Fig. 1.   Comparison of the activity of Ba2+-permeable channels in myotubes from normal and Bio14.6 hamsters. Single-channel recordings were performed in the cell-attached configuration at a membrane potential of -60 mV with 110 mM BaCl2 in the pipette. The patch current was filtered at 5 kHz, and the acquisition sampling rate was 20 kHz. A: representative traces obtained from 10-s continuous recordings using normal (left) and Bio14.6 (right) hamster myotubes. Dotted lines represent the 0 current level. B: summary results for mean open probability (NPo) averaged from 22 and 24 patches from normal and Bio14.6 hamsters, respectively.

To determine the single-channel conductance, the unitary amplitude of the channel was determined by constructing histograms after events-list analysis. This procedure was performed for recordings at different membrane potentials (examples of current traces obtained at -60 mV are shown in Fig. 2B). The single-channel current was plotted as a function of the membrane potential to obtain the unitary conductance (Fig. 2C). With 110 mM BaCl2 as the pipette solution, the unitary current-voltage (I-V) relationship was linear in the voltage range studied (-20 to -100 mV). The mean slope conductance, which was obtained at voltages more negative than -20 mV, was 20.4 ± 2.3 pS (n = 4) in normal myotubes and 19.4 ± 1.8 pS (n = 4) for Bio14.6 myotubes (Fig. 2C). These values are not statistically different. These results suggest that a channel that permeates Ba2+, with a unitary conductance of ~20 pS, can be recorded in both groups but that its activity is substantially higher in skeletal muscles from Bio14.6 hamsters. Similarly, when the pipette was filled with 110 mM CaCl2, channel activity was recorded in both groups (with an increased activity in Bio14.6 myotubes; results not shown). However, with Ca2+ as the charge carrier, channel kinetics were extremely flickery, and the unitary current amplitude was much smaller (~8 pS), which precluded further detailed analysis.


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Fig. 2.   Single-channel conductance of the channel. Recordings were obtained using cell-attached patches from normal (left) or Bio14.6 (right) hamsters with 110 mM BaCl2 in the pipette solution. A: representative traces measured at -60, -80, and -100 mV of the membrane potentials. Dotted lines depict the single-channel amplitude. Note the different time scales used in this figure and in Fig. 1. B: amplitude histograms of the channel measured at -60 mV obtained from normal (left) and Bio14.6 (right) myotubes. An events-list histogram was constructed from 10-s recordings measured at different membrane potentials. Amplitude histograms were fitted to double Gaussian equations. C: unitary current-voltage relationship of the channel. A least-squares fit of the data was performed (dotted line) to obtain the slope conductance (n = 4 each).

Extrapolation of the I-V relationships shown in Fig. 2 toward more positive potentials yields reversal potentials that are significantly lower than the calculated equilibrium potentials of Ba2+, suggesting that the channel might also be permeable to other ions. To test this possibility, we used NaCl or KCl in the pipette in a separate group of experiments. As shown in Fig. 3A, a 28.6-pS channel was detected when 165 mM KCl was used as the pipette solution. Under these conditions, outward current was observed at positive membrane potentials, resulting in a linear I-V relationship. This channel activity was, therefore, not caused by the classic inward rectifier K+ channel, IK1, which was expected to exhibit strong inward rectification in the cell-attached configuration due to block by intracellular polyamines and Mg2+ (8, 21). With 165 mM NaCl in the pipette solution, a large 47.3-pS channel was observed. To avoid activation of the tetrodotoxin-sensitive Na+ current that opens at membrane potentials more positive than -60mV (29), we restricted our measurements to potentials more negative than this value (Fig. 3B). Separate experiments were also performed to exclude the possibility that the channel current was mediated by Cl- ion fluxes rather than by an influx of cations. We kept the Na+ concentration constant (and removed Cl-) by using sodium glutamate in the pipette. Under these conditions, a channel was observed having a unitary conductance identical to when NaCl was used in the pipette, thus eliminating Cl- as a significant charge carrier. These results demonstrate that the channel was also permeable to Na+ and to K+.


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Fig. 3.   Channel activities measured from Bio14.6 hamster myotubes using K+ or Na+ as the charge carrier. A: 165 mM KCl was used in the pipette solution. Currents were measured at +100 mV (a) or -60 mV (b-d), either at atmospheric pressure (a, b, and d) or with suction (-40 mmHg) applied to the pipette (c). In a different experiment, 500 µM GdCl3 was included in the pipette (d). B: NaCl (165 mM) was used in the pipette solutions. Currents were measured at -80 mV, either at atmospheric pressure (a) or with suction (-40 mmHg) applied to the pipette (b). Some experiments were performed with 500 µM GdCl3 in the pipette (c). C: sodium glutamate (165 mM) was used as the pipette solution. Currents were measured at a membrane potential of -60 mV, either at atmospheric pressure (a and c) or with negative pressures (-40 mmHg) applied to the pipette (b). In some experiments, 500 µM GdCl3 was included in the pipette (c). Dotted lines indicate the 0 current level. Unitary amplitude-voltage relationships and slope conductances, obtained by linear regression, are shown (right).

Using Ba2+ as the charge carrier, we performed a kinetic analysis of this channel and compared its properties in normal and Bio14.6 myotubes (Fig. 4). The distributions of open and closed times were best fitted with the sum of two exponential functions, suggesting that there might be multiple open and closed states. The mean open time was longer in patches isolated from Bio14.6 myotubes; most of this difference was attributable to the fast exponential (Table 1). The fast and slow time constants of closed duration were significantly smaller in Bio14.6 compared with normal channels (Table 1). These values reflect the longer opening and shorter closing times of the channels, which result in higher Po of the channels in Bio14.6 myotubes.


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Fig. 4.   Histograms of the cumulative distributions of open (A and C) and closed (B and D) times of channel activity recorded from normal (A and B) and Bio14.6 (C and D) hamster myotubes. Kinetic analysis was performed on a 20-s segment of recordings obtained at -60 mV, during which only one level of channel activity was observed. The distributions of open and closed times were best fitted with the sum of 2 exponential functions.


                              
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Table 1.   Time constants of open and closed durations

We next examined the effect of pressure-induced stretch on the activity of these channels in normal and Bio14.6 myotubes and compared the responses between these two groups (Fig. 5). Graded positive or negative pressures were applied to the pipette (which was filled with 110 mM BaCl2) in cell-attached patch configurations while the membrane was held at -60 mV. In normal myotubes, no channel activity was detected under atmospheric pressure in this particular patch (Fig. 5A, left, no applied pressure). Application of pressure to the pipette evoked channel activity, which had a similar unitary amplitude and kinetics at positive and negative pressures (Fig. 5A, left). Similarly, application of pressure induced a strong enhancement of the already higher basal channel activity in Bio14.6 myotubes (Fig. 5A, right). Data summarized in Fig. 5B show that channel activity was enhanced by both positive and negative pressures; this effect occurred in both experimental groups. However, the greater slopes of the positive and negative NPo-pressure relationships in the Bio14.6 myotubes indicate that the channels are more sensitive to stretch activation compared with those in normal myotubes. Application of patch pressure also enhanced the activities of the 28-pS channel or the 48-pS channel, respectively, when the monovalent cations K+ or Na+ were used as the charge carrier (Fig. 3, Ac, Bb, and Cb). These data suggest that the channel activity recorded with monovalent cations in the pipette may originate from the same channel that is active when using divalent cations (Ba2+) as the charge carrier. Because applied pressure further increases activity regardless of the charge carrier, this channel can be classified as a stretch-activated nonspecific cation channel.


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Fig. 5.   Effect of pressure-induced stretch on channel activities. Graded positive or negative pressures were applied to the pipette using cell-attached patches that were held at -60 mV. BaCl2 (110 mM) was used as the charge carrier. A: representative trace is a continuous recording in one patch from a normal (left) or a Bio14.6 (right) hamster myotube. Dotted lines indicate the closed level. B: stretch-activity relationships were obtained from 8 normal (left) and 8 Bio14.6 (right) hamster myotubes. *Statistically different (P < 0.05) from values obtained at atmospheric pressure.

To characterize the pharmacological properties of the channel, we examined the effect of GdCl3, a known blocker of stretch-activated channels (Fig. 6). Bio14.6 myotubes were first incubated in 5 µM GdCl3 (10 min to 1 h), and channel activity was tested using different concentrations of GdCl3 in the pipette (with Ba2+ as the charge carrier at a patch potential of -60 mV). This ensured that GdCl3 was applied to the extracellular face of the membrane. A similar experimental strategy was followed to examine the reversibility of GdCl3. For the latter experiments, GdCl3-incubated myotubes were patched using a pipette solution devoid of GdCl3 (to allow washout of bound gadolinium). When the pipette solution did not contain GdCl3, channel activity appeared within ~2 min after seal formation (due to the washout effect). With a pipette solution containing 500 µM GdCl3, however, no channel activity was observed even 10 min after seal formation (n = 6). We performed a parallel set of experiments using various concentrations of GdCl3 in the pipette solution, and we measured channel activity 9-12 min after seal formation. These results demonstrate that the inhibitory effect of GdCl3 was concentration dependent (Fig. 6). Furthermore, GdCl3 also blocked the channel during application of pressure to the pipette (Fig. 6). Similarly, no K+- or Na+-permeable channel activity was observed when 500 µM GdCl3 was included in the pipette solution either in the absence (Fig. 3, Ad, Bc, and Cc) or presence of pressure (data not shown).


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Fig. 6.   Concentration-dependent inhibition of channel activity by GdCl3. Myotubes from Bio14.6 hamsters were incubated in 5 µM GdCl3 for ~1 h, and channel activities were tested using different concentrations of GdCl3 in the pipette. BaCl2 (110m) was used as the charge carrier, and currents were measured at -60 mV. A: representative traces with varying concentrations of GdCl3 in the pipette obtained 9-12 min after the seal formation. Currents were measured under atmospheric conditions (0 mmHg) or under the condition that pressure was applied (-20 mmHg). B: data were obtained from 6 patches and summarized. A concentration-dependent inhibitory effect of GdCl3 on the channel activity was observed in both the absence () or presence (open circle ) of pressure (-20 mmHg).

We also examined the effects of blockers for other Ca2+-permeable channels. The L-type Ca2+-channel blocker nifedipine (1 µM) or its solvent DMSO (0.1%), applied from outside of myotubes as described earlier, had no effect on channel activity (data not shown). Because the spontaneous opening of acetylcholine receptor channels were observed more frequently in dy/dy mouse myotubes than in normal or mdx myotubes (12), we also used the blocker of this channel, alpha -bungarotoxin (31). This toxin similarly had no effect on channel activity (data not shown).

The physiological role of the SG subcomplex (which is disrupted in Bio14.6 hamsters) is largely unknown. SGs may have interactions with submembrane actin cytoskeleton, and disruption of the actin microfilaments may therefore lead to similar electrophysiological abnormalities as observed in Bio14.6 hamster myotubes. Accordingly, we tested the response of these channel activities to depolymerization of F-actin using cytochalasin D. Before performing electrophysiological measurements, however, we examined whether this reagent in fact depolymerized F-actin. Staining with phalloidin, which binds with high affinity to F-actin, revealed no apparent differences in the overall cell shape or macroscopic actin organization in either normal or Bio14.6 myotubes (Fig. 7, top). When using a more sensitive biochemical assay, however, we found that the F-actin content in membrane fractions was decreased and the G-actin content in the cytosolic fractions was significantly increased after cytochalasin D treatment (top panels, bottom insets), suggesting that under these conditions depolymerization of membrane-associated F-actin (i.e., cortical actin) occurred in both experimental groups.


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Fig. 7.   Effects of cytochalasin D (Cyt.D) on actin filaments in normal and Bio14.6 myotubes. Normal and Bio14.6 myotubes were treated with vehicle (top) or cytochalasin D (10 µM; bottom) for 15 min and then stained with rhodamine-phalloidin to visualize actin filaments using a confocal microscope. Insets: F-actin content in high-spin pellets (right lane) and depolymerized G-actin in high-spin supernatants (left lane) prepared from myotubes treated with DMSO or cytochalasin D were analyzed by Western blotting with anti-actin antibody (see MATERIALS AND METHODS).

We next examined the effect of cytochalasin D on the activity of the stretch-activated cation channel. As before, low levels of channel activity were observed under atmospheric conditions in normal myotubes (Fig. 8A, top left). Application of cytochalasin D (10 µM, ~15 min) led to a marked enhancement of channel activity even under atmospheric conditions (Fig. 8A, top right). This effect was further enhanced when pressure was applied to the patch membrane (Fig. 8A, middle right). Channel activity was similarly increased by treatment with cytochalasin D in the presence or absence of pressure in Bio14.6 myotubes (Fig. 8B, top and middle). The effect of cytochalasin D was completely prevented when 500 µM GdCl3 was included in the pipette in both normal and Bio14.6 myotubes (Fig. 8, A and B, bottom). The average values of NPo were 0.03 ± 0.01 (before) and 0.25 ± 0.09 (after) cytochalasin D treatment in normal myotubes and 0.13 ± 0.04 (before) and 0.37 ± 0.15 (after) cytochalasin D treatment in Bio14.6 myotubes, respectively (n = 4 each). The vehicle, DMSO (0.1%, ~15 min), had no such effects on the channel activity (data not shown).


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Fig. 8.   Effect of cytochalasin D on channel activity in normal (A) or Bio14.6 (B) hamster. Currents were measured at -60 mV with 110 mM BaCl2 as the pipette solution. Recordings were made in the absence (0 mmHg) or presence (+20 mmHg) of pressure applied to the pipette before (left) and ~15 min after application of cytochalasin D (10 µM; right). In different experiments, currents were measured in the presence of 500 µM GdCl3 in the pipette. Dotted lines depict the closed levels.

The results so far show that a cation-permeable channel, with low levels of basal activity, exists in skeletal muscle myotubes from hamster. We also demonstrate that the stationary activity of this channel is higher in muscle from Bio14.6 hamsters, which is expected to lead to an increased unidirectional cation influx. To examine whether these patch phenomena translate to an increased cation conductance in intact myotubes, we initiated Ca2+-influx experiments, using standard techniques. Our data show that relative to control, Ca2+ influx (as assessed by Gd3+-sensitive 45Ca2+ influx) was significantly higher in myotubes isolated from Bio14.6 hamsters (Fig. 9). The Gd3+ sensitivity of this assay is in strong support of the concept that Ca2+ influx occurred directly through the stretch-activated cation channels or that Ca2+ influx occurred through a pathway that is sensitive to the activity of these channels.


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Fig. 9.   Gd3+-sensitive 45Ca2+ uptake into intact myotubes under resting conditions. Normal and Bio14.6 myotubes were incubated in the balanced salt solution containing 45Ca2+ with or without 0.5 mM GdCl3 for 5 min, and 45Ca2+ uptake was measured. Gd3+-inhibitable fraction of 45Ca2+ uptake was calculated as described in MATERIALS AND METHODS. For each group, n = 3. *Significantly different compared with normal (P < 0.05).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In this study, we demonstrate the presence of a nonspecific cation channel in cultured skeletal muscle myotubes from hamsters. The basal activity of this channel was substantially higher in myotubes isolated from Bio14.6 hamsters, which lack the dystrophin-associated SG subcomplex. We found that channel activity was further enhanced by pressure-induced stretch of the membrane patch and that it was inhibited by a known blocker of other stretch-activated channels, GdCl3. Depolymerization of F-actin by cytochalasin D enhanced basal channel activity in both normal and Bio14.6 myotubes. This effect was further potentiated by the application of stretch. In addition, we found that Ca2+ influx was significantly higher in the Bio14.6 group compared with the normal group, suggesting that these patch phenomena also occur in intact cells.

Comparison of the mechanosensitive cation channels observed in this study and in other skeletal muscle preparations. In our study, we observed channels under a variety of experimental conditions. The unitary conductance ranged between 8 and 47 pS, depending on the nature of the predominant charge carrier. The unitary events under the various experimental conditions were presumably caused by activity of the same stretch-activated, nonspecific cation channel for the following reasons. First, the reversal potentials (extrapolated from the linear portion of the I-V relationships) for these channels differed from the calculated equilibrium potential, irrespective of the nature of the predominant charge carrier. This suggests that these channels selected poorly among different cations. Evidence for cation selectivity was obtained when NaCl in the pipette was replaced by sodium glutamate. The fact that the unitary conductance was unchanged strongly suggests that Cl- permeates poorly through this channel. Second, under all experimental conditions, channels were activated by pressure-induced membrane stretch and inhibited by a low concentration of the stretch-activated channel blocker, GdCl3. Third, in none of these conditions were channels blocked by nifedipine (1 µM) or alpha -bungarotoxin (50 nM), which are known to inhibit L-type Ca2+ currents and nicotinic acetylcholine receptor channels, respectively. Although the L-type Ca2+ channel has a unitary conductance similar to that of the cation channel we observed (25 pS with Ba2+ as the charge carrier) (10) and can be activated by membrane stretch (22), our data suggest that the channel observed in our study was not the L-type Ca2+ channel. As noted earlier, the channel reported here is not sensitive to nifedipine. In addition, the voltage dependency of activation is totally different between these channels (most of our recordings were performed at membrane potentials beyond the activation voltage of L-type Ca2+ channels).

The channel activity observed in our study had striking similarities to that which was recorded in cultured myotubes and acutely isolated flexor digitorum brevis fibers from mdx mouse (12). Similar to the channel we observed here, skeletal muscle from the mdx mouse has a channel with a respective single-channel conductance of 7 or 19 pS, with Ca2+ or Ba2+ as the charge carrier. These authors also reported that channel activity was strongly enhanced by membrane stretch. Furthermore, basal activities of these channels were significantly higher in both mdx mice and Bio14.6 hamsters. Thus there are reasons to believe that channels with similar electrophysiological characteristics are activated in these two different forms of muscular dystrophy. Confirmation of their identity will await further characterization of the mdx channel (i.e., unitary conductance when using various cations as well as their ability to be blocked by GdCl3).

Role of the SG-sarcospan subcomplex in the activity of mechanosensitive channels. Significant basal activity of mechanosensitive channels was observed in mdx mice (that lack dystrophin). This observation suggests that a strong correlation may exist between the activity of mechanosensitive channels and the lack of dystrophin. However, in addition to the lack of dystrophin and dystroglycan, these mice also have a greatly reduced content of (or lack of) the SG-sarcospan subcomplex (4, 5, 26, 27). Therefore, the specific role of the SG-sarcospan subcomplex remains unclear. Our data suggest that similar mechanosensitive channels exist in Bio14.6 hamsters, which lack the SG-sarcospan subcomplex but still retain significant levels of dystrophin and beta -dystroglycan (19, 20, 30). These data are, therefore, in support of the concept that the SG-sarcospan subcomplex is involved in the modulation of mechanosensitive cation channels. At present, it is not clear how this may take place. One possibility is that the SG-sarcospan subcomplex may be interacting with these channels via submembrane actin cytoskeleton. In support of this hypothesis, we found that disruption of membrane-associated F-actin with cytochalasin D substantially increased the channel activity in both normal and Bio14.6 groups. It has recently been reported that the actin-binding protein filamin directly binds to delta -SG (35). Another study revealed that filamin also binds to one of the voltage-gated ion channels and regulates the expression level of this channel by interaction with actin cytoskeleton (28). It is therefore possible that some intermediate protein that can bind actin cytoskeleton (such as filamin) may be involved in the interaction of SGs with stretch-activated channels. Another possibility is that disruption of some or all of the components of the DGC may lead to membrane deformation, causing channel activity by general (and nonspecific) stretch of the membrane. This possibility is less likely, though, since independent reports exist that membrane fragility (as judged by the pressure required to rupture the membrane patch) is unaffected in mdx mice (18). In addition, in another mouse model (dy/dy mice that lack laminin-2), membrane fragility is expected to be equally affected, yet this mouse muscle shows no enhanced mechanosensitive channel activity (12). This suggests that a specific mechanism may exist for the activation of these channels in the Bio14.6 hamster.

Possible mechanisms of cell damage occurring in Bio14.6 skeletal muscle. There are several possible mechanisms that may underlie cellular damage occurring in Bio14.6 skeletal muscle fibers. We observed a high basal activity of cation-selective channels in myotubes isolated from Bio14.6 hamsters. These channels are also permeable to Ca2+, which could provide a leak pathway for Ca2+ to enter the cell and thus to cause cellular damage. These ideas are consistent with our present results showing that 45Ca2+ influx is about twofold higher in Bio14.6 hamsters than in normal controls. Because these channels are mechanosensitive, membrane stretch (as would occur during muscle contractions and relaxations) could further exacerbate this process. Another possibility is that opening of these cation-selective channels may cause membrane depolarization and consequent opening of voltage-gated Ca2+ channels and, hence, Ca2+ overload and cellular damage. Yet another possibility is suggested by the report that a Ca2+-specific leak channel is activated by increased proteolysis in dystrophic myocytes, leading to cell Ca2+ overload (23, 37). The cellular mechanisms of cell damage observed in Bio14.6 hamsters remain to be elucidated.

Possible limitations of this study. Our patch-clamp data show an enhanced basal activity and mechanosensitivity of a stretch-activated cation channel in the SG-deficient hamster, Bio14.6. One should be aware of possible confounding factors that may impact the interpretation of these data. It is possible, for example, that the viscoelastic properties of membrane in the myotubes from the Bio14.6 group might be different and that the increased basal channel activity and increased response to pressure may not have been due to specific alterations in SG components (discrepancy of the mechanosensitivity was recently reported between whole cell and membrane patch in Xenopus oocytes) (41). Although this is an interesting alternative mechanism that can account for increased cation-selective channel activity in Bio14.6 myotubes, this is unlikely to be the sole reason for increased basal channel activity. Our data show that there is an enhanced Gd3+-sensitive Ca2+ uptake in intact myotubes from Bio14.6 hamsters, consistent with the idea that this influx occurred though the Gd3+-sensitive cation channels or that the activity of these channels was responsible for Ca2+ influx through a different pathway. If so, these data suggest that the increased channel activity under patch-clamp conditions was due not only to changes in membrane viscoelastic properties but rather originated from specific alterations in SG components.

In conclusion, a novel finding in our present study is that the resting activity of stretch-activated nonspecific cation channels is markedly elevated in Bio14.6 hamster myotubes, which lack the SG-sarcospan subcomplex in the membrane. Increased Ca2+ influx was also observed in intact myotubes from the Bio14.6 group. Our data indicate that intact submembrane cytoskeletal architecture, including DGC components and the actin cytoskeleton, is important to regulate the activity of this mechanosensitive cation channel. We propose that increased activity of these channels may ultimately cause Ca2+ overload, which contributes to the cell damage observed in this hamster model of muscular dystrophy.


    ACKNOWLEDGEMENTS

This work was supported by Special Coordination Funds from the Science and Technology Agency of Japan and Research Grant for Cardiovascular Diseases 9C-6.


    FOOTNOTES

We thank Dr. Bernardo Rudy for useful discussions.

Address for reprint requests and other correspondence: M. Shigekawa, Dept. of Molecular Physiology, National Cardiovascular Center Research Institute, Fujishiro-dai 5-7, Suita, Osaka 565-8565, Japan.

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.

Received 18 September 2000; accepted in final form 26 March 2001.


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Am J Physiol Cell Physiol 281(2):C690-C699
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