Fiber type conversion alters inactivation of voltage-dependent sodium currents in murine C2C12 skeletal muscle cells

Eva Zebedin,1 Walter Sandtner,1 Stefan Galler,2 Julia Szendroedi,1 Herwig Just,1 Hannes Todt,1 and Karlheinz Hilber1

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


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Each skeletal muscle of the body contains a unique composition of "fast" and "slow" muscle fibers, each of which is specialized for certain challenges. This composition is not static, and the muscle fibers are capable of adapting their molecular composition by altered gene expression (i.e., fiber type conversion). Whereas changes in the expression of contractile proteins and metabolic enzymes in the course of fiber type conversion are well described, little is known about possible adaptations in the electrophysiological properties of skeletal muscle cells. Such adaptations may involve changes in the expression and/or function of ion channels. In this study, we investigated the effects of fast-to-slow fiber type conversion on currents via voltage-gated Na+ channels in the C2C12 murine skeletal muscle cell line. Prolonged treatment of cells with 25 nM of the Ca2+ ionophore A-23187 caused a significant shift in myosin heavy chain isoform expression from the fast toward the slow isoform, indicating fast-to-slow fiber type conversion. Moreover, Na+ current inactivation was significantly altered. Slow inactivation less strongly inhibited the Na+ currents of fast-to-slow fiber type-converted cells. Compared with control cells, the Na+ currents of converted cells were more resistant to block by tetrodotoxin, suggesting enhanced relative expression of the cardiac Na+ channel isoform Nav1.5 compared with the skeletal muscle isoform Nav1.4. These results imply that fast-to-slow fiber type conversion of skeletal muscle cells involves functional adaptation of their electrophysiological properties.

muscle plasticity; myosin heavy chain expression; sodium channel expression


ADULT SKELETAL MUSCLE has a remarkable capacity to adapt in response to altered functional demands such as enhanced activity or disuse. On the basis of contraction kinetics, a distinction is drawn between "fast" and "slow" skeletal muscles, which contain predominantly fast- and slow-type muscle fibers, respectively. These fiber types express specific sets of fast and slow protein isoforms (for review, see Refs. 5, 36, 47) and are generally categorized according to the specific myosin heavy chain (MHC) isoforms that they express. In adaptation to altered functional demands, muscle fibers can switch from the fast to the slow fiber type and vice versa (i.e., fiber type conversion). Fiber type conversion involves morphological and biochemical changes that result in altered contractile properties and endurance capacities. It is well established that motor neuron firing patterns control the expression of isoforms of contractile proteins and metabolic enzymes of muscle fibers in vivo (6, 44). These firing patterns have been mimicked successfully in vitro in cell culture studies by imposed electrical stimulation (28, 58). The cellular signaling mechanisms involved in muscle fiber type conversions are only beginning to be understood (for a recent review, see Ref. 48). However, changes in intracellular free Ca2+ concentration ([Ca2+]i) seem to be essentially involved. Importantly, Kubis et al. (27) showed that a modest but sustained rise in [Ca2+]i caused by low concentrations of the Ca2+ ionophore A-23187 in the culture medium induced fast-to-slow fiber type conversion in a rabbit primary skeletal muscle cell culture. This clearly emphasizes the importance of Ca2+ for phenotypic adaptations in skeletal muscle.

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 {alpha}-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).


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. The C3H murine skeletal muscle cell line C2C12 (CRL-1772; American Type Culture Collection, Manassas, VA) was propagated in growth medium consisting of Dulbecco's modified Eagle's medium (Invitrogen, Lofer, Germany) containing 4.5 g/l glucose, 4 mM L-glutamine, 50 U/ml penicillin, 50 µg/ml streptomycin, and 20% fetal calf serum (PAA Laboratories, Pasching, Austria). The cells were incubated at 37°C and 5% CO2, and when ~50–70% confluence was reached, undifferentiated muscle cells (myoblasts) were plated onto Matrigel (Becton Dickinson, Schwechat, Austria)-coated culture dishes (3.5 cm; Sarstedt, Wiener Neudorf, Austria). Differentiation was induced by serum reduction. For this purpose, myoblasts were incubated in differentiation medium that was identical to the growth medium, except that it contained 2% horse serum (Invitrogen) instead of 20% fetal calf serum. The media were changed three times per week. Before being used to coat the dishes, Matrigel was diluted 1:10 with differentiation medium without horse serum. Coating was performed by transferring 1 ml of this ice-cold, Matrigel-containing medium into the dishes via pipettes. Immediately afterward, most of the medium was removed again, leaving a thin (~70 µl) Matrigel medium layer coating the dishes, a procedure that greatly reduced the amount of Matrigel required. Matrigel coating considerably improved cell culture performance. Compared with uncoated or collagen-coated dishes, myoblast proliferation as well as the development of well-differentiated, longitudinally shaped, multinuclear muscle cells (myotubes) was greatly enhanced in Matrigel-coated dishes. Importantly, the long-term survival of myotubes also was improved. These myotubes could be kept for at least 3 wk without any obvious signs of deterioration.

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 (1,400–1,600 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 {beta}-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 15–19 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{Omega} 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 10–20 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[(xV0.5)/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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
MHC expression in control cells. Differentiation of C2C12 myoblasts was induced by serum reduction. Two to three days after the induction of differentiation, myoblasts began to fuse to each other and formed small multinuclear myotubes. After 5–6 days, large longitudinal myotubes containing tens of nuclei had appeared, and after 7–8 days, the cells had reached a well-differentiated state showing cross-striation. A few myotubes also showed spontaneous contractions. These differentiated myotube cultures could be maintained for at least 3 wk.

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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Fig. 1. Fast and slow myosin heavy chain (MHC) isoform expression in differentiated C2C12 cells. Fast (A) and slow (B) MHC isoforms were detected by immunofluorescence with the use of the monoclonal anti-MHC antibodies MY32 (anti-fast MHC) and NOQ7.5.4D (anti-slow MHC), respectively, and secondary Alexa Fluor 488 antibody. Shown are C2C12 cells that were differentiated for 22 days. Identical microscope settings were used in A and B. Bars, 40 µm. C: immunoblot analysis of fast (MHCf) and slow (MHCs) isoforms. The C2C12 myotubes (MT) were differentiated for 17 days. Undifferentiated myoblasts (MB) were used as control. Antibodies MY32 (left) and NOQ7.5.4D (right) were used to detect MHCf and MHCs isoforms, respectively.

 
Effects of Ca2+ ionophore A-23187 on MHC expression. To study the effect of a long-term rise in [Ca2+]i, after 3 days of differentiation, A-23187 was added to the differentiation medium of part of the dishes at a final concentration of either 12.5 or 25 nM, with the remainder of the dishes serving as control. The cells were then kept in this medium for 12–16 days, and no obvious differences regarding culture growth, differentiation, and cell morphology were observed between control and ionophore-treated cultures. Slightly higher ionophore concentrations (50 or 100 nM), however, led to marked deterioration of the cultures. The control experiment in Fig. 2 shows that the low ionophore concentrations (12.5 or 25 nM) applied in our experiments were sufficient to cause a significant sustained increase in [Ca2+]i. This is indicated by a rise in the fura 2 signal ratios (R340/380) at 340- and 380-nm excitation wavelengths after ionophore application.



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Fig. 2. Effect of A-23187 on intracellular free Ca2+ concentration ([Ca2+]i). Shown is the fura 2 signal ratio (R340/380) at 340- and 380-nm excitation wavelengths over time. C2C12 MT that had been differentiated for 18 days were loaded with 5 µM fura 2-AM (Calbiochem, Vienna, Austria) for 30 min. A microscope photometer (Axiovert 200; Carl Zeiss, Oberkochen, Germany) was used to obtain the optical measurement over segments of 2 single MT. The dye was successively excited with light at wavelengths of 340 and 380 nm. The emitted light was measured at 510 ± 10 nm, and R340/380, which is related to the [Ca2+]i of the MT, was calculated. Arrows indicate the start of MT superfusion at various A-23187 concentrations.

 
Figure 3, A–C, shows representative immunofluorescence images of C2C12 cells that were incubated in differentiation medium containing 25 nM A-23187. Immunolabeling was obtained with both the MY32 (Fig. 3A) and NOQ7.5.4D (Fig. 3, B and C) antibodies. This indicated that, like control myotubes, ionophore-treated myotubes express fast and slow MHC isoforms. In addition, Fig. 3C shows that a spherically shaped myoball exhibited immunolabeling similar to that of longitudinally shaped myotubes. When the fluorescence images in Figs. 1 and 3 are compared, a relative enhancement of the anti-slow MHC antibody signal due to ionophore treatment can be recognized. This effect was observed in all six experiments that we performed. A more quantitative analysis of MHC isoform expression in ionophore-treated cells was performed by immunoblotting. Figure 3D shows a marked decrease in the anti-fast MHC antibody signal due to incubation of the cells in 25 nM A-23187. In contrast, Fig. 3E shows that the anti-slow MHC antibody signal was increased. A-23187 (12.5 nM) did not produce any significant effects. A summary of the results of a series of such experiments is shown in Fig. 3F. The results suggest that long-term application of A-23187 in sufficiently high (25 nM) concentrations causes strong downregulation in the expression of the fast MHC isoforms as well as marked upregulation in the expression of the slow MHC isoform in C2C12 cells.



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Fig. 3. Effect of A-23187 on MHC isoform expression. Immunofluorescence staining of MHCf (A) and MHCs (B and C) was performed with the primary antibodies MY32 and NOQ7.5.4D, respectively, and the secondary Alexa Fluor antibody. Shown are differentiated C2C12 cells that were treated with 25 nM A-23187 for 12 (A and B) or 14 (C) days. Identical microscope settings were used in AC. Bars, 40 µm. D and E: immunoblot analysis of MHCf and MHCs isoforms was performed in C2C12 cells that had been differentiated for 17 days with antibodies MY32 (D) and NOQ7.5.4D (E). Actin signal was used as a loading control. Control cells (lane C) were compared with cells that had been treated with 25 nM (lane 1) or 12.5 nM (lane 2) of A-23187 for 14 days. F: comparison of the actin-corrected relative densities of immunoblot signals obtained in a series of experiments as described in D and E. The effect of 25 nM A-23187 is illustrated.

 
Chronic low-frequency electrical stimulation (CLFES) has been shown to induce fast-to-slow fiber type conversion in various skeletal muscle cell types (for review, see Ref. 37), probably also via a sustained, modest increase in [Ca2+]i (8). We tested the effects of CLFES on MHC isoform expression. For this purpose, cells were stimulated with two different pulse protocols for 1 wk. One pulse protocol (PP1) consisted of 10-min stimulation periods (10-ms pulses, 3 Hz, 7.5 V) followed by resting periods of 30 min; the other protocol (PP2) was continuous stimulation (10-ms pulses, 1 Hz, 7.5 V). These pulse protocols caused visible contractions in at least 50% of the cells. After stimulation, immunoblot analysis was performed. Similarly to ionophore treatment, both stimulation protocols caused a marked decrease in the anti-fast MHC antibody signal (Fig. 4) compared with control, most likely indicating downregulation of fast MHC isoform expression. However, unlike ionophore treatment, electrical stimulation did not result in an obvious increase in the anti-slow MHC antibody signal.



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Fig. 4. Effect of chronic low-frequency electrical stimulation (CLFES) on MHC isoform expression. Immunoblot analysis of MHCf and MHCs isoforms was performed on C2C12 cells that had been differentiated for 6 days and, except for control cells (C), were electrically stimulated afterward for 1 wk according to different pulse protocols (PP1 and PP2; see RESULTS). First, analysis of MHCf was performed with antibody MY32. Thereafter, the blot was reprobed for MHCs with antibody NOQ7.5.4D after being stripped with stripping buffer (62.5 mM Tris·HCl, pH 6.8, 2% SDS, 100 mM 2-mercaptoethanol). Actin signal was used as a loading control.

 
Effects of A-23187 on functional parameters of Na+ currents. Figure 5A shows a summary of the I-V relationships of control C2C12 cells and cells that were incubated in differentiation medium containing 25 nM A-23187 for 12–16 days. Typical original traces of inward currents elicited by various depolarizing voltage steps of a control cell are also shown (Fig. 5A, inset). The rapid kinetics of the current decay, as well as the fact that TTX blocked the currents (see below), confirmed that Na+ ions were the carriers of these currents. The I-V relationships were similar in control and ionophore-treated cells. Both the voltages at which half-maximum activation occurred (V0.5) and the reversal potentials (Vrev) were not significantly different under control and experimental conditions (Table 1). These results suggest that ionophore treatment does not affect the process of macroscopic Na+ current activation in C2C12 cells. In a separate set of experiments, the Na+ current densities of control and ionophore-treated cells were estimated. The respective values obtained were 54 ± 8 pA/pF (n = 15) and 44 ± 6 pA/pF (n = 15), which were not significantly different.



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Fig. 5. Effect of A-23187 on functional Na+ current properties. A: summary of current-voltage (I-V) relationships for control C2C12 cells ({circ}) and C2C12 cells that were treated with 25 nM A-23187 for 12–16 days ({blacksquare}). From a holding potential of –120 mV, voltage steps to potentials in the range between –90 and +20 mV were applied. Inset shows typical original traces of inward currents elicited by such depolarizing voltage steps in a control cell. To reduce current amplitude, a low (15 mM)-Na+ bath solution was used. B: plot showing voltage dependencies of steady-state fast inactivation of a typical control C2C12 cell ({circ}) and an ionophore-treated (25 nM, 14 days) cell ({blacksquare}). From a holding potential of –120 mV, 50-ms inactivating prepulses to potentials in the range between –100 and –40 mV were applied. Lines connecting the data points represent curve fits to a Boltzmann equation. Channel availability was assessed by 10-ms test pulses to –20 mV. Peaks of inward currents (see inset) elicited by these test pulses were plotted against prepulse voltage. Bathing solutions contained 140 mM Na+. C: summary of voltage dependencies of slow inactivation of control C2C12 cells ({circ}) and A-23187-treated cells ({blacksquare}; 25 nM, 12–16 days). Ten-second inactivating prepulses to potentials in the range from –120 to –10 mV were applied from a holding potential of –120 mV. Thereafter, the channels were allowed to recover from fast inactivation during a 20-ms period at –140 mV. Control experiments showed that at –140 mV, this time was long enough to allow complete recovery from fast inactivation (data not shown). Channel availability after the 10-s inactivating prepulse and the 20-ms recovery period was then assessed by 10-ms test pulses to –20 mV. The peaks of the inward currents (see inset) elicited by such test pulses were plotted against the prepulse voltage. Bathing solutions contained 140 mM Na+.

 

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Table 1. Parameters of Na+ current activation, fast inactivation, and slow inactivation of control and A-23187-treated (12–16 days) C2C12 cells

 
Brief depolarizations (≤50 ms) cause inactivation of Na+ channels from which these channels recover with a single kinetic phase whose time constant is on the order of a few milliseconds. This inactivation process is called Na+ channel "fast inactivation." Figure 5B shows a comparison of the voltage dependence of fast inactivation of a typical control cell with that of a typical ionophore-treated cell (25 nM). The relationship of the ionophore-treated cell is slightly less steep than that of the control. Statistical analysis of a series of such experiments revealed a significant increase in the slope factor (Boltzmann fit) of the voltage dependence of fast inactivation after ionophore treatment (Table 1), indicating reduced voltage dependence. In contrast, V0.5 was independent of ionophore treatment. These results suggest that ionophore treatment significantly affects the process of Na+ current fast inactivation in C2C12 cells.

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, 12–16 days) cells. Figure 6, A–C, 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 ~1–2 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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Fig. 6. Effect of A-23187 on Na+ channel isoform expression. A: Na+ currents elicited by 10-ms test pulses from –120 to –20 mV in the absence (bottom traces) or presence of various concentrations of tetrodotoxin (TTX; 1 nM, 3 nM, 10 nM, 30 nM, 100 nM, 300 nM, 1 µM, 3 µM, and 10 µM) in a control cell and an ionophore-treated cell (25 nM, 14 days). B: summary of TTX sensitivities of Na+ currents in control and ionophore-treated cells (25 nM, 12–16 days). Current peak amplitudes in the presence of TTX were normalized to the respective amplitudes in the absence of TTX and plotted against TTX concentration. C: plot of current inhibition by TTX in the experiment shown in A. The solid lines show free fits of data with 2-site (Bmax1 and Bmax2) binding function. Insets demonstrate the fitting parameters obtained when Bmax1 and Bmax2 are the relative contributions of a TTX-sensitive (Nav1.4) and a TTX-resistant (Nav1.5) Na+ channel fraction, respectively. k1 and k2 represent the respective 50% inhibitory constant (IC50) values in µM. D: semiquantitative RT-PCR of Na+ channel isoforms Nav1.4 and Nav1.5 (independent experiments, 1 and 2). Control (CTL) cells were compared with ionophore-treated cells (25 nM, 14 days). Total cell RNA was extracted with the use of the RNeasy extraction kit (Qiagen, Hilden, Germany) and reverse transcribed with AMV reverse transcriptase and oligo(dT) primers (first-strand cDNA synthesis kit; Roche, Basel, Switzerland). PCR primers were designed in the 3' untranslated regions of the transcripts, where the isoforms share almost no homology (Nav1.4: forward 5'-TGAAGATCCCGCCTCCTGA-3'; Nav1.4: reverse 5'-AGTTTGTCTTTGTCCTGGCTA-3'; Nav1.5: forward 5'-CGTGCCCTGTTGTATCCTG-3'; Nav1.5: reverse 5'-CCAGCCAGGGTTGCTCGA-3'). PCR amplification of cDNA (25 cycles; i.e., within linear range of this PCR) was performed with Expand High Fidelity PCR System (Roche) with both primer pairs (cDNA lanes). A parallel reaction was performed with isolated total cellular RNA, omitting the reverse transcription step to demonstrate the absence of contaminating genomic DNA (RNA lanes). PCR products were visualized on 2% agarose gel. The Nav1.5 signal is stronger than that of Nav1.4 in both control and ionophore-treated cells. However, this RT-PCR allows for comparison only within the same isoform and not between expression levels of 2 different isoforms. Different intensities of Nav1.5 and Nav1.4 bands reflect differences in the efficiency of the RT reaction and the PCR. In the bottom lanes, RT-PCR for {beta}-actin is shown to control for prepared cDNA.

 

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Table 2. Parameters of tetrodotoxin sensitivity of Na+ current in control and A-23187-treated (12–16 days) C2C12 cells

 
Effects of A-23187 on functional parameters of Na+ currents in N1E-115 cells. To test whether the reported effects of A-23187 on Na+ current inactivation in C2C12 cells were linked to muscle fiber type conversion or were simply caused by an unspecific mechanism, we also treated N1E-115 neuroblastoma cells with low concentrations (25 nM, 100 nM) of A-23187 for 12–16 days. We then investigated the voltage dependencies of fast and slow inactivation in control and ionophore-treated N1E-115 cells by using the same experimental protocols as described above. Table 3 lists the parameters of fast inactivation (V0.5, K) and slow inactivation (V0.5, fractSI) for control cells and ionophore-treated N1E-115 cells. All these parameters were similar and were not significantly different from each other. These results indicate that, unlike in C2C12 cells, ionophore treatment does not affect the inactivation properties of Na+ currents in N1E-115 cells.


View this table:
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Table 3. Parameters of Na+ current fast and slow inactivation of control and A-23187-treated (12–16 days) NIE-115 cells

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
A-23187 treatment causes fast-to-slow fiber type conversion. The expression of different MHC isoforms has been used extensively and has proved to be an appropriate marker for discriminating between the fast and slow contracting skeletal muscle fiber types (see, e.g., Refs. 36, 39, 46). In mammalian fast skeletal muscle, three different fast MHC isoforms (MHC IIa, MHC IIb, and MHC IId/x) have been found, whereas slow skeletal muscle expresses only a single slow MHC isoform (MHC I{beta}). However, there is also evidence for different MHC I isoforms in rabbit (18) and human (23, 24) skeletal muscle.

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 1–3 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, A–C) 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).


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by Austrian Fonds zur Förderung der Wissenschaftlichen Forschung Grant P15063 [GenBank] (to K. Hilber).


    ACKNOWLEDGMENTS
 
We thank Anton Karel for technical assistance and Boris Kovacic, Helmut Kubista, Oliver Kudlacek, Rene Ott, and Dagmar Stoiber for helpful scientific comments.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Hilber, Institut für Pharmakologie, Medizinische Universität Wien, Währinger Strasse 13A, A-1090 Vienna, Austria (E-mail: karlheinz.hilber{at}meduniwien.ac.at).

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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