1Institut für Pharmakologie, Medizinische Universität Wien, A-1090 Vienna; and 2Institut für Zoologie, Universität Salzburg, A-5020 Salzburg, Austria
Submitted 12 January 2004 ; accepted in final form 19 March 2004
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ABSTRACT |
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muscle plasticity; myosin heavy chain expression; sodium channel expression
Whereas changes in the expression of contractile proteins and metabolic enzymes in the course of fiber type conversion are well described (for review, see Ref. 37), little is known about possible adaptations in the electrophysiological properties of skeletal muscle cells. Ion channels of surface and intracellular membranes are crucially involved in the control of muscle cell excitability and consequently of muscle contraction. Thus changes in the expression and/or function of ion channels during fiber type conversion would have major consequences in the physiology of muscle cells. Froemming et al. (17) found a decrease in the expression of the ryanodine receptor Ca2+ release channel isoform ryanodine receptor 1 during fast-to-slow fiber type conversion in rabbit skeletal muscle. Other authors (10) reported enhanced Na+ current density paralleled by a transient increase in the mRNA concentration of the Na+ channel -subunit during slow-to-fast fiber type conversion in rat skeletal muscle. Recently, the same group showed that enhanced chloride channel expression (38) as well as reduced stretch-activated Ca2+ channel expression (16) also are associated with slow-to-fast fiber type conversion. Taken together, these findings suggest that fiber type conversion in skeletal muscle does indeed involve changes in the expression of ion channels. However, to our knowledge, alterations in the functional parameters of ion currents in the course of fiber type conversion have not yet been reported. Such alterations would be represented by changes in the kinetics and/or the voltage dependency of current activation and/or inactivation.
In this study, we tested the hypothesis that fast-to-slow fiber type conversion affects the functional parameters of Na+ currents. We found that Na+ current inactivation properties were significantly altered in fast-to-slow fiber type-converted cells. These results suggest that fiber type conversion of skeletal muscle cells involves functional adaptations of their electrophysiological properties. Some of the data reported herein were published previously in abstract form (61).
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MATERIALS AND METHODS |
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Electrical stimulation of C2C12 cells was performed via custom-built platin/iridium plate electrodes that were integrated into culture dishes (3.5-cm diameter). Electrical pulses were generated by a computer with an analog-to-digital/digital-to-analog (A/D-D/A) converter card (ACL-6128; Distrelec, Vienna, Austria). Media of stimulated cultures were changed every day to avoid possible negative effects of enhanced cellular metabolism due to increased contractile activity.
N1E-115 neuroblastoma cells were propagated in Dulbecco's modified Eagle's medium containing 4.5 g/l glucose, 4 mM L-glutamine, 50 u/ml penicillin, and 50 µg/ml streptomycin with 10% fetal calf serum and incubated at 37°C and 5% CO2.
Immunofluorescence. The cell culture dishes were washed twice with phosphate-buffered saline (PBS) and fixed with 4% paraformaldehyde/PBS for 30 min. Next, cells were washed twice with PBS and incubated for 5 min in 50 mM NH4Cl/PBS to remove the remaining paraformaldehyde. After a single wash in PBS, cells were permeabilized for 5 min in 0.1% Triton/PBS and washed twice in PBS before nonspecific binding sites were blocked with 2% bovine serum albumin/PBS (blocking solution) for 45 min. Incubation with the primary antibody (1:200 dilution in blocking solution) lasted 90 min. Two different monoclonal mouse antibodies were applied: MY32 (Biomedica, Vienna, Austria) and NOQ7.5.4D (Chemicon, Hofheim, Germany). Antibody MY32 recognizes neonatal and all adult fast MHC isoforms, whereas antibody NOQ7.5.4D is specific for slow MHC (20). After incubation with the primary antibody, cells were washed three times with blocking solution. This was followed by 60-min incubation with the secondary antibody, Alexa Fluor 488-conjugated goat anti-mouse IgG (1:500 dilution in blocking solution; Eubio, Vienna, Austria), labeled with a fluorophore. After two washings, with blocking solution and PBS, respectively, cells were briefly exposed to ddH2O to prevent the formation of salt crystals. Next, cells were dried in air and then mounted onto the dishes with the use of mowiol (Sigma, Vienna, Austria) overlaid with coverslips. After drying overnight, these preparations were examined under a fluorescence microscope (Axiovert 135M; Carl Zeiss, Oberkochen, Germany). All of these procedures were performed at room temperature.
MHC electrophoresis and immunoblotting.
Protein was extracted from C2C12 cells with the use of a denaturing lysis buffer containing 10 mM Tris·HCl (pH 7.5), 50 mM NaCl, 30 mM sodium pyrophosphate, 50 mM NaF, 2 mM EDTA, 1% (vol/vol) Triton X-100, 1 mM phenylmethylsulfonyl fluoride, 0.1 mM sodium vanadate, 1 µg/ml pepstatin, 0.5 µg/ml leupeptin, and 3 µg/ml aprotinin. The 3.5-cm dishes were washed with ice-cold PBS, and cells were collected in 1 ml of fresh PBS with the use of a cell scraper. The cell suspension was centrifuged (1,4001,600 rpm, 4°C, 1 min), and the pellet was resuspended in 60 µl of lysis buffer and shaken for 30 min at 4°C. This was followed by another centrifugation step (14,000 rpm, 4°C, 15 min). The supernatant was collected, frozen in liquid nitrogen, and stored at 80°C. Protein concentrations were determined by using a bicinchoninic acid kit according to the manufacturer's protocol (Pierce, Rockford, IL). Protein samples were mixed with Laemmli buffer and then heat denatured at 95°C for 5 min. Protein (15 µg/sample) was electrophoretically resolved on polyacrylamide gels (7.5 or 12% separating gel, 4% stacking gel) containing SDS. Electrophoresis was performed at a constant voltage of 120 V for 2 h. The electrophoretically separated samples were transferred to a nitrocellulose membrane (Schleicher & Schuell, Gassel, Germany). The blots were blocked with 5% nonfat dry milk in TBST (1 mM Tris, 15 mM NaCl, 0.1% Tween). After blocking, the blots were probed with specific anti-MHC antibodies MY32 or NOQ7.5.4D at 1:500 dilution in blocking solution. After the reaction with the antibodies and being washed three times in TBST for 5 min, the nitrocellulose sheet was incubated with the goat anti-mouse horseradish peroxidase-linked secondary antibody NA931V (Amersham Biosciences, Little Chalfont, UK) at 1:50,000 dilution in blocking solution. The antigen-antibody complex was visualized by staining with enhanced chemiluminescence (ECL) solution (ECL detection kit; Amersham, Arlington Heights, IL). Specific signals revealed by ECL were analyzed by densitometry, and the ratios of intensities were calculated, with the signal of
-actin (monoclonal anti-actin antibody, clone AC-40; Sigma) serving as a reference for the signal of MHC.
Electrophysiology. Na+ currents from C2C12 cells were recorded with the whole cell patch-clamp technique. The cells were differentiated for 1519 days in the absence or in the presence of 25 nM Ca2+ ionophore A-23187 (Sigma). The ionophore was always applied 3 days after the cells had been incubated in differentiation medium. In cultures grown on Matrigel-coated dishes, most of the differentiated cells showed a longitudinal shape (myotubes). However, in all dishes, several cells with a spherical shape (myoballs) could be found. Myoballs were selected for the electrophysiological experiments. In contrast to the longitudinal cell shape of myotubes, the spherical shape of myoballs allowed proper voltage control of the cell membrane to be obtained in whole cell patch-clamp experiments.
Na+ currents were recorded at room temperature (22 ± 1.5°C) with an Axoclamp 200B patch-clamp amplifier (Axon Instruments, Union City, CA). Recording was begun 10 min after whole cell access was attained to minimize time-dependent shifts in gating. Pipettes were formed from aluminosilicate glass (AF150-100-10; Science Products, Hofheim, Germany) with a P-97 horizontal puller (Sutter Instruments, Novato, CA), heat polished on a microforge (MF-830; Narishige, Japan), and had resistances between 1 and 2 M
when filled with the recording pipette solution (105 mM CsF, 10 mM NaCl, 10 mM EGTA, and 10 mM HEPES, pH 7.3). Substitution of K+ with Cs+ in the pipette solution eliminated K+ currents. Voltage-clamp protocols and data acquisition were performed with pCLAMP 6.0 software (Axon Instruments) via a 12-bit A/D-D/A interface (Digidata 1200; Axon Instruments). Data were low-pass filtered at 2 kHz (3 dB) and digitized at 1020 kHz. Curve fitting was performed with the use of ORIGIN 5.0 software (MicroCal Software, Northampton, MA). Current-voltage (I-V) relationships were fit with the function Gmax·(x Vrev)·[1 (1/{1 + exp[(x V0.5)/K]})], where Gmax is the maximum conductance, Vrev is the reversal potential, V0.5 is the voltage at which half-maximum activation occurs, and K is the slope factor. Na+ current density was calculated by dividing the maximum peak current amplitude (normally 20 mV) of a cell by its membrane capacitance. Cell capacitance was estimated by integrating the area under the capacitive transient (31) elicited by a 20-ms voltage step from 120 to 80 mV that did not activate the channels. This area was then divided by the applied change in voltage (40 mV). Steady-state inactivation data were fit with the Boltzmann function 1/{1 + exp[(x V0.5)/K]}, where V0.5 is the voltage at which half-maximum inactivation occurs and K is the slope factor. The fractions of channels that could be inactivated by the process of slow inactivation were detected by measuring the smallest test pulse peak currents observed after 10-s inactivating prepulses between 40 and 10 mV (followed by 20-ms periods at 140 mV to allow for recovery from fast inactivation) in each experiment. Data from experiments performed to assess tetrodotoxin (TTX) sensitivity were fit with the two-site binding function y = Bmax1·x/(k1 + x) + Bmax2·x/(k2 + x), where Bmax1 and Bmax2 are the relative contributions of a TTX-sensitive and a TTX-resistant Na+ channel fraction, respectively. Variables k1 and k2 represent the respective 50% inhibitory constant (IC50) values. A bathing solution consisting of 140 mM NaCl, 2.5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES, pH 7.4, was used to obtain recordings. Some experiments were executed in low-Na+ bath solution containing 15 mM Na+ to minimize current amplitudes. In those experiments, we used the impermeant monovalent cation N-methyl-D-glucamine as a substitute for 125 mM Na+. Chemicals were purchased from Sigma. Rapid solution changes were performed with a DAD-8-VC superfusion system (ALA Scientific Instruments, Westbury, NY).
Experiments on N1E-115 neuroblastoma cells were performed according to the same experimental procedures described above, except that different solutions were used. The pipette solution for these experiments consisted of 100 mM CsF, 40 mM CsCl, 10 mM NaCl, and 10 mM HEPES, pH 7.3. The bathing solution consisted of 140 mM NaCl, 2 mM CaCl2, and 10 mM HEPES, pH 7.4. Data are expressed as means ± SE. Statistical comparisons were performed with two-tailed Student's unpaired t-tests. P < 0.05 was considered significant.
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RESULTS |
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Figure 1 shows representative immunofluorescence images of well-differentiated multinuclear C2C12 cells. The MHC was targeted by either a primary antibody specific for fast MHC isoforms (MY32) (Fig. 1A) or an antibody specific for the slow MHC isoform (NOQ7.5.4D) (Fig. 1B). Immunolabeling was obtained with both antibodies. Control cells that were treated only with the secondary antibody did not show any fluorescence signal when identical microscope settings were used. These experiments indicated that our cells contained both fast and slow MHC isoforms. Expression of fast and slow MHC isoforms was further confirmed by immunoblot analysis (Fig. 1C). Immunoreactive signals also were obtained with the anti-fast MHC MY32 antibody (Fig. 1C, left) or the anti-slow MHC NOQ7.5.4D antibody (Fig. 1C, right). Undifferentiated myoblasts did not show any signal.
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Prolonged depolarization (seconds to minutes) causes inactivation of Na+ channels from which the channels recover with multiple kinetic phases whose time constants range across several orders of magnitude, from tens of milliseconds to tens of seconds. These kinetic phases of recovery are summarized by the term "slow inactivation." Figure 5C shows that a striking difference can be observed between the voltage dependence of slow inactivation of control and ionophore-treated cells (25 nM). In ionophore-treated cells, the fraction of channels that were slowly inactivated (fractSI) by strongly depolarized voltages (40 to 10 mV) was markedly decreased compared with the corresponding fraction in control cells (Table 1). This indicates that the Na+ currents of ionophore-treated cells are less affected by, or more resistant to, the process of slow inactivation. V0.5 was independent of ionophore treatment, however.
To test whether ionophore treatment produces a shift in the expression of different Na+ channel isoforms, we compared the TTX sensitivities of Na+ currents in control and ionophore-treated (25 nM, 1216 days) cells. Figure 6, AC, shows that ionophore treatment significantly reduced TTX sensitivity. Two distinct populations of Na+ channels, a TTX-sensitive fraction and a TTX-resistant fraction, could clearly be separated by fitting the data with a two-site binding function (Fig. 6C). As shown in Table 2, the IC50 values of Na+ channel block by TTX of control and ionophore-treated cells were similar. This was true for both the TTX-sensitive channel fraction (IC50 20 nM) and the TTX-resistant channel fraction (IC50
12 µM). In contrast to the IC50 values, the respective relative contributions of the two channel fractions were significantly different from each other. Thus the TTX-sensitive Na+ channel fraction in control cells amounted to 84% of the total channel fraction, whereas the corresponding value in ionophore-treated cells was reduced to 52% (Table 2). Figure 6D shows two experiments involving RT-PCR of the adult skeletal muscle voltage-gated Na+ channel isoform (Nav1.4) and the cardiac voltage-gated Na+ channel isoform (Nav1.5) in control and ionophore-treated cells. In both experiments, ionophore treatment decreased and increased the amounts of PCR product of Nav1.4 and Nav1.5, respectively. This suggests downregulation of Nav1.4 expression and upregulation of Nav1.5 expression at the mRNA level.
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DISCUSSION |
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Our data demonstrate that under control conditions, well-differentiated C2C12 cells express both fast and slow MHC isoforms. This was shown in immunofluorescence and immunoblot experiments by using the anti-MHC antibodies MY32 and NOQ7.5.4D, which are specific for fast and slow MHC isoforms, respectively. These antibodies have been used extensively to discriminate between fast and slow MHC isoform expression in mammalian skeletal muscle cells (12). Our data support the findings of other investigators who have reported the presence of fast and slow MHC isoforms in C2C12 cells (2, 32, 52, 59). On the basis of these findings, C2C12 cells cannot be classified as pure fast- or slow-type skeletal muscle fibers but must be categorized as hybrids. The fast MHC isoforms may predominate (2).
Kubis et al. (27) showed that a modest but sustained rise in [Ca2+]i levels induced by 400 nM of the Ca2+ ionophore A-23187 caused marked fast-to-slow fiber type conversion in a rabbit primary skeletal muscle cell culture. This crucial study not only emphasized the importance of [Ca2+]i for phenotypic adaptations in mammalian skeletal muscle but also provided an easy method by which to induce fast-to-slow fiber type conversion in muscle cell cultures. This method allows phenotypic changes that occur during fiber type conversion to be studied in in vitro models.
In the present study, we used A-23187 to induce fast-to-slow fiber type conversion in murine C2C12 skeletal muscle cells. We found a strong decrease in the fast MHC isoform content and an increase in the slow MHC isoform content, probably because of down- and upregulation of fast and slow MHC isoform expression, respectively. These results strongly suggest that, as it does in rabbit muscle cell cultures (27, 30), A-23187 induces considerable fast-to-slow fiber type conversion in C2C12 cells. However, significantly lower (25 instead of 400 nM) concentrations of ionophore had to be used in our study to prevent deterioration of the cultures. Interestingly, at 12.5 nM unlike at 25 nM, A-23187 did not significantly affect MHC isoform expression, although it induced a slight rise in [Ca2+]i (Fig. 2). Obviously, this slight change in [Ca2+]i was not sufficient to induce considerable fast-to-slow fiber type conversion. Allen and Leinwand (1) reported contrary effects of A-23187 treatment on the expression of MHC isoforms in C2C12 cells. These authors reported upregulation of the expression of fast MHC isoforms, whereas we found strong downregulation. This discrepancy may be due to the much lower (25 instead of 400 nM) ionophore concentrations as well as to the longer (2 wk instead of 36 h) treatment durations used in our study. It is possible that the fast MHC genes are activated at high [Ca2+]i and during an early phase of adaptation (1) but eventually are downregulated as cells progressively shift toward a slower phenotype (27).
The incubation of cultures in medium containing a Ca2+ ionophore represents a rather artificial stimulus to trigger fast-to-slow fiber type conversion. In contrast, CLFES mimicking the firing patterns of slow motor neurons in vivo may provide a more physiological approach (53). We found that chronic CLFES had a similar effect of downregulation (Fig. 4) on the expression of the fast MHC isoforms as ionophore treatment. This is not surprising, because, similarly to A-23187, CLFES may modestly increase [Ca2+]i (8), which, in both experimental approaches, most likely represents the trigger for fast-to-slow fiber type conversion. In contrast to ionophore treatment, we did not observe obvious upregulation of slow MHC isoform expression with the use of electrical stimulation. This could be due to an insufficient stimulation period (1 wk).
Fast-to-slow fiber type conversion affects functional Na+ current parameters. After we found that long-term application of low A-23187 concentrations induced fast-to-slow fiber type conversion in C2C12 cells, we investigated the hypothesis that alterations in the functional parameters of ionic currents accompany fiber type conversions in skeletal muscle. We chose to investigate Na+ currents because these currents play a central role in determining basal functional properties of muscle cells. They are responsible for the rapid depolarization of the cell membrane, which forms the basis for the propagation of action potentials. C2C12 cells were selected for this study because the morphological and biochemical characteristics of this cell line closely resemble those of differentiated skeletal muscle (7, 33, 49, 52, 59). Moreover, C2C12 cells exhibit developmental regulation in the expression of voltage-gated ion channels much like they do in vivo and show TTX-sensitive Na+ currents indicating the expression of Nav1.4 (7, 11, 29, 60). Spherically shaped, differentiated cells (myoballs) were selected for the electrophysiological experiments. We think that the physiological properties of myoballs were similar to normal, longitudinally shaped myotubes because myoballs were multinuclear, occasionally showed spontaneous contractions, and exhibited immunofluorescence signals similar to those of myotubes (see Figs. 1B and 3C). Moreover, myoballs resemble the electrophysiological properties of differentiated myotubes (4), and in C2C12 cells, even unfused myoblasts were shown to produce developmentally regulated, voltage-gated ion channels that resembled those of intact skeletal muscle (7).
Several authors have investigated the effects of increased [Ca2+]i induced by the application of relatively high concentrations of Ca2+ ionophores on the expression of voltage-gated ion channels in different cell types. Hirsh and Quandt (25) found that treatment of N1E-115 neuroblastoma cells with 1 µM A-23187 for 2 days led to downregulation of Na+ channel expression. Furthermore, treatment of C2C12 cells with 0.5 µM of the Ca2+ ionophore ionomycin for as long as 6 h activated the expression of IRK1, an inwardly rectifying K+ channel (51). These reports suggested that as a result of the application of relatively high concentrations of a Ca2+ ionophore, altered [Ca2+]i affects the expression of voltage-gated ion channels both in neurons and in muscle cells.
In this study, we tested the effects of long-term (2 wk) treatment with a comparably low (25 nM) A-23187 concentration (inducing fast-to-slow fiber type conversion) on the functional parameters of Na+ currents in C2C12 cells. We found a slightly reduced voltage dependence of fast inactivation and a higher resistance to slow inactivation in Na+ currents of ionophore-treated cells (Table 1). To our knowledge, this study is the first in which significant alterations in functional parameters of ionic currents have been found in association with skeletal muscle fiber type conversion. This finding cannot be explained simply by fiber type conversion-induced changes in the expression level of skeletal muscle Na+ channels (10); it most likely is a result of the expression of different Na+ channel isoforms (discussed below). Importantly, these considerations imply that the level of plasticity of skeletal muscle cells (i.e., the potential to adapt to changes in functional demands) can be extended to functional adaptations of their electrophysiological properties (for review, see Ref. 37).
Control experiments were performed in nonmuscle neuroblastoma (N1E-115) cells to exclude the possibility of a nonspecific A-23187 effect that is not directly related to fast-to-slow fiber type conversion. In these cells, the identical (25 nM) ionophore concentration used for the C2C12 experiments did not alter the inactivation parameters of Na+ currents (Table 3). An even higher (100 nM) ionophore concentration also failed to produce any significant effect. These control experiments strongly suggest that the altered inactivation parameters of Na+ currents that we found in ionophore-treated C2C12 cells are connected to the process of fast-to-slow fiber type conversion.
Our finding that Na+ current inactivation can be altered by experimentally induced fiber type conversion is in contrast to the finding of Desaphy et al. (10), who induced slow-to-fast fiber type conversion by hindlimb unloading in rat skeletal muscle. These authors found no evidence for differences in Na+ current inactivation between normal and slow-to-fast-converted muscle fibers. This discrepancy may be due to differences in the electrophysiological techniques applied or to the use of muscle fibers of different animal species. Alternatively, the lack of an effect on Na+ current inactivation reported by Desaphy et al. (10) may be due to insufficient fiber type conversion. Only partial fiber type conversion may be inducible by hindlimb unloading for 13 wk, and this may not provide a sufficient stimulus to significantly affect the functional Na+ current parameters. Indeed, evidence that more complete fiber type conversion alters functional Na+ current parameters can be discovered in early studies that compared Na+ currents between muscle fibers isolated from either fast or slow skeletal muscle (13, 14, 4042). The authors of these studies reported differences in the voltage dependencies of inactivation processes that are similar to our findings in the present study. First, slow muscle fibers were more resistant than fast muscle fibers to the process of slow inactivation (41, 42). This is in accordance with the findings in our fast-to-slow fiber type-converted C2C12 cells (Table 1). Second, the steepness of the steady-state fast inactivation curves was significantly reduced in slow fibers, which also is in agreement with our data. However, we observed only a minor effect on fast inactivation, and therefore its significant functional importance may be doubtful. In studies by Ruff (40, 41), Na+ currents were inactivated at less negative potentials in slow than in fast muscle fibers. We could not induce a corresponding difference with fiber type conversion in our system (Table 1). This discrepancy might be due to the fact that we were able to generate only partial fast-to-slow fiber type conversion by A-23187 treatment, whereas in the work of Ruff, pure fast muscle fibers probably were compared with pure slow fibers.
The reason for different Na+ current inactivation properties in control and fast-to-slow converted skeletal muscle fibers is unknown. In principle, all factors that affect inactivation are possible candidates. These involve Na+ channel isoform expression, channel subunit expression, G proteins, calmodulin, phosphorylation, glycosylation, nitric oxide-dependent nitrosylation, and cytosolic ion concentrations. All of these factors might affect or modulate Na+ current inactivation in a fiber type-specific manner. In the present study, we tested the hypothesis that the expression of different Na+ channel isoforms with specific inactivation properties is altered in the course of fast-to-slow fiber type conversion. Na+ currents of differentiated C2C12 cells were previously reported to consist of a TTX-sensitive as well as a TTX-resistant component (7, 29), suggesting that both Nav1.4 and Nav1.5 are expressed. Our main finding that Na+ currents of fast-to-slow fiber type-converted cells showed higher resistance to slow inactivation is in accord with relative enhancement of the expression of Nav1.5, because this Na+ channel isoform is highly resistant to slow inactivation (35, 56, 57). Like other authors, we found a TTX-sensitive as well as a TTX-resistant Na+ channel fraction (Fig. 6, AC) in differentiated C2C12 cells. Moreover, we observed a highly significant increase in the TTX-resistant Na+ channel fraction of fast-to-slow fiber type-converted cells (Table 2), suggesting an enhanced contribution of Nav1.5 to the total Na+ current. This may be explained by a relative shift in Na+ channel expression from Nav1.4 toward Nav1.5. Accordingly, we found molecular biological evidence that Nav1.4 expression is downregulated, whereas Nav1.5 expression is upregulated, in converted cells (Fig. 6D). This finding is in agreement with Desaphy et al. (10), who induced fiber type conversion in the opposite (i.e., slow to fast) direction and found upregulation of Nav1.4. To summarize, these results strongly suggest that enhanced relative expression of Nav1.5 compared with Nav1.4 in fast-to-slow fiber type-converted cells accounts for the different inactivation properties observed. One might argue that this mechanism of adaptation in the course of fiber type conversion is limited to the cells that we studied, the murine cell line C2C12, which might not fully resemble the properties of adult skeletal muscle. Accordingly, in addition to being expressed in the heart, Nav1.5 was originally thought to be expressed in embryonic and denervated, but not adult innervated, skeletal muscle (19, 26, 54). However, more recent studies (3, 15) have provided strong evidence that Nav1.5 is expressed and functions in normal innervated adult skeletal muscle as well. Thus Fletcher et al. (15) detected significant Nav1.5 mRNA levels by performing RT-PCR in vastus lateralis muscle biopsy specimens obtained from healthy humans. The expression of Nav1.5 was reduced 20-fold in biopsy specimens obtained from patients susceptible to malignant hyperthermia, which resulted in a higher sensitivity of muscle twitches to TTX and thus had considerable functional consequences. In addition to being present in human skeletal muscle, TTX-resistant Na+ current also was present in normal adult murine and equine skeletal muscle (3), suggesting functional Nav1.5 expression. According to these studies, relative shifts in the expression of Nav1.5 compared with Nav1.4 may well represent an as yet unknown common mechanism by which to regulate cell excitability in skeletal muscle. Such shifts in Na+ channel subtype expression also may play a role in the manifestation of certain myopathies.
Finally, our Na+ current density measurements revealed no significant differences between control and Ca2+ ionophore-treated cells. This finding is in disagreement with the results of numerous other studies (34, 50) in which the investigators found a decrease in Na+ current densities caused by ionophore treatment. In our study, with the use of comparably low ionophore concentrations, upregulation of Nav1.5 seems to compensate for downregulation of Nav1.4, which may result in similar Na+ current densities.
Physiological and pathophysiological implications. Slow inactivation is thought to play an important role in membrane excitability and firing properties (43, 45). The main finding of the present study is that the Na+ currents of fast-to-slow fiber type-converted cells showed higher resistance to slow inactivation. This specific inactivation property may have important physiological consequences in that it may help to allow converted cells, in contrast to control cells, to fire action potentials continuously, because firing is less likely to be terminated by the process of slow inactivation.
Inheritable mutations that alter Na+ current slow inactivation have been identified in human skeletal muscle. These mutations underlie diseases such as hyperkalemic periodic paralysis and paramyotonia congenita (9, 21, 22). Altered inactivation properties of Na+ currents that accompany fast-to-slow fiber type conversion may change a muscle's susceptibility to be affected by these diseases. Accordingly, fast and slow skeletal muscles have been shown to exhibit different susceptibility to myotonia (55).
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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2. Artaza JN, Bhasin S, Mallidis C, Taylor W, Ma K, and Gonzalez-Cadavid NF. Endogenous expression and localization of myostatin and its relation to myosin heavy chain distribution in C2C12 skeletal muscle cells. J Cell Physiol 190: 170179, 2002.[CrossRef][ISI][Medline]
3. Beech J, Fletcher JE, Erwin K, and Lindborg SR. Comparison of sensitivity of sodium currents to tetrodotoxin in equine muscle specimens with that in murine and human muscle specimens. Am J Vet Res 61: 133138, 2000.[ISI][Medline]
4. Boldin S, Jager U, Ruppersberg JP, Pentz S, and Rudel R. Cultivation, morphology, and electrophysiology of contractile rat myoballs. Pflügers Arch 409: 462467, 1987.[ISI][Medline]
5. Bottinelli R and Reggiani C. Human skeletal muscle fibres: molecular and functional diversity. Prog Biophys Mol Biol 73: 195262, 2000.[CrossRef][ISI][Medline]
6. Buller AJ, Eccles JC, and Eccles RM. Interactions between motoneurones and muscles in respect of the characteristic speeds of their responses. J Physiol 150: 417439, 1960.[ISI][Medline]
7. Caffrey JM, Brown AM, and Schneider MD. Ca2+ and Na+ currents in developing skeletal myoblasts are expressed in a sequential program: reversible suppression by transforming growth factor beta-1, an inhibitor of the myogenic pathway. J Neurosci 9: 34433453, 1989.[Abstract]
8. Carroll S, Nicotera P, and Pette D. Calcium transients in single fibers of low-frequency stimulated fast-twitch muscle of rat. Am J Physiol Cell Physiol 277: C1122C1129, 1999.
9. Cummins TR and Sigworth FJ. Impaired slow inactivation in mutant sodium channels. Biophys J 71: 227236, 1996.[Abstract]
10. Desaphy JF, Pierno S, Léoty C, George AL Jr, De Luca A, and Camerino DC. Skeletal muscle disuse induces fibre type-dependent enhancement of Na+ channel expression. Brain 124: 11001113, 2001.
11. Deschênes I, Neyroud N, DiSilvestre D, Marbán E, Yue DT, and Tomaselli GF. Isoform-specific modulation of voltage-gated Na+ channels by calmodulin. Circ Res 90: E49E57, 2002.[CrossRef][ISI][Medline]
12. Dusterhoft S and Pette D. Satellite cells from slow rat muscle express slow myosin under appropriate culture conditions. Differentiation 53: 2533, 1993.[ISI][Medline]
13. Duval A and Léoty C. Comparison between the delayed outward current in slow and fast twitch skeletal muscle in the rat. J Physiol 307: 4357, 1980.[Abstract]
14. Duval A and Léoty C. Ionic currents in slow twitch skeletal muscle in the rat. J Physiol 307: 2341, 1980.[Abstract]
15. Fletcher JE, Wieland SJ, Karan SM, Beech J, and Rosenberg H. Sodium channel in human malignant hyperthermia. Anesthesiology 86: 10231032, 1997.[ISI][Medline]
16. Fraysse B, Desaphy JF, Pierno S, De Luca A, Liantonio A, Mitolo CI, and Camerino DC. Decrease in resting calcium and calcium entry associated with slow-to-fast transition in unloaded rat soleus muscle. FASEB J 17: 19161918, 2003.
17. Froemming GR, Murray BE, Harmon S, Pette D, and Ohlendieck K. Comparative analysis of the isoform expression pattern of Ca2+-regulatory membrane proteins in fast-twitch, slow-twitch, cardiac, neonatal and chronic low-frequency stimulated muscle fibers. Biochim Biophys Acta 1466: 151168, 2000.[ISI][Medline]
18. Galler S, Hilber K, Gohlsch B, and Pette D. Two functionally distinct myosin heavy chain isoforms in slow skeletal muscle fibres. FEBS Lett 410: 150152, 1997.[CrossRef][ISI][Medline]
19. George AL Jr, Komisarof J, Kallen RG, and Barchi RL. Primary structure of the adult human skeletal muscle voltage-dependent sodium channel. Ann Neurol 31: 131137, 1992.[ISI][Medline]
20. Harris AJ, Fitzsimons RB, and McEwan JC. Neural control of the sequence of expression of myosin heavy chain isoforms in foetal mammalian muscles. Development 107: 751769, 1989.[Abstract]
21. Hayward LJ, Brown RH Jr, and Cannon SC. Slow inactivation differs among mutant Na channels associated with myotonia and periodic paralysis. Biophys J 72: 12041219, 1997.[Abstract]
22. Hayward LJ, Sandoval GM, and Cannon SC. Defective slow inactivation of sodium channels contributes to familial periodic paralysis. Neurology 52: 14471453, 1999.
23. Hilber K and Galler S. Mechanical properties and myosin heavy chain isoform composition of skinned skeletal muscle fibres from a human biopsy sample. Pflügers Arch 434: 551558, 1997.[CrossRef][ISI][Medline]
24. Hilber K, Galler S, Gohlsch B, and Pette D. Kinetic properties of myosin heavy chain isoforms in single fibers from human skeletal muscle. FEBS Lett 455: 267270, 1999.[CrossRef][ISI][Medline]
25. Hirsh JK and Quandt FN. Down-regulation of Na channel expression by A23187 [GenBank] in N1E-115 neuroblastoma cells. Brain Res 706: 343346, 1996.[CrossRef][ISI][Medline]
26. Kallen RG, Sheng ZH, Yang J, Chen LQ, Rogart RB, and Barchi RL. Primary structure and expression of a sodium channel characteristic of denervated and immature rat skeletal muscle. Neuron 4: 233242, 1990.[ISI][Medline]
27. Kubis HP, Haller EA, Wetzel P, and Gros G. Adult fast myosin pattern and Ca2+-induced slow myosin pattern in primary skeletal muscle culture. Proc Natl Acad Sci USA 94: 42054210, 1997.
28. Kubis HP, Scheibe RJ, Meissner JD, Hornung G, and Gros G. Fast-to-slow transformation and nuclear import/export kinetics of the transcription factor NFATc1 during electrostimulation of rabbit muscle cells in culture. J Physiol 541: 835847, 2002.
29. Kubo Y. Comparison of initial stages of muscle differentiation in rat and mouse myoblastic and mouse mesodermal stem cell lines. J Physiol 442: 743759, 1991.[Abstract]
30. Meissner JD, Kubis HP, Scheibe RJ, and Gros G. Reversible Ca2+-induced fast-to-slow transition in primary skeletal muscle culture cells at the mRNA level. J Physiol 523.1: 1928, 2000.
31. Meza U, Avila G, Felix R, Gomora JC, and Cota G. Long-term regulation of calcium channels in clonal pituitary cells by epidermal growth factor, insulin, and glucocorticoids. J Gen Physiol 104: 10191038, 1994.[Abstract]
32. Miller JB. Myogenic programs of mouse muscle cell lines: expression of myosin heavy chain isoforms, MyoD1, and myogenin. J Cell Biol 111: 11491159, 1990.[Abstract]
33. Minty A, Blau H, and Kedes L. Two-level regulation of cardiac actin gene transcription: muscle-specific modulating factors can accumulate before gene activation. Mol Cell Biol 6: 21372148, 1986.[ISI][Medline]
34. Offord J and Catterall WA. Electrical activity, cAMP, and cytosolic calcium regulate mRNA encoding sodium channel alpha subunits in rat muscle cells. Neuron 2: 14471452, 1989.[ISI][Medline]
35. O'Reilly JP, Wang SY, Kallen RG, and Wang GK. Comparison of slow inactivation in human heart and rat skeletal muscle Na+ channel chimaeras. J Physiol 515: 6173, 1999.
36. Pette D and Staron RS. Cellular and molecular diversities of mammalian skeletal muscle fibers. Rev Physiol Biochem Pharmacol 116: 176, 1990.[Medline]
37. Pette D and Staron RS. Transitions of muscle fiber phenotypic profiles. Histochem Cell Biol 115: 359372, 2001.[ISI][Medline]
38. Pierno S, Desaphy JF, Liantonio A, De Bellis M, Bianco G, De Luca A, Frigeri A, Nicchia GP, Svelto M, Léoty C, George AL Jr, and Camerino DC. Change of chloride ion channel conductance is an early event of slow-to-fast fibre type transition during unloading-induced muscle disuse. Brain 125: 15101521, 2002.
39. Reggiani C, Bottinelli R, and Stienen GJM. Sarcomeric myosin isoforms: fine tuning of a molecular motor. News Physiol Sci 15: 2633, 2000.
40. Ruff RL. Na current density at and away from end plates on rat fast- and slow-twitch skeletal muscle fibers. Am J Physiol Cell Physiol 262: C229C234, 1992.
41. Ruff RL. Sodium channel slow inactivation and the distribution of sodium channels on skeletal muscle fibres enable the performance properties of different skeletal muscle fibre types. Acta Physiol Scand 156: 159168, 1996.[CrossRef][ISI][Medline]
42. Ruff RL, Simoncini L, and Stuhmer W. Comparison between slow sodium channel inactivation in rat slow- and fast-twitch muscle. J Physiol 383: 339348, 1987.[Abstract]
43. Ruff RL, Simoncini L, and Stuhmer W. Slow sodium channel inactivation in mammalian muscle: a possible role in regulating excitability. Muscle Nerve 11: 502510, 1988.[ISI][Medline]
44. Salmons S and Sreter FA. Significance of impulse activity in the transformation of skeletal muscle type. Nature 263: 3034, 1976.[ISI][Medline]
45. Sawczuk A, Powers RK, and Binder MD. Spike frequency adaptation studied in hypoglossal motoneurons of the rat. J Neurophysiol 73: 17991810, 1995.
46. Schiaffino S and Reggiani C. Myosin isoforms in mammalian skeletal muscle. J Appl Physiol 77: 493501, 1994.
47. Schiaffino S and Reggiani C. Molecular diversity of myofibrillar proteins: gene regulation and functional significance. Physiol Rev 76: 371423, 1996.
48. Schiaffino S and Serrano A. Calcineurin signaling and neural control of skeletal muscle fiber type and size. Trends Pharmacol Sci 23: 569575, 2002.[CrossRef][ISI][Medline]
49. Semsarian C, Sutrave P, Richmond DR, and Graham RM. Insulin-like growth factor (IGF-I) induces myotube hypertrophy associated with an increase in anaerobic glycolysis in a clonal skeletal-muscle cell model. Biochem J 339: 443451, 1999.[CrossRef][ISI][Medline]
50. Sherman SJ and Catterall WA. Electrical activity and cytosolic calcium regulate levels of tetrodotoxin-sensitive sodium channels in cultured rat muscle cells. Proc Natl Acad Sci USA 81: 262266, 1984.[Abstract]
51. Shin KS, Park JY, Kwon H, Chung CH, and Kang MS. Opposite effect of intracellular Ca2+ and protein kinase C on the expression of inwardly rectifying K+ channel 1 in mouse skeletal muscle. J Biol Chem 272: 2122721232, 1997.
52. Silberstein L, Webster SG, Travis M, and Blau HM. Developmental progression of myosin gene expression in cultured muscle cells. Cell 46: 10751081, 1986.[ISI][Medline]
53. Thelen MH, Simonides WS, and van Hardeveld C. Electrical stimulation of C2C12 myotubes induces contractions and represses thyroid-hormone-dependent transcription of the fast-type sarcoplasmic-reticulum Ca2+-ATPase gene. Biochem J 321.3: 845848, 1997.
54. Trimmer JS, Cooperman SS, Tomiko SA, Zhou JY, Crean SM, Boyle MB, Kallen RG, Sheng ZH, Barchi RL, Sigworth FJ, Goodman RG, Agnew SA, and Mandel G. Primary structure and functional expression of a mammalian skeletal muscle sodium channel. Neuron 3: 3349, 1989.[ISI][Medline]
55. Vergara C and Ramirez BU. Age-dependent expression of the apamin-sensitive calcium-activated K+ channel in fast and slow rat skeletal muscle. Exp Neurol 146: 282285, 1997.[CrossRef][ISI][Medline]
56. Vilin YY, Fujimoto E, and Ruben PC. A single residue differentiates between human cardiac and skeletal muscle Na+ channel slow inactivation. Biophys J 80: 22212230, 2001.
57. Vilin YY, Makita N, George AL Jr, and Ruben PC. Structural determinants of slow inactivation in human cardiac and skeletal muscle sodium channels. Biophys J 77: 13841393, 1999.
58. Wehrle U, Dusterhoft S, and Pette D. Effects of chronic electrical stimulation on myosin heavy chain expression in satellite cell cultures derived from rat muscles of different fiber-type composition. Differentiation 58: 3746, 1994.[CrossRef][ISI][Medline]
59. Weydert A, Barton P, Harris AJ, Pinset C, and Buckingham M. Developmental pattern of mouse skeletal myosin heavy chain gene transcripts in vivo and in vitro. Cell 49: 121129, 1987.[ISI][Medline]
60. Wieland SJ, Fletcher JE, and Gong QH. Differential modulation of a sodium conductance in skeletal muscle by intracellular and extracellular fatty acids. Am J Physiol Cell Physiol 263: C308C312, 1992.
61. Zebedin E, Sandtner W, Szendroedi J, Galler S, Todt H, and Hilber K. A possible link between myosin isoform expression and sodium channel function in skeletal muscle cells (Abstract). Naunyn Schmiedebergs Arch Pharmacol 367, Suppl 1: R6, 2003.