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
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ABSTRACT |
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Deficiency of -sarcoglycan
(
-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
-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
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INTRODUCTION |
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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, -dystroglycan forms the
membrane-spanning linkage between
-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 (
-,
-,
-, and
-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
-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 -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
-SG in Bio14.6 hamsters causes disruption of the DGC and almost complete loss of other SGs as well as
reduction of
-dystroglycan (16, 20, 30, 34). However,
dystrophin and
-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
-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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MATERIALS AND METHODS |
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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 M, 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. -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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RESULTS |
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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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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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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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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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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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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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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, -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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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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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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DISCUSSION |
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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
-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).
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 -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
-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 |
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This work was supported by Special Coordination Funds from the Science and Technology Agency of Japan and Research Grant for Cardiovascular Diseases 9C-6.
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FOOTNOTES |
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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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