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Department of Biochemistry, Cinvestav-IPN, AP 14-740, Mexico City, DF 07000, Mexico

	Abstract

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In skeletal muscle myogenesis, precursor cells or myoblasts fuse to form multinucleated cells (myotubes), which then further develop into functional muscle. We investigated if the inhibition of myogenesis by transforming growth factor-ß1 (TGF-ß1) and bone morphogenetic protein-2 (BMP-2) involve regulation of voltage-dependent Ca²⁺ channels. Primary cultured myoblasts were kept in fusion medium (0–6 days) in either the absence (control conditions) or the presence of 40 pM TGF-ß1 or 5 nM BMP-2. Subsequently, the developing myotubes were transferred to a growth factor-free recording solution, and subjected to whole cell patch-clamp experiments. At day 0, 14% of non-fusing myoblasts exhibited T-current, whereas the L-current was practically absent. Under control conditions, however, the percentage of T- and L-channel-expressing myotubes increased sharply, from 25% at day 1 to 100% at days 2–6. In addition, parallel increases were determined for Ca²⁺-currents density and cell membrane capacitance (C_m), which is proportional to the size of myotubes. Interestingly, at days 1–2 TGF-ß1 and BMP-2 eliminated the T-current on initial 14% of T-channel-expressing myoblasts. Moreover, at day 6 the growth factors significantly reduced the maximal values of both T-current density (80%) and C_m (60%). The effect of BMP-2 was selective on T-channels, whereas TGF-ß1 decreased also the L-current density (90%). A similar reduction in maximal conductance of the Ca²⁺ channels was determined, in the absence of significant alterations in other essential properties of the channels, including the time course and voltage dependence of activation and inactivation. The results suggest these growth factors markedly reduce the number of functional T- (both TGF-ß1 and BMP-2) and L-channels (only TGF-ß1) in the surface of the plasma membrane, and contribute to explaining the associated effects on myogenesis.

(Received 20 April 2004; accepted after revision 14 June 2004; first published online 24 June 2004)
Corresponding author G. Avila: Department of Biochemistry, Cinvestav-IPN, AP 14-740, Mexico City, DF 07000, Mexico. Email: gavila{at}mail.cinvestav.mx

	Introduction

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The influx of Ca²⁺ through voltage-dependent Ca²⁺ channels underlies many physiological processes. Developing mammalian skeletal muscle cells express both L-type and T-type voltage-dependent Ca²⁺ channels (Cognard et al. 1986; Beam & Knudson, 1988a; Dirksen & Beam, 1995; Strube et al. 2000). The principal subunits of these channels are _1S or Ca_v1.1 (L-channels) (Tanabe et al. 1987, 1988) and _1H or Ca_v3.2 (T-channels) (Bijlenga et al. 2000; Berthier et al. 2002). In contrast to the well-established role for the L-channel _1S subunit in transmitting the electrical stimulus, resulting in excitation–contraction (EC) coupling (Tanabe et al. 1988; for reviews see Melzer et al. 1995; Dirksen, 2002), the physiological relevance of skeletal muscle T-channels is just beginning to be resolved. In skeletal muscle, significant levels of T-channels are normally expressed during prenatal development (Strube et al. 2000; Berthier et al. 2002) and then practically disappear 3–4 weeks after birth (Beam & Knudson, 1988b; Gonoi & Hasegawa, 1988). A similar transient expression of T-channels is also observed in developing myocytes in vitro (Cognard et al. 1986; Caffrey et al. 1989). Thus, the physiological role of skeletal muscle T-channels is thought to be tightly related to the skeletal muscle development (Beam & Knudson, 1988b; Perez-Reyes, 2003).

In developing skeletal muscle, precursor cells or embryonic myoblasts proliferate and then withdraw from the cell cycle to fuse, forming an ordered array of multinucleated cells, termed myotubes, which then further develop into functional muscle (for recent reviews see Chen & Goldhamer, 2003; Parker et al. 2003; Horsley & Pavlath, 2004). In response to muscle injury, quiescent satellite cells are activated to enter the cell cycle and to regenerate a pool of proliferating myogenic precursors, analogous to the embryonic myoblasts. Therefore, a parallelism has been suggested between adult muscle regeneration and embryonic myogenesis (Chen & Goldhamer, 2003). The fusion of myoblasts and the subsequent myotube formation is a Ca²⁺-dependent process (Shainberg et al. 1969), requires an increase on intracellular Ca²⁺ (David et al. 1981), and depends on Ca²⁺ influx through T-channels (Bijlenga et al. 2000; reviewed by Horsley & Pavlath, 2004).

Bone morphogenetic protein-2 (BMP-2) and transforming growth factor-ß1 (TGF-ß1) are members of the transforming growth factor-ß (TGF-ß) superfamily. At least 20 members of this superfamily have been discovered in mammals. These growth factors are structurally related and are essential in mammalian reproduction and development (Matzuk, 1995). Their central role on mammalian development is emphasized by the fact that the majority of ‘knock-out’ mice for TGF-ß1 (Shull et al. 1992; Kulkarni et al. 1993) and BMP-2 (Zhang & Bradley, 1996) die during embryogenesis. In regard to myogenesis, the continuous exposure of myoblasts to TGFß-1 or BMP-2, significantly decreases myotube formation and suppresses the expression level of myogenic transcription factors (Florini et al. 1986; Olson et al. 1986; Massague et al. 1986; Katagiri et al. 1994, 1997). We investigated here if these inhibitory effects on myogenesis, by TGFß-1 and BMP-2, on primary cultured myoblasts, involve alterations in the functional expression of voltage-gated Ca²⁺ channels. A chronic exposure (up to 6 days) of myoblasts to either TGFß-1 (40 pM) or BMP-2 (5 nM) significantly prevented myoblast fusion, as determined by measurements of cell membrane capacitance (C_m). Interestingly, the T-current density was reduced 80% by both TGFß-1 and BMP-2, whereas the L-current density was only significantly inhibited (90%) by TGFß-1. We determined a similar extent of inhibition for the Ca²⁺ channels maximal conductance or G_max. However, the voltage dependence and time course for the activation and inactivation of the channels were practically unaltered, suggesting these effects cannot be explained by alterations in the functional properties of the channels, but might probably involve a reduced number of functional channels in the surface of the membrane.

	Methods

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Primary culture of myocytes

Skeletal myocytes were obtained from 1- to 3-day-old newborn mice, following a slightly modified method of Beam & Franzini-Armstrong (1997). Briefly, the mice were decapitated, according to local ethical guidelines, and limb muscle minced and incubated in a calcium–magnesium-free Ringer solution, containing 0.3% trypsin and 0.01% DNAse (45 min, 37°C). The digested tissue was transferred to ‘plating medium’, consisting of Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% horse serum (HS), 10% fetal bovine serum (FBS), 100 U ml^–1 penicillin, 100 µg ml^–1 streptomycin, and 4 mML-glutamine. Subsequently, the tissue was dispersed by trituration, using fire-polished and silicone-coated Pasteur pipettes of decreasing tip diameter, centrifuged, resuspended, and filtered through a stainless steel mesh. The resulting cells (myoblasts and fibroblasts) were pre-plated for 2 h in a 10 mm plastic dish, and mildly resuspended, to selectively eliminate the most adhesive cells, corresponding to fibroblasts. The purified myoblasts were subsequently plated in 35 mm primary-treated plastic dishes, containing 2 ml of plating medium (0.4 x 10⁵ cells per dish). Twenty-four hours later, the plating medium was replaced by ‘fusion medium’, which was similar to the plating medium but only contained 2% HS, instead of 10% HS and 10% FBS. Unless specified, experiments were carried out on myocytes cultured 6 days in standard fusion medium (control condition) and fusion medium supplemented with either 5 nM BMP-2 or 40 pM TGF-ß1. The cells were kept at 37°C in a water-saturated tissue culture incubator, in the presence of 95% air and 5% CO₂. Fusion media were renewed every other day, to avoid loss of active nutrients and growth factors.

Voltage-clamp experiments

The myocytes were transferred from the culture medium to a growth factor-free external solution (see Recording solutions), and between the following 15 and 120 min, subjected to the whole cell patch-clamp technique (Hamill et al. 1981). The patch electrodes were elaborated from borosilicate glass capillaries using a two-stage vertical puller (L/M-3P-A, List-Electronic; Darmstadt/Eberstadt, Germany). The electrodes were filled with the internal solution described below (Recording solutions), and exhibited electrical resistances of 1.3–1.8 M, following immersion in the external solution. The holding potential was set to –80 mV, unless otherwise specified. The linear capacitative currents associated with charging the membrane from –80 mV to –100 mV were analysed to estimate the cells membrane capacitance (C_m). Briefly, 25 ms hyperpolarizing pulses were applied under cell-attached (A) and whole-cell (B) conditions. The associated capacitative currents were subtracted off-line (B – A), integrated, and divided by –20 mV. Once the C_m values were determined, the capacitative currents were > 90% analogically cancelled by the membrane capacitance cancellation feature of the patch-clamp amplifier (Axopatch 200B, Axon Instruments; Union City, CA, USA). All experiments were carried out under phase contrast microscopy (at 600x magnification), using an inverted microscope (TE-300, Nikon Instruments Inc.; Melville, NY, USA). This allowed us to visually corroborate the inhibitory effects of TGF-ß1 and BMP-2 on myotube formation, by directly observing a reduced number of nuclei per myocyte, as well as a smaller size of myocytes. Thus, although cell geometry might contribute to intrinsic variability of C_m within a particular experimental condition, significant differences across the mean values of C_m are primarily due to alterations in myoblasts fusion. An on-line subtraction procedure (P/–4) was used to eliminate all remaining linear components. The time constant for charging the membrane was in the range of 80 ± 30 µs, following analogical compensation of the series resistance (R_s). The current signals were filtered at 2 kHz with a 4-pole Bessel filter, and recorded at 100 kHz (capacitative currents) and 10 kHz (Ca²⁺ currents). We used a conditioning pre-pulse (1 s, –20 mV) to dissociate the contribution of T- and L-channels to the total Ca²⁺ current, as previously described (Avila et al. 2001). A 25 ms interpulse to –50 mV separated the test pulse and pre-pulse. To attenuate the tail current amplitudes and thus decrease the voltage-drop error associated with large currents, the membrane potential at the end of the test pulses was set to –50 mV for 150 ms. The absolute amplitudes for the T- and L-currents were normalized to the cells membrane capacitance (pA pF^–1), plotted independently as a function of test potential (I–V curves), and fitted according to the following Boltzmann equation:

(1)

where G_max is the maximal channel conductance, V_m is the test potential, V_rev is the extrapolated reversal potential, V is the potential for half-maximal activation of G_max, and k is a slope factor. To investigate the L-channels activation rate, the L-current activation phase was fitted according to a second order exponential equation:

(2)

where t represents the time after the onset of the test pulse, A_fast and A_slow are the contributions of a fast and a slow component to the total current amplitude, and _fast and _slow are the time constants associated with each component (for details see Avila & Dirksen, 2000). The T- and L-channels' voltage dependence of inactivation was accessed by modifying the holding potential (HP) 30–60 s before applications of test pulses to –20 mV and +40 mV. Subsequently, the absolute T- (–20 mV) and L-current (+40 mV) amplitudes were plotted as a function of the HP, and fitted according to:

(3)

I_max represents the maximal current amplitude, V the potential for half-maximal inactivation of I_max, and k is a slope factor.

Recording solutions

The bath or external solution contained (mM): 145 TEA-Cl, 10 CaCl₂ and 10 Hepes. The internal solution consisted of (mM): 145 caesium aspartate, 10 Cs₂EGTA, 5 MgCl₂ and 10 Hepes. The pH was adjusted to 7.4 with either TEA-OH (external solution) or CsOH (internal solution). All data were acquired and analysed with the pCLAMP software suite (v. 7.0, Axon Instruments), and are expressed as the mean ±S.E.M. Significant differences were determined at the P < 0.05 level, unless specified, using Student's unpaired t test.

	Results

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TGF-ß1 and BMP-2 significantly decrease myotube formation and Ca²⁺ currents

High serum-containing medium (plating medium) promotes myoblast proliferation and strongly inhibits myotube formation. However, myoblasts start to fuse after they are transferred to a low serum-containing medium (‘fusion medium’). In order to evaluate the capability of myoblasts to fuse under our experimental conditions, we estimated the membrane capacitance (C_m), which is proportional to the surface area of the plasma membrane (Fig. 1). The control values of C_m started to increase within the first 24 h in fusion medium, continued increasing and tended to reach a maximum value between days 4 and 6. On average, the control C_m values increase 8-fold from days 0 to 6 (Fig. 1B; filled symbols). However, the average C_m values obtained from BMP-2- and TGF-ß1-treated cells started to increase only after 3–4 days in fusion medium, and by day 6 were 65% smaller, compared to the respective control condition (Fig. 1A and B). We conclude therefore that the myoblasts were actively fusing in our control conditions, and both BMP-2- and TGF-ß1 severely decreased myotube formation.

View larger version (18K): [in this window] [in a new window]   Figure 1.  A chronic exposure of skeletal myocytes to BMP-2 and TGF-ß1 significantly decreases myotube formation A, representative capacitative currents associated with charging the cells plasma membrane from –80 mV to –100 mV. The current traces were obtained from myotubes cultured 6 days in control fusion medium (left, •) and fusion medium supplemented with 5 nM BMP-2 (centre, ) or 40 pM TGF-ß1 (right, grey circles). The estimated membrane capacitances (Cm) for these myotubes were 85 pF (control), 30 pF (BMP-2), and 34 pF (TGF-ß1). B, average Cm values for myocytes cultured 0–6 days in the control fusion medium (•) and fusion medium supplemented with BMP-2 () or TGF-ß1 (grey circles). The Cm values were estimated from capacitative currents as in A, and represent the means ±S.E.M., from a minimum of six (TGF-ß-1, 1 day) to a maximum of 36 (control, 6 days) experiments.

As a first test to determine if the inhibitory effects of BMP-2 and TGF-ß1 on myotube formation might involve a parallel regulation in the development of voltage-gated Ca²⁺ channels, we compared the amplitude of Ca²⁺ currents at –20 mV and +40 mV (Fig. 2A). At –20 mV, the Ca²⁺ currents exhibited all the hallmarks of a T-current, including fast activation and inactivation (Dirksen & Beam, 1995; Perez-Reyes, 2003). Accordingly, we evaluated the T-channel activity as the peak amplitude of the current traces at –20 mV (Fig. 2B). By contrast, the Ca²⁺ current at +40 mV activated slowly and did not inactivate, strongly suggesting it is mainly carried through L-channels (Tanabe et al. 1988). The L-channel activity was therefore evaluated as the amplitude of the current remaining at the end of the 200 ms, at +40 mV (Fig. 2C). The results of applying this protocol to 36 control, 27 BMP-2- and 13 TGF-ß1-treated myotubes (6 days) are plotted in Fig. 2B and C. As can be seen, both growth factors drastically decreased the T-current amplitude, to 5% of the control condition. By contrast, the L-current amplitude was differentially affected by TGF-ß1 and BMP-2. Specifically, the L-current practically disappeared in the TGFß-1-treated myotubes, but was only 60% reduced in response to BMP-2 (Fig. 2C).

View larger version (22K): [in this window] [in a new window]   Figure 2.  A 6-days treatment of myotubes with BMP-2 and TGF-ß1 severely decreases the Ca2+ current amplitude at –20 mV and +40 mV A, representative Ca2+ currents from myotubes cultured 6 days under control conditions (top) and in the presence of BMP-2 (middle) or TGF-ß1 (bottom). Ca2+ currents were elicited in response to 200 ms test pulses to –20 mV and +40 mV, which were delivered from a holding potential of –80 mV. B and C, average Ca2+ current amplitudes obtained from 36 control (filled bars), 27 BMP-2-treated (open bars), and 13 TGF-ß1-treated myotubes (grey bars). The current amplitudes were measured from current traces recorded as in A, and represent the peak of the current at –20 mV (B) and the current remaining at the end of the 200 ms at +40 mV (C).

It was recently proposed that Ca²⁺ influx through the T-channels stimulates the fusion index, defined as the number of nuclei in myotubes divided by the total number of nuclei counted (Bijlenga et al. 2000). If that is the case, then functional T-channels should be expressed at or before myoblast fusion. Figure 3 shows experimental data in support of this view. We estimate that 10 pA is the minimal current amplitude which can be unambiguously resolved in the present recording conditions. By taking into account this limitation, we classified the investigated cells, as expressing (> 10 pA) and not expressing ( 10 pA) Ca²⁺ currents. As a result, we detected T-channel activity in 2 out of 14 (i.e. 14%) non-fusing myoblasts, which means in myoblasts that were kept in plating medium for 24 h (day 0; Fig. 3A and B). The proportion of control cells expressing T-current exhibited a 2-fold increase at day 1 (i.e. from 14% to 25%), and by days 2–6, virtually all control myotubes expressed T-currents (Fig. 3B, filled circles). Likewise, the control T-current amplitude (Fig. 3C) and current density (Fig. 3D) increased as a function of the number of days in fusion medium. A very interesting observation is that the T-currents from original 14% subpopulation of T-current-expressing myoblasts (at day 0) practically disappeared following 1 and 2 days' exposures to either TGF-ß1 or BMP-2 (Fig. 3B). In the case of TGF-ß1, the abolition of T-currents was extended until day 3. Eventually, a significant proportion of the treated myotubes (50%; day 6) technically expressed T-channels, but the current amplitude remained barely resolved (Fig. 3B and C). Figure 3D shows also T-current amplitudes divided by the respective values of C_m (T-current density). It is clear that the growth factors strongly restrained development of the T-current density. In particular, significant differences were determined at days 3–6, between control myotubes and myotubes treated with BMP-2 or TGF-ß1. These results indicate that the T-channel activity is drastically inhibited (80%) in response to a long-term exposure of myotubes to BMP-2 and TGF-ß1.

View larger version (32K): [in this window] [in a new window]   Figure 3.  Onset of the inhibitory actions of BMP-2 and TGF-ß1 on the development of T-channels A, representative T-currents from myotubes that were kept 24 h in plating medium (i.e. non-fusing conditions; day 0, top), or 24 h in plating medium plus 1–6 days in fusion medium (days 1–6). The vertical scale at day 0 was 4-fold amplified, for clarity. B–D, time course for the development of T-current-expressing myocytes (B), T-current amplitude (C), and T-current density (D); for control myocytes (•) and myocytes treated with BMP-2 () or TGF-ß1 (grey circles). The current density (D) was estimated by dividing the current amplitudes in C by the corresponding Cm values reported in Fig. 1. The experiments shown in C for day 6 correspond to the same data set as in Fig. 2B.

The L-channel time course of development is presented in Fig. 4. In contrast to the T-current, which is present in 14% of non-fusing myoblasts (day 0; Fig. 3A and B), the L-current is practically absent in all myoblasts at day 0 (Fig. 4A and B). One day thereafter, the L-current was expressed in a 25% of the control cells, and by days 3–6, practically all control myotubes expressed this current (Fig. 4B, filled circles). Figure 4C shows the average L-current amplitudes from days 0 through 6. In general, TGFß-1 and BMP-2 tended to decrease L-current amplitude at every developmental stage. Significant differences, however, were only resolved for days 3–6. To determine if these effects on the L-current amplitude could be explained by parallel changes in the size of the myocytes, the L-current amplitudes were normalized by the corresponding values of C_m (L-current density; Fig. 4D). Interestingly, the L-current density was not significantly altered by BMP-2, at most of the developmental stages, but was practically eliminated by TGF-ß1. Taken together, the results from Figs 3 and 4 show a differential inhibition on the activity of T- (by BMP-2 and TGFß-1) and L-channels (by only TGFß-1).

View larger version (31K): [in this window] [in a new window]   Figure 4.  Onset of the inhibitory actions of BMP-2 and TGF-ß1 on the development of L-channels A, representative L-currents recorded from control myocytes at different stages of development. For clarity, the current trace at day 0 is been amplified 20-fold, in the vertical scale (note different calibration bar). The L-current was totally absent at day 0 (i.e. 10 pA). B–D, percentage of myocytes expressing L-current (B), average L-currents amplitude (C), and average L-current density (D), plotted as a function of the days in fusion medium. To estimate the L-current density (D), the L-current amplitudes (C) were normalized by the corresponding Cm values from Fig. 1. Experiments shown in Fig. 2C are also illustrated in C (day 6).

We next investigated if the effects of BMP-2 and TGFß-1 on the activity of Ca²⁺ channels could be explained by alterations in the macroscopic kinetic properties of the T- and L-channels. Figure 5A illustrates normalized T-currents, which were obtained from a control myotube (top) and myotubes treated with BMP-2 (middle) and TGF-ß1 (bottom). The time course of the current traces was very similar in the three experimental conditions, strongly suggesting the activation and inactivation rates of the T-channels have not been altered. Accordingly, neither the time to peak (Fig. 5B) nor the time constant for the inactivation of the current (_ina; Fig. 5C) were significantly modified.

View larger version (28K): [in this window] [in a new window]   Figure 5.  Neither BMP-2 nor TGF-ß1 significantly alters the time course for the activation and inactivation of T-channels A, normalized T-currents from myotubes cultured 6 days in control conditions (top) and in the presence of BMP-2 (middle) or TGF-ß1 (bottom). The current traces were amplified 3.7-fold (BMP-2) and 6-fold (TGF-ß1) to match the peak amplitude of the control trace. B and C, average T-current time to peak (B) and time constants for the inactivation of T-channels (ina, C). The values of ina were estimated by fitting the decreasing phase of T-currents according to the following equation: I=Imax[exp(–t/ina)]. t represents the time after the onset of depolarization, Imax is the extrapolated T-current amplitude at t= 0, and ina is the time constant for T-channel inactivation.

View larger version (25K): [in this window] [in a new window]   Figure 6.  The L-channels activation rate is not significantly altered by BMP-2 and TGF-ß1 A, normalized L-currents recorded from control (top), BMP-2- (middle), and TGF-ß1-treated (bottom) myotubes. The current traces were elicited following a 1 s inactivating prepulse to –20 mV, and multiplied by 1.95 (BMP-2) and 5.60 (TGF-ß1). For clarity, the tail currents are not fully displayed. B and C, average values for kinetic parameters describing the L-channels activation rate. A second order exponential equation (eqn (2)) was used to fit the activation phase of current traces recorded as in A. The resulting average values for Aslow, Afast (B), slow and fast (C), were obtained from a total of nine control (filled bars), eight BMP-2- (open bars) and seven TGF-ß1-treated (grey bars) myotubes. The relative values of Aslow (Relative Aslow) and Afast (Relative Afast) were estimated according to: Relative Aslow=Aslow/Aslow+Afast, and Relative Afast=Afast/Aslow+Afast.

The voltage-dependent activation of L- and T-channels is presented in Fig. 7. Families of current traces and the corresponding current to voltage (I–V) curves are shown in Fig. 7A and 7B, respectively. The currents were acquired in response to test pulses in the absence (Fig. 7Aa) or the presence (Fig. 7Ab) of a 1 s inactivating prepulse to –20 mV. As a result of this conditioning pre-pulse, the inactivating component (T-current) of the total Ca²⁺ current was eliminated, allowing us to dissociate the L-channel activity during the following test pulses (Fig. 7Ab). The T-channel activity is subsequently isolated, by means of algebraically subtracting (off-line) the L-current traces from the total Ca²⁺ current (Fig. 7Ac).

View larger version (34K): [in this window] [in a new window]   Figure 7.  The effects of TGF-ß1 and BMP-2 on the Ca2+ channels activity are entirely explained by significant alterations in the maximal channels conductance (Gmax) A, experimental procedure used to dissect the T- and L-channel activity. The current traces were obtained from a control myotube (6 days). B, I–V curves from the current traces recorded as in A. Families of current traces were elicited from –80 mV to record the total Ca2+ current (Aa and Ba). Subsequently, 1 s inactivating prepulses to –20 mV were used to inactivate T-channels, and a second set of test pulses applied, to activate only the ‘inactivation-resistant’ or L-currents (Ab and Bb). The contribution of T-channels to the total Ca2+ current (Ac and Bc) was subsequently isolated by off-line subtracting L-currents (Ab), from the corresponding total Ca2+ currents (Aa). Table 1 shows kinetic parameters resulting from fitting the T- (Bc) and L-channel (Bb) I–V curves according to a Boltzmann equation (eqn (1)). Both BMP-2 and TGF-ß1 significantly reduced (80%) the T-channel maximal conductance (Gmax), whereas only TGF-ß1 significantly reduced the L-channel Gmax (90%; Table 1).

To evaluate possible alterations in the voltage-dependent activation of Ca²⁺ channels, we fitted the L- and T-channel I–V curves according to a Boltzmann equation (eqn (1)). The resulting kinetic parameters are shown in Table 1. Both BMP-2 and TGF-ß1 significantly reduced (80%) the control values for the T-channel maximal conductance (G_max), whereas the G_max values for the L-channels were significantly reduced (90%) only by TGF-ß1. Conversely, no robust changes were found in the voltage required for 50% activation of G_max (V), the slope factor (k), and the extrapolated reversal potential (V_rev; Table 1). These selective reductions in G_max fully explain the corresponding effects on the Ca²⁺ channel activity.

The results shown in Fig. 7 and Table 1 strongly suggest that the long-term effects of BMP-2 and TGF-ß1 on myotube formation involve a down-regulation of either the Ca²⁺-channel unitary conductance or the number of functional channels. Alternatively, the results might be explained by the presence of significantly higher percentages of inactivated channels at the holding potential (HP; –80 mV). We decided therefore to explore this possibility, by investigating the voltage-dependent inactivation of T- and L-channels.

Even though TGF-ß1- and BMP-2-treated myotubes expressed only barely resolved T-currents, we generated complete voltage-dependent inactivation curves for T-channels from three TGF-ß1- and three BMP-2-treated myotubes (Fig. 8). Except for a marked reduction in the maximal current amplitude, I_max (4-fold; for TGF-ß1, and 6-fold for BMP-2), the other Boltzmann parameters describing the inactivation curves (i.e. V and k) were very similar to the corresponding average values obtained from 12 control myotubes (see the legend of Fig. 8).

View larger version (17K): [in this window] [in a new window]   Figure 8.  The voltage-dependent inactivation of T-channels is practically unaltered by TGF-ß1 and BMP-2 A, T-currents recorded from a 6-day TGF-ß1-treated myotube. The currents were elicited from five different holding potentials (left), in response to test pulses to –20 mV. The holding potential was changed in increments of 10 mV, and set 30–60 s before test pulse applications. B, average peak current amplitudes plotted as a function of the holding potential. Experimental data were obtained from 12 control (•), three TGF-ß1- (grey circles), and three BMP-2-treated myotubes (). Fitting the experimental data to eqn (3) yielded the following Boltzmann parameters: for control, BMP-2-, and TGF-ß1-treated myotubes, respectively: Imax (pA): 340 ± 91, –76 ± 11 and –97 ± 24; k (mV): –3.3 ± 0.6, –4.4 ± 1.2 and –3.7 ± 0.7; V (mV): –49 ± 2.0, –55 ± 2.1 and –52 ± 2.2.

View larger version (21K): [in this window] [in a new window]   Figure 9.  The regulatory effects of TGF-ß1 and BMP-2 do not involve major alterations in the voltage-dependent inactivation of L-channels A, representative L-currents from a BMP-2-treated myotube. The current traces were elicited from five different holding potentials (left). The holding potential was applied 30–60 s before test pulse applications. B, average L-channel inactivation curves obtained from control (n= 8), BMP-2- (n= 7), and TGF-ß1-treated (n= 1) myotubes. The current amplitude at the end of 200 ms test pulses recorded as in A is plotted as a function of the holding potential. The L-channel inactivation curves were fitted according to a Boltzmann equation (eqn (3)); and the resulting kinetic parameters are as follows, for control, BMP-2-, and TGF-ß1-treated myotubes, respectively: Imax (pA) –1315 ± 209, –503 ± 188 and –104; k (mV) –6.7 ± 0.8, –7.4 ± 1.2 and –3.2; V (mV) –13.5 ± 6.5, –7.0 ± 3.7 and –10.3. No significant differences were found, for the Boltzmann parameters V and k, between control and BMP-2-treated myotubes.

	Discussion

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Comparison with previous studies

Caffrey et al. (1989) investigated the chronic effects of TGF-ß1 on current densities carried through Ca²⁺ and Na⁺ channels, in a myogenic cell line (C2). In that study, C2 myoblasts were exposed for up to 14 days to fusion medium supplemented with 1 nM TGF-ß1. The treatment with TGF-ß1 literally abolished the Na⁺ and Ca²⁺ currents, in both amplitude and percentage of cells exhibiting inward currents. It has been reported that 1 nM TGF-ß1 reduces the fusion index virtually to zero. In fact, the EC₅₀ for the inhibition of myotube formation is close to 20 pM TGF-ß1, and 4 nM BMP-2 (Katagiri et al. 1994). We have therefore used 40 pM TGF-ß1 and 5 nM BMP-2, two concentrations at which the growth factors exert only a 60% reduction in myotube formation, but do not totally prevent the fusion, as determined by membrane capacitance measurements (this study) and counting the number of newly formed myotubes (Katagiri et al. 1994). By using these concentrations, we were able to characterize the macroscopic kinetic properties of T- and L-channels, in both TGF-ß1- and BMP-2-treated myotubes. Moreover, even though 6-day-treated myotubes are 60% smaller, compared to the corresponding 6-day-control myotubes, the treated myotubes still fuse poorly. This is based on the observation that the C_m values for the 6-day-treated myotubes are 3-fold higher (P < 0.05) than those for the non-fusing myoblasts at day 0 (i.e. myoblasts unexposed to the fusion medium). The absolute C_m values were as follows: 13.9 ± 3.2 pF (control myoblasts, day 0), 38.4 ± 12.1 pF (TGF-ß1, day 6), and 42.0 ± 5.4 pF (BMP-2, day 6; see also Fig. 1). Thus, the regulation of Ca²⁺ channels with 40 pM TGF-ß1 and 5 nM BMP-2 is most likely not due to a global alteration in the synthesis of proteins, since the treated myoblasts still fuse (TGF-ß1 and BMP-2) and normally express L-channels (BMP-2).

Several other intercellular chemical messengers, in addition to TGF-ß1 and BMP-2, also regulate the fusion of myoblasts. For example, insulin like growth factor 1 (IGF-1), which exert a stimulatory effect (Florini et al. 1996). Interestingly, Wang et al. (1999) investigated the effects of IGF-1 on Ca²⁺ channels from 5-day-old myotubes, following a 3-days exposure to IGF-1 (i.e. from days 2–5). As a result, the growth factor significantly increased the expression level of L-channels, but was unable to regulate the T-channel activity. Apparently, IGF-1 failed also to stimulate the formation of myotubes, as deduced by the similar values of C_m reported for the control and IGF-1-treated myotubes (151 ± 21 pF and 140 ± 18 pF, respectively). Unfortunately, speculation about a possible relationship between this absence of effects on T-channels and the fusion of myoblasts is complicated, since IGF-1 was only present 2 days after the onset of myotube formation. Thus, it will be important for future studies to determine if an appropriated treatment with IGF-1, which effectively increases myogenesis, is also able to up-regulate the functional expression of T-channels. Likewise, it will be important to investigate if other voltage-dependent ion channels are being regulated by 40 pM TGF-ß1 and 5 nM BMP-2. For instance, the growth factors might also exert a fast modulation of K⁺ channels, because these channels are particularly important for the earliest events of myogenesis, by means of determining myoblast membrane potential (reviewed in Bernheim & Bader, 2002).

Possible molecular mechanisms involved

Our experiments show that TGFß-1 and BMP-2 significantly reduce the current amplitudes associated with both T- and L-channels. These inhibitory effects are parallel to the inhibition of myotube formation. Thus, the smaller size of the myotubes might contribute to explaining the observed reduction in current amplitudes. In fact, the inhibition by BMP-2 of the L-current is entirely explained by the smaller size of the myotubes, since BMP-2 was unable to significantly alter the L-current density. By contrast, the effects of BMP-2 and TGF-ß1 on the T-current, as well as the effect of TGF-ß1 on the L-current, are only explained to a minor degree by the reduced values of C_m (Figs 3D and 4D). This indicate that even though the onset of the inhibitory effects on Ca²⁺ channels parallels the inhibition on myotube formation, the regulatory effects on the T- (TGF-ß1 and BMP-2) and L-channels (TGF-ß1) cannot be solely explained by the smaller size of the myotubes.

On the other hand, all patch-clamp experiments were carried out in the absence of growth factors (i.e. external solution), and thus the observed effects persist up to 2 h (see Methods). This suggests that slow molecular mechanisms are involved, as opposed to a fast modulation of the function of a fixed population of channels. In keeping with this view, the functional properties of the channels were practically unaltered (Figs 6–9). Only major alterations were determined in the maximal Ca²⁺ channels conductance (G_max) (Table 1), which depends on the number of functional channels. Thus conceivably, alterations in the number of Ca²⁺ channel forming proteins (like the principal _1S and _1H subunits), or even a possible regulatory protein(s), which could be altering the surface expression and/or unitary conductance of the channels, might account for the effects. In this regard, it is worth mentioning that the mRNA levels of myogenic transcription factors, like myogenin and MyoD, are regulated following 1–24 h treatments of myoblasts with BMP-2 and TGF-ß1 (Katagiri et al. 1994). It is possible therefore that this long-term regulation of Ca²⁺ channels might represent a down-stream effect, resulting from alterations on the expression level of these transcription factors.

TGF-ß1 and BMP-2 regulate a diverse array of cellular processes, and are predominantly produced by platelets and osteoblasts, respectively. The signalling pathways that mediate their effects have been previously reviewed extensively (e.g. Matzuk, 1995; Kawabata et al. 1998; Yamaguchi et al. 2000; ten Dijke et al. 2003; Serra & Chang, 2003). In brief, the growth factors bind to two types of functional receptors (I and II), which are single-pass transmembrane serine/threonine kinases. At least eight mammalian signal-transducing molecules of the TGF-ß superfamily, termed Smads, have been identified. Smads belonging to a pathway-restricted subgroup (RSmad) are activated by binding of specific ligands to type I receptors. Once phosphorylated, Smads are translocated into the nucleus, where they regulate the transcription of targeted genes by directly binding to DNA. The fact that different Smads are selectively involved in BMP (Smad 1, Smad 5 and Smad 8) and TGF-ß (Smad 2 and Smad 3) signalling might contribute to explaining why the functional expression of L-channels is only inhibited by TGF-ß1.

Skeletal muscle EC coupling involves a physical interaction between the _1S subunit of L-channels (DHPRs) with ryanodine receptors (RyR1s), located at the terminal cisternae of the sarcoplasmic reticulum (SR). The DHPRs undergo conformational changes in response to electrical depolarizations of the sarcolemma, resulting in the activation of nearby RyR1s, and a massive release of Ca²⁺ from the SR (orthograde coupling; reviewed in Melzer et al. 1995; Dirksen, 2002). In addition, the RyR1s exert a reciprocal regulation of the L-channel activity (retrograde coupling). Specifically, RyR1s increase the L-channel G_max, and reduce activation rate (Nakai et al. 1996; Avila & Dirksen, 2000). Thus, conceivably, alterations in DHPR/RyR1 physical interaction might contribute to explaining the observed effects on L-channel G_max (Table 1). Nevertheless, the determination of entirely normal L-channel activation kinetics, in both TGF-ß1- and BMP-2-treated myotubes (Fig. 6), points to the absence of a DHPR/RyR1 functional expression mismatch, and rules out a possible contribution of the retrograde coupling to the observed effects.

Physiological relevance

The physiological relevance for the specific regulation of L-channels by TGF-ß1 but not BMP-2 is not clear yet. As outlined in the previous section, the growth factors did not alter the L-channel activation kinetics, which is consisting with the DHPRs still interacting with RyR1s, and probably transmitting the electrical stimulus in EC coupling. However, since TGF-ß1 but not BMP-2 drastically reduces the L-channel activity, TGF-ß1-treated myotubes are most unlikely to exhibit EC coupling, whereas the BMP-2-treated myotubes might be still conserving a normal skeletal muscle-type EC coupling. On the other hand, it has been proposed that BMP-2 functions to establish a sufficient number of dividing myoblasts, which eventually will fuse and enlarge the muscle tissue (Parker et al. 2003). In this regard, the possibility of normal EC coupling in the BMP-2-treated myotubes fits well with this interpretation. Since the T- and L-channel activity is practically absent in the TGF-ß1-treated myotubes, it is tempting to speculate that TGF-ß1 might be acting to guarantee regeneration of the original satellite or progenitor cells, in the absence of an irreversible commitment of these cells to the skeletal muscle phenotype.

As mentioned before, a pivotal role has been categorically established for the principal subunit of the L-channels (_1S) in transmitting the electrical stimulus in skeletal muscle EC coupling (Tanabe et al. 1988; Melzer et al. 1995; Dirksen, 2002; see also the previous section). By contrast, the physiological role of T-channels in skeletal muscle is just beginning to be unravelled. Beam & Knudson (1988b) hypothesized that the T-channels might be important in regulating the fusion of myoblasts. Experimental evidence in support of this view was generated more recently. Specifically, the fusion of myoblasts is drastically reduced in response to a long-term exposure (1–3 days) of human myoblasts to Ni²⁺ and amiloride, two T-channel antagonists. This suggests a critical role for Ca²⁺ influx through T-channels in stimulating myotube formation (Bijlenga et al. 2000). Within this context, our results strongly suggest that the inhibitory effects of BMP-2 and TGF-ß1 on myogenesis might depend, at least partially, on the observed reduction in the functional expression of T-channels.

How might a significant reduction in the T-channel functional expression, in this case induced by TGF-ß1 and BMP-2, contribute to preventing the fusion of myoblasts? It has been reported that intracellular Ca²⁺ ([Ca²⁺]_i) increases prior to myotube formation and induces the fusion of myoblasts (David et al. 1981). It is also known that a small proportion of competent (i.e. ready to fuse) myoblasts (20%) exhibit a significantly higher [Ca²⁺]_i, compared to the other 80% (105 ± 21 nMversus 65 ± 14 nM, respectively) (Bijlenga et al. 2000). Interestingly, this subpopulation of myotubes with a higher [Ca²⁺]_i is not present following 3–4 day treatments with T-channel blockers (Ni²⁺ and amiloride). Moreover, the same treatment drastically reduces the fusion index, strongly suggesting that the increased [Ca²⁺]_i and fusion index both depend on Ca²⁺ influx through T-channels (Bijlenga et al. 2000). The higher [Ca²⁺]_i might be activating a Ca²⁺-dependent signalling pathway, involving perhaps calmodulin as a Ca²⁺ sensor (Bar-Sagi & Prives, 1983), and the nuclear factor of activated T cell (NFAT) family of transcription factors (Pavlath & Horsley, 2003). Eventually, activation of this signalling pathway might lead to the fusion of myoblasts (reviewed in Horsley & Pavlath, 2004).

On the other hand, if Ca²⁺ influx through T-channels effectively represents an important modulator of myoblast fusion, then this influx of Ca²⁺ must be somehow specifically regulated. To achieve this, an efficient way would be to prevent the expression of T-channels. Thus, by keeping low the T-channel functional expression, and the consequent reduction in the influx of Ca²⁺, through a previously determined window current (Bijlenga et al. 2000; Berthier et al. 2002), the growth factors might be acting to reduce the fusion index.

Within this context, the 15–25% of myoblasts exhibiting T-currents during the earliest steps of myogenesis, and the subsequent T-currents' abolition in response to TGF-ß1 and BMP-2 (Fig. 3B), are equivalent to the 20% of competent myoblasts exhibiting higher [Ca²⁺]_i, and the corresponding [Ca²⁺]_i decrease by Ni²⁺ and amiloride (Bijlenga et al. 2000). This suggests the growth factors might decrease also the proportion of myoblasts exhibiting a higher [Ca²⁺]_i. If that is the case, TGF-ß1 and BMP-2 might prevent activation of the proposed Ca²⁺–calmodulin signalling pathway (Horsley & Pavlath, 2004). More work is obviously needed to clarify the specific molecular mechanisms involved. Nevertheless, the reported effects of TGF-ß1 and BMP-2 on Ca²⁺ channels contribute to explaining the associated effects on myogenesis.

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	Acknowledgements

 
We are in debt to Dr Gabriel Cota for allowing us to use his patch-clamp system. We also thank Dr Ricardo Felix for helpful suggestions to improving the manuscript, and M. in Sc. Jose Luis Martinez and Felipe Cruz for technical assistance. This study was supported by the CONACyT grant 39512 to G.A.

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