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Functional Interaction of Ca_V Channel Isoforms with Ryanodine Receptors Studied in Dysgenic Myotubes

Ralph Peter Schuhmeier ^*, Elodie Gouadon ^*, Daniel Ursu ^*, Nicole Kasielke , Bernhard E. Flucher , Manfred Grabner  and Werner Melzer ^*

^* Department of Applied Physiology, University of Ulm, Ulm, Germany; Department of Physiology and Medical Physics, and Department of Medical Genetics, Molecular and Clinical Pharmacology, Innsbruck Medical University, Innsbruck, Austria

Correspondence: Address reprint requests to Werner Melzer, University of Ulm, Dept. of Applied Physiology, Albert-Einstein-Allee 11, D-89069 Ulm, Germany. Tel.: 49-731-500-23248; Fax: 49-731-500-23260; E-mail: werner.melzer{at}medizin.uni-ulm.de.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
The L-type Ca²⁺ channels Ca_V1.1 (_1S) and Ca_V1.2 (_1C) share properties of targeting but differ by their mode of coupling to ryanodine receptors in muscle cells. The brain isoform Ca_V2.1 (_1A) lacks ryanodine receptor targeting. We studied these three isoforms in myotubes of the _1S-deficient skeletal muscle cell line GLT under voltage-clamp conditions and estimated the flux of Ca²⁺ (Ca²⁺ input flux) resulting from Ca²⁺ entry and release. Surprisingly, amplitude and kinetics of the input flux were similar for _1C and _1A despite a previously reported strong difference in responsiveness to extracellular stimulation. The kinetic flux characteristics of _1C and _1A resembled those in _1S-expressing cells but the contribution of Ca²⁺ entry was much larger. _1C but not _1A-expressing cells revealed a distinct transient flux component sensitive to sarcoplasmic reticulum depletion by 30 µM cyclopiazonic acid and 10 mM caffeine. This component likely results from synchronized Ca²⁺-induced Ca²⁺ release that is absent in _1A-expressing myotubes. In cells expressing an _1A-derivative (₁Aas(1592-clip)) containing the putative targeting sequence of _1S, a similar transient component was noticeable. Yet, it was considerably smaller than in _1C, indicating that the local Ca²⁺ entry produced by the chimera is less effective in triggering Ca²⁺ release despite similar global Ca²⁺ inward current density.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
The mobilization of Ca²⁺ for force activation in muscle cells involves the rapid communication of voltage-dependent Ca²⁺ channels with the ryanodine receptors of the sarcoplasmic reticulum (SR) (Rüegg, 1986; Melzer et al., 1995; Bers, 2001; Dirksen, 2002; Beam and Horrowitz, 2004). In cardiac cells, a Ca²⁺ inward flux mediated by voltage-operated L-type Ca²⁺ channels triggers a larger secondary Ca²⁺ efflux from the SR (DelPrincipe et al., 1999; Cheng and Wang, 2002). This mechanism requires the close proximity of the pore-forming subunit _1C of the cardiac L-type channel (Ca_V1.2) with the Ca²⁺ release channel of the SR (ryanodine receptor RyR2) (Franzini-Armstrong et al., 1999; Bers, 2001). In contrast, the skeletal muscle L-type Ca²⁺ channel (_1S = Ca_V1.1) interacts with the ryanodine receptor (RyR1) via a direct physical link (Dirksen, 2002; Beam and Horrowitz, 2004). The protein-protein interaction is reflected in a regular tetrameric pattern (junctional tetrads) formed by the Ca_v1.1 in the transverse tubular (TT) membrane opposite the RyR1 clusters (Franzini-Armstrong et al., 1998, 2004). The correct targeting to the TT-SR junction that is essential for both types of interaction mechanisms seems to depend on amino acid sequences in the C-terminal regions of _1C and _1S (Flucher et al., 2000; Proenza et al., 2000). The skeletal muscle-specific type of direct interaction with the ryanodine receptor, on the other hand, has been attributed to specific sequences in the II–III loop of the _1S isoform (Nakai et al., 1998; Grabner et al., 1999; Dirksen, 2002) and may involve further cytoplasmic regions of this protein (Ahern et al., 2001) as well as the participation of the auxiliary ß_1a subunit (Beurg et al., 1999a,b).

The _1A-subunit (Ca_V2.1) of neuronal P/Q-type Ca²⁺ channels is thought to lack both a specific junctional targeting sequence as well as a RyR interaction domain (Flucher et al., 2000). Immunolabeling after heterologous expression in _1S-deficient dysgenic myotubes showed colocalization with RyR1 for _1S and _1C but not for _1A. However, a chimeric construct consisting of a truncated form of _1A fused with the distal half of the C-terminal _1S sequence, ₁Aas(1592-clip), restored targeting to the ryanodine receptor based on the immunostaining results (Flucher et al., 2000).

The scheme of Fig. 1 summarizes putative functional properties of the four ₁ variants in dysgenic myotubes: All constructs except _1S should lead to intracellular Ca²⁺ signals that depend on the size of the voltage-activated Ca²⁺ inward current rather than on membrane voltage alone. Because of the lack of specific junctional targeting, _1A should be least effective in triggering secondary Ca²⁺ release and may even not be able to cause Ca²⁺ release at all. Finally, ₁Aas(1592-clip), like _1C, would be expected to restore cardiac-type EC coupling exhibiting a significant secondary Ca²⁺ release component.

View larger version (14K): [in this window] [in a new window]   FIGURE 1  Expression of different Ca2+ channel 1-subunits in dysgenic (GLT) myotubes. The four Ca2+ channel 1-subunits that were expressed in the 1-deficient GLT myotubes are known to differ in their location (specific targeting to the TT-SR junction or not) and their mode of interaction with the ryanodine receptor (direct mechanical coupling or not). Based on previous structural and functional investigations, 1S controls release from the SR without the participation of a Ca2+ inward flux. 1C and 1Aas(1592-clip) which are colocalized with RyR1 should support Ca2+ inward current-induced Ca2+ release whereas 1A which was reported not to be targeted to the junction should not be able to functionally interact with the junctional ryanodine receptors.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
Solutions and media
The following solutions were used in the cell culture:

Growth medium—Dulbecco's Modified Eagle Medium (DMEM) with 1% glutamine (200 mM), 10% fetal calf serum (FCS), and 10% horse serum (HS).

Trypsin solution—Trypsin-EDTA (0.05–0.53 µM) in Ca²⁺- and Mg²⁺-free phosphate-buffered saline (PBS).

Fusion medium—DMEM with 1% glutamine (200 mM) and 2% HS.

Transfection medium—100 µl per 35-mm dish, containing 94 µl serum-free DMEM, 6 µl Fugene 6 (see below), and 2 µg DNA.

The experimental recording solutions had the following compositions:

External solution—130 mM tetraethylammonium hydroxide, 127 mM HCl, 10 mM CaCl₂, 1 mM MgCl₂, 2.5 mM 4-aminopyridine, 0.00125 mM tetrodotoxin, and 10 mM HEPES, pH-adjusted to 7.4 with HCl.

Internal (pipette) solution—145 mM CsOH, 145 mM aspartic acid, 0.1 mM EGTA, 0.01 mM CaCl₂ (free Ca²⁺, 10^–5 mM), 1.14 mM Na₂ATP, 3.86 mM MgATP (Mg/ATP ratio according to supplier, 1.4; free Mg²⁺, 1 mM), 5 mM Na creatine phosphate, 0.2 mM K₅-Fura-2, and 10 mM HEPES, pH-adjusted to 7.4 with CsOH.

DMEM, HS, and Trypsin-EDTA were purchased from Gibco (Karlsruhe, Germany), FCS, and PBS from PAA Laboratories (Cölbe, Germany), rat tail collagen (Typ 1, C 7661), EGTA, ATP, caffeine, cyclopiazonic acid, and CsOH from Sigma-Aldrich (Taufkirchen, Germany), Fugene 6 from Roche Biochemicals (Mannheim, Germany), TEA and HEPES from Merck (Darmstadt, Germany), tetrodotoxin and Fura-2 from Molecular Probes (Leiden, Netherlands), creatine phosphate from Boehringer (Mannheim, Germany), 4-aminopyridine from Fluka (Neu-Ulm, Germany), and L-glutamine from Biochrom (Berlin, Germany).

Cell culture and transfection
For heterologous expression of the Ca²⁺ channel ₁-subunits, we used myotubes of the homozygous dysgenic (mdg/mdg) cell line GLT that was originally generated by transfection of mdg myoblasts with a plasmid-encoding Large-T Antigen (Powell et al., 1996). GLT myoblasts were expanded in growth medium at 10% CO₂ and 37°C and passaged before reaching 80% confluence using trypsin detachment. To obtain myotubes, cells were plated onto carbon- and gelatin-coated coverslips in 35-mm dishes. After reaching confluence, growth medium was exchanged for fusion medium. We used mammalian expression plasmids coding for N-terminally GFP-tagged fusion proteins of the Ca²⁺ channel pore-forming subunits _1S, _1C, _1A, and ₁Aas(1592-clip), respectively, as described by Flucher et al. (2000). Cells were transfected at the onset of fusion. Measurements were made from the myotubes four days later.

Electrophysiology and fluorimetry
Whole-cell patch-clamp experiments and microfluorimetric recordings were performed at room temperature (20–23°C) as described by Schuhmeier et al. (2003) and Schuhmeier and Melzer (2004). Briefly, myotubes were loaded with Fura-2 by diffusion from a patch-pipette containing internal solution (see above). Leak-resistance and capacitance were determined by using small positive and negative pulse excursions (10 mV amplitude, 25 ms duration) from the holding potential. Linear leak correction of current recordings was performed by a standard p/n method (n = 4 or 8). Ca²⁺-dependent fluorescence changes were recorded at 515 nm while exciting at 380 nm (F₃₈₀). After background correction, the F₃₈₀ recordings were normalized either by F₃₈₀ measurements preceding each voltage-clamp activation (Figs. 2–4, displayed as –F/F₀), or by F₃₆₀ (Figs. 5–8), as described in Schuhmeier et al. (2003).

View larger version (20K): [in this window] [in a new window]   FIGURE 2  Simultaneously recorded Ca2+ inward currents and intracellular Ca2+ signals in GLT myotubes. (A) Recordings of leak-corrected Ca2+ inward currents (a and c) and fluorescence changes (b and d) in GLT myotubes transfected with expression plasmids carrying cDNA encoding 1S and 1C, respectively. (B) Peak Ca2+ current densities (a and c) and Ca2+ transients (b and d) as functions of voltage. Continuous lines are least-squares fits with Eqs. 1 and 2, i.e., formally the same functions were used to describe the voltage-dependence of transmembrane Ca2+ current density and intracellular Ca2+ transient.

View larger version (23K): [in this window] [in a new window]   FIGURE 3  Voltage-dependence of Ca2+ inward current density and intracellular Ca2+ transient. Mean data obtained from recordings in transfected GLT myotubes. (A) 1S (n = 6), (B) 1C (n = 5), (C) 1A (n = 5), and (D) 1Aas(1592-clip) (n = 10). The continuous curves show least-squares fits according to Eqs. 1 and 2 (see also Fig. 2).

View larger version (22K): [in this window] [in a new window]   FIGURE 4  Voltage-dependence of activation. The inward current data of Fig. 3 were converted to fractional conductance using Eq. 1 and solving for f(V) (solid symbols). A formally equivalent conversion was performed using the optically recorded Ca2+ signal data of Fig. 3 (open symbols). The two types of activation curves differ clearly for 1S (A) and to a lesser degree for 1C (B), whereas for 1A (C) and 1Aas(1592-clip) (D), the difference is below 5 mV.

View larger version (12K): [in this window] [in a new window]   FIGURE 5  Comparison of Ca2+ input flux and Ca2+ entry flux. (A) Averaged Ca2+ input flux calculations for the constructs shown in Fig. 1 during 100 ms depolarizations to +30 mV. (B) Averaged Ca2+ inward current densities for the same pulses that elicited the records in A. Number of experiments from left to right: n = 17, 16, 16, and 15. Thick lines indicate mean values; shaded areas indicate mean ± SE.

View larger version (20K): [in this window] [in a new window]   FIGURE 6  Effect of SR depletion on EC coupling gain. (A) 30 µM CPA and 10 mM caffeine were applied to deplete the SR of stored Ca2+ to study the effect on Ca2+ fluxes and gain in a number of myotubes expressing the four 1-subunits of Fig. 1. (B) Time-dependent changes in mean baseline free Ca2+ concentration indicating comparable Ca2+ release in all cases. (C) Mean Ca2+ input flux and Ca2+ entry flux (average from 25 to 75 ms during the voltage pulse) for each of the three intervals, labeled a, before drug application; b, after CPA application; and c, after caffeine application. Entry flux plotted in opposite direction of input flux. (D) A gain factor was calculated representing the ratio of Ca2+ input flux (derived from the optical signals) and Ca2+ entry flux (derived from the inward current density) using the data of C. Because small current amplitudes produced large variances, current density amplitudes <0.5 A/F were excluded for the gain calculation of 1S in D. From left to right, the numbers of cells were 7, 4, 5, and 8, respectively.

View larger version (24K): [in this window] [in a new window]   FIGURE 7  Time course of Ca2+ input and entry flux during SR depletion. Time course of calculated Ca2+ fluxes (Ca2+ input flux drawn upward, Ca2+ entry flux downward). Columns (a), (b), and (c) show the averaged responses to pulse depolarization (100 ms, +30 mV) in the correspondingly labeled intervals of Fig. 6 B, respectively. Column (d) shows the depletion-sensitive flux components. After scaling with a factor f to correct for a rundown in the current traces, the depolarization-induced responses persisting after CPA and caffeine application (column c) were subtracted from the responses before drug application (column a). The subtraction was performed for each individual experiment and the differences were averaged. Shaded areas indicate point by point SE.

View larger version (17K): [in this window] [in a new window]   FIGURE 8  Correlation between Ca2+ entry flux and Ca2+ input flux. Mean values of Ca2+ input flux plotted versus corresponding means of Ca2+ entry flux, evaluated at the input flux peak (A) and near the end of the pulse (95 ms) (B). Each connected data points correspond to Fig. 7, a, b, and c.

The leak-corrected ionic currents I(V) were normalized by the linear capacitance to obtain current densities i(V). The voltage-dependence of the Ca²⁺ current density i_Ca(V) was least-squares-fitted with Eq. 1:

(1)

(2)

The values g_Ca,max and V_Ca are maximal normalized conductance and reversal potential, respectively, of the L-type Ca²⁺ current; f(V), voltage-dependence of activation; and V_0.5 and k, voltage of half-maximal activation and voltage sensitivity parameter, respectively. The fit also included correction for a small offset resulting from noise effects in the minimum detection.

Ca²⁺ current densities i_Ca were converted to Ca²⁺ entry flux using Eq. 3:

(3)

Here, z is the valence of the Ca²⁺ ion, F is the Faraday's constant, V_C is the intracellular volume per membrane capacitance, and f_V is the fraction of the total volume that is immediately accessible to Ca²⁺.

Free Ca²⁺ concentration was determined using background- and bleaching-corrected fluorescence ratio signals (R = F₃₈₀/F₃₆₀) according to Eq. 4 (Klein et al., 1988):

(4)

The following values were used for R_min (fluorescence ratio at zero dye saturation), R_max (ratio at full dye saturation), and K_D,Fura (dissociation constant of the dye): 2.84, 0.68, and 276 nM, respectively (see Schuhmeier et al., 2003). For the Fura-2 dissociation rate constant k_off,Fura we used the value of 46.4 s^–1 (Schuhmeier and Melzer, 2004).

An estimate of the flux of Ca²⁺ mobilization in the myoplasm (called Ca²⁺ input flux) during strong depolarizations was calculated with a Ca²⁺ binding model using the general method of Baylor et al. (1983). The free Ca²⁺ transient (Eq. 4), averaged for three sequential voltage pulses (+30 mV) of 100-ms duration, applied at 30-s intervals, was used to calculate the Ca²⁺ occupancies of the model components. Free Ca²⁺ and the estimated occupancies were summed and the time derivative calculated. Differential equations were solved using Euler's method. The calculation employed a digital filter that adjusted its bandwidth automatically to the signal dynamics (Schuhmeier et al., 2003). In the model we used troponin C with concentration (120 µM) and kinetic properties as reported for skeletal muscle fibers (Baylor and Hollingworth, 1998). Each molecule of troponin C has two fast, Ca²⁺-specific binding sites (T-sites) and two slow Ca²⁺-Mg²⁺-binding sites (P-sites) with rate constants k_on,T,Ca = 88.5 µM^–1 s^–1, k_off,T,Ca = 115 s^–1, k_on,P,Ca = 41.7 µM^–1 s^–1, k_off,P,Ca = 0.5 s^–1, k_on,P,Mg = 0.033 µM^–1 s^–1, and k_off,P,Mg = 3 s^–1. In contrast to mature muscle fibers, there is no evidence for the presence of parvalbumin in developing muscle (Leberer and Pette, 1986). EGTA (0.1 mM) in the myoplasm was described by using the means of in situ rate constants (k_on,S = 20 µM^–1 s^–1 and k_off,S = 2.71 s^–1) determined empirically in C2C12 myotubes loaded with large excess of EGTA (Schuhmeier and Melzer, 2004). Because SR uptake rates in myotubes seem to be small compared to release rates during depolarization (García and Beam, 1994), a component simulating the SERCA pump was not included in the model.

Statistics
Data are presented as means ± SE (n = number of experiments) for averaged values, and as parameter ± SE for best-fit parameters.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
To determine the voltage-dependence of both Ca²⁺ inward currents and Ca²⁺ transients, the pulse paradigm described in Fig. 2 was used. The recordings in Fig. 2 A were obtained from one GLT cell expressing _1S (a and b) and another one expressing _1C (c and d). The length of the voltage-clamp pulse was 200 ms in all cases (horizontal bar). Leak-corrected Ca²⁺ inward current (I_Ca) and normalized change in fluorescence (–F/F₀) are displayed next to each other for a series of pulses from –20 mV to +70 mV.

Fig. 2 B depicts the voltage-dependence obtained from each series of recordings, upon taking the signal peak for the inward current (solid symbols) and the average of the last 10 ms during the pulse for the fluorescence (open symbols). Clearly, in the case of _1S the threshold for activation of Ca²⁺ transients (b) is considerably more negative than for the activation of inward current (a) and Ca²⁺ transients are of almost equal amplitude between +50 and +80 mV. In contrast, in the case of _1C, Ca²⁺ current (c) and Ca²⁺ transient (d) start at a similar potential and show a similar decline in amplitude at large depolarizing potentials.

Fig. 3 compares voltage-dependence of Ca²⁺ current (solid symbols) and Ca²⁺ transient (open symbols) for each of the ₁-subunit types of Fig. 1 using averaged data of individual experiments. _1S produced the smallest Ca²⁺ current densities but the largest Ca²⁺ transients (Fig. 3 A). The _1S-transients show only a small decrease in amplitude at large voltages whereas the decrease is substantial in the three other cases (Fig. 3, B–D). The bell-shaped fluorescence-voltage relations for _1C, _1A, and ₁Aas(1592-clip) indicate a close relation between inward current and Ca²⁺ signal. The maximal mean Ca²⁺ signal of _1S was significantly larger than that of _1C and _1A (but not ₁Aas(1592-clip)). The putative junctionally targeted ₁Aas(1592-clip) showed a significantly larger maximal current density than the nontargeted _1A channel. On the other hand, the amplitude of the ₁Aas(1592-clip) Ca²⁺ signal was not found to be significantly larger than that of _1A.

To better compare the voltage-dependence of activation of Ca²⁺ current and intracellular Ca²⁺ transients we formally fitted both leak-corrected current and fluorescence data in Fig. 3 with Eq. 1. The best-fit functions are superimposed on the data points as continuous lines using the means of the individual best-fit parameters.

Fig. 4 displays the data of Fig. 3 after conversion to f(V) to show the voltage-dependence of fractional activation according to Eq. 2. If the Ca²⁺ transient is the immediate result of the Ca²⁺ entry flux, the two activation curves are expected to be similar. As can be seen from these plots, for the skeletal muscle ₁-subunit (_1S), the optically recorded Ca²⁺ transient reaches its half-maximal value at a considerably more negative potential than the Ca²⁺ conductance activation (Fig. 4 A, open and solid circles, respectively). The midpoint voltages of activation are separated by 29.9 mV. In the case of the cardiac ₁-subunit (_1C) the activation curves are closer together, but still separated by a gap of 8.6 mV (Fig. 4 B). In contrast, the two activation curves show much smaller differences for _1A and its chimeric construct, 3.3 and 4.1 mV, respectively (Fig. 4, C and D).

Table 1 summarizes the parameters describing the voltage-dependence obtained from the data of Figs. 3 and 4. The two L-type channels can be easily distinguished from _1A and its chimeric derivative by the lower steepness of activation (larger k). Within each group, steepness of activation of Ca²⁺ entry and intracellular Ca²⁺ signals were similar.

In some experiments with _1S-expressing myotubes, we also applied the approach described recently to determine Ca²⁺ input flux in C2C12 myotubes equilibrated with a high concentration (15 mM) of EGTA in the pipette solution (Schuhmeier and Melzer, 2004). In this method, removal model parameters were determined by a least-squares fitting of model-calculated to measured Ca²⁺ transients in the time intervals after depolarizing pulses. This method was not generally used in the present investigation, because of low signal/noise ratios that made the numerical calculations difficult and because of possible suppression of Ca²⁺-induced Ca²⁺ release by the strong EGTA buffering. Two _1S-expressing cells that were analyzed in this way showed removal parameter results and a time course of Ca²⁺ input flux comparable to those determined in C2C12 cells (Schuhmeier and Melzer, 2004). They also showed a phasic-tonic time course similar to that estimated for the _1S-expressing cells in the present experiments with 0.1 mM EGTA in the pipette solution.

In addition to the Ca²⁺ input flux derived from the optical indicator signal, the flux of Ca²⁺ entry was determined from the electrically measured Ca²⁺ inward current. The transmembrane Ca²⁺ inward current density was converted to flux according to Eq. 3. The necessary volume/capacitance ratios (V_C) were measured by scanning dye-loaded and whole-cell patch-clamped GLT myotubes with a confocal microscope as described by Schuhmeier et al. (2003). Volume (corrected for the space occupied by nuclei) was approximately proportional to capacitance in the range 200–800 pF, with a best-fit proportionality factor of V_C = 0.26 l/F that was used for the calculation. The factor f_V was arbitrarily set to 1 (the upper bound of this value).

Fig. 5 A shows the averaged Ca²⁺ input flux records obtained at +30 mV in a number of cells expressing the four different ₁-subunits shown in Fig. 1. Mean values are displayed as thick lines and their standard errors as shaded areas. The corresponding Ca²⁺ entry flux traces derived from the Ca²⁺ inward currents are plotted in Fig. 5 B with the same scale (for comparison of flux amplitudes) but opposite in sign.

Even though the calculated absolute flux amplitudes are somewhat questionable because of uncertainties in the model parameters, a relative comparison shows some interesting details. Consistent with Fig. 3, the Ca²⁺ input flux was found to be largest in the _1S-expressing myotubes whereas the Ca²⁺ entry flux was very small, indicating that essentially all the Ca²⁺ input flux resulted from SR Ca²⁺ release. ₁Aas(1592-clip), the brain ₁-subunit carrying the putative signal sequence for SR-TT junctional targeting, showed the second-largest flux amplitude. Here, however, the Ca²⁺ entry flux from the extracellular space was many times larger than in the case of _1S. Therefore, a much larger part of the total estimated Ca²⁺ input flux resulted from Ca²⁺ entry and it appears difficult to determine which component of the Ca²⁺ input flux results from SR Ca²⁺ release and which from Ca²⁺ entry. A similar situation exists for _1A and _1C.

Based on previous physiological data, one should expect clear differences between the junctionally targeted and nontargeted channels in their effectiveness to elicit intracellular Ca²⁺ transients. When activated by extracellular electrical stimulation, _1A-expressing myotubes had been reported to show much weaker force and Ca²⁺ responses than _1C-expressing myotubes which was attributed to the lack of specific junctional targeting (Flucher et al., 2000). In contrast to this, under voltage-clamp conditions in the present experiments, both channel types produced approximately equal input flux amplitudes.

Estimating EC coupling gain
The ratio of Ca²⁺ transient amplitude to Ca²⁺ inward current has frequently been used as a measure of EC coupling gain (for references, see Bers, 2001). On the other hand, the ratio between total Ca²⁺ input flux, which includes Ca²⁺ release, and the flux of Ca²⁺ entry, defines a physically more meaningful gain (Wier et al., 1994). Using the flux determinations of Fig. 5, we determined average EC coupling gain factors by calculating the mean ratio between total Ca²⁺ input flux and Ca²⁺ entry flux in a broad time interval during the pulse (from 25 ms to 75 ms) excluding the rapid phases of activation and deactivation. If the absolute amplitudes of the fluxes were correctly determined, a value of 1 of the gain factor would mean that no secondary release of Ca²⁺ from intracellular stores is present, i.e., all measured Ca²⁺ changes result from Ca²⁺ entering from the extracellular space. For the _1S-subunit, this gain factor was many times >1 (mean value 31.5 ± 4.4), consistent with the major contribution of the SR Ca²⁺ release to the Ca²⁺ transient and the negligible role of the Ca²⁺ current. For the three subunits that show no skeletal-type conformational coupling, the ratios were 3.2 ± 0.5 (_1C), 3.9 ± 0.4 (_1A), and 2.0 ± 0.3 (₁Aas(1592-clip)). That is, quite consistent with the experiments of Fig. 3, there were no large differences between the three non-skeletal-type channels in their mean EC coupling gain.

Effects of SR depletion
These results indicate that Ca²⁺ entry makes a substantial contribution to the total Ca²⁺ input flux, with the exception of _1S. However, considering the uncertainties in determining the absolute flux amplitudes (see above), it seems difficult to quantify the true fractional contribution. In particular, the value larger than unity for the gain factor of the non-junctionally targeted _1A-subunit may result from a Ca²⁺ release component in addition to Ca²⁺ entry or from false assumptions in the calculation of the absolute flux amplitudes leading to overestimation of Ca²⁺ input flux, underestimation of Ca²⁺ entry flux, or both. We therefore performed experiments in which the SR was depleted of its stored Ca²⁺ by applying 30 µM cyclopiazonic acid (CPA), a blocker of the SERCA Ca²⁺ pump (Schuhmeier and Melzer, 2004), and 10 mM of the ryanodine receptor agonist caffeine (Herrmann-Frank et al., 1999). This procedure has been shown to drastically reduce the flux component resulting from SR Ca²⁺ release in C2C12 myotubes and should reveal the component that consists only of the Ca²⁺ entry through the voltage-activated Ca²⁺ channels (Schuhmeier and Melzer, 2004).

Fig. 6, A and B, demonstrates the depletion protocol. The measurements shown covered a time interval of 10 min. Application of CPA and caffeine (as indicated by the bars in Fig. 6 A) caused changes in baseline free [Ca²⁺] values calculated from the fluorescence ratio immediately before each voltage pulse that was applied (Fig. 6 B). The mean values for each group of cells are shown. In all cases a similar increase in baseline Ca²⁺ concentration occurred when CPA was applied, indicating that a discharge of similar amounts of Ca²⁺ from the SR took place and that the loading state of the SR had not been considerably different before the application of CPA.

Fig. 6, C and D, summarize the mean results for fluxes and gains for all four ₁ isoforms. Fig. 6 C shows the means of Ca²⁺ input flux and Ca²⁺ entry flux (plotted upward and downward, respectively), obtained in the three regions labeled a, b, and c (i.e., before and after CPA application and after caffeine application). Fig. 6 D displays the means of the individually calculated gains from the data in C. The initial gains were very similar to those determined in the previous series of experiments (Fig. 5). In the _1S case (left column), the input flux amplitude dropped substantially after application of CPA, whereas the entry flux (L-type current) amplitude changed only a little. Little further change was observed when applying caffeine in addition to CPA, indicating that the CPA treatment had already released most of the Ca²⁺ stored in the SR, consistent with results in C2C12 myotubes (Schuhmeier and Melzer, 2004). In the other cases (columns 2–4), the relative change in input flux was considerably smaller on CPA application and showed only small further changes on application of caffeine. The fractions of the initial gain that remained after CPA application showed mean values of 12%, 58%, 54%, and 65% for _1S, _1C, _1A, and ₁Aas(1592-clip), respectively. After caffeine application the fractional values were 8.7%, 64%, 56%, and 60%, respectively.

Fig. 7 presents the mean time course of Ca²⁺ input flux and Ca²⁺ entry flux induced by the +30 mV step depolarizations averaged over the intervals a, b, and c indicated in Fig. 6 B, respectively. During depletion, Ca²⁺ input flux remained phasic (i.e., exhibits a peak) in _1C, _1A, and ₁Aas(1592-clip) but loses the peak in _1S (see inset). Fig. 8 evaluates both peaks (A) and end level (values at 95 ms, B) by plotting Ca²⁺ input flux versus Ca²⁺ entry flux. The dotted lines indicate a ratio (i.e., gain) of 1. In all four cases, the gain at the pulse end approaches unity after the CPA/caffeine treatment, as would be expected for full SR depletion and a correct description of cytoplasmic Ca²⁺ binding by the model. The reason for the larger deviation from unity gain in CPA/caffeine observed at the peak (Fig. 8 A) is unclear, but might be a remaining small reuptake activity of the SR that permits a transient residual Ca²⁺ release at the onset of the depolarization.

The records in Fig. 7, column 3 (c), represent, for each series of experiments, the maximum of depletion that could be obtained under these conditions. By subtracting this component from the record in column 1 (a) we calculated the fraction of the control flux that is eliminated by CPA and caffeine (Fig. 7, column 4 (d)). Before subtracting the full-depletion response (c), we compensated for the rundown in Ca²⁺ entry flux by multiplying entry and input flux with scaling factors to obtain equal entry flux amplitudes for each row. The depletion-sensitive component showed two phases, a rapidly and a slowly declining one. The rapidly declining phase, i.e., the peak above the dashed line in Fig. 7, column 4 (d), was largest in _1S (A), intermediate in _1C (B), and small in ₁Aas(1592-clip) (D). It was absent in _1A (C). The slow component (indicated by the dashed lines) was similar in size for _1C, _1A, and ₁Aas(1592-clip), but considerably larger for _1S. A tentative interpretation for these observations will be given in the Discussion in conjunction with Fig. 9.

View larger version (19K): [in this window] [in a new window]   FIGURE 9  Ca2+ mobilization from extra- and intracellular sources by voltage-dependent Ca2+ channels of different properties. The scheme summarizes the interpretation of experimental data obtained in this study. (A) The skeletal muscle-type 1S responds to depolarization with an intramolecular charge movement that initiates direct conformational coupling to the RyR1 causing voltage-dependent Ca2+ release (VDCR). This leads to a primary Ca2+ signal in the junctional gap. The small and slow Ca2+ inward current plays a negligible role in Ca2+ mobilization and is omitted in the scheme. The released Ca2+ enhances further Ca2+ release leading to a secondary Ca2+-induced Ca2+ release flux component (CICR) that has phasic and tonic components. VDCR and CICR together make up the Ca2+ input flux. (B) The cardiac muscle-type DHPR likewise responds to depolarization with an intramolecular charge movement that also results in a primary Ca2+ signal but by voltage-induced Ca2+ inward current (Ca2+ entry) instead of VDCR. This also leads to CICR, which, together with the Ca2+ entry flux, makes up total Ca2+ input flux. (C) The brain 1A-subunit, which is not positioned close to the ryanodine receptors, generates a rapid global Ca2+ entry flux signal, although probably a much slower and smaller primary junctional Ca2+ signal than 1S or 1C, leading to ineffective CICR (lacking the phasic component). 1Aas(1592-clip) shows a phasic component unlike 1A but not as large as in 1S or 1C, probably because not enough 1Aas(1592-clip) subunits become positioned close to the RyRs. See Discussion for further explanations.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

Comparison between _1S and _1C
The _1S channels generated Ca²⁺ input flux with a time course similar to that found in normal mammalian myotubes (Dietze et al., 1998; Ursu et al., 2001; Schuhmeier et al., 2003; Schuhmeier and Melzer, 2004) and mature muscle fibers (Delbono and Stefani, 1993; García and Schneider, 1993; Shirokova et al., 1996; Csernoch et al., 1999a,b; Ursu et al., 2004), providing evidence that the rescued Ca²⁺ release in this expression system shows the typical kinetic hallmarks of skeletal muscle type EC coupling, including the rapid partial inactivation (Melzer et al., 1984).

Despite the fundamentally different mechanisms of EC coupling in _1S- and _1C-expressing cells, the time courses of Ca²⁺ input flux were surprisingly similar (Fig. 7). As a major difference, depletion of the SR reduced primarily the phasic component in _1C, whereas it eliminated both phasic and tonic components in _1S (Fig. 7, (d), B and A, respectively). The reason is that the tonic component of _1C-expressing cells results largely from the Ca²⁺ inward current, whereas in _1S-expressing cells it is a flux from the SR, controlled by the DHPR voltage sensor (Csernoch et al., 1993). This is depicted in the idealized scheme of Fig. 9 (A and B). In both _1S- and _1C-expressing GLT myotubes, a pedestal flux of Ca²⁺ is activated by the membrane depolarization. Yet, in the _1C case this flux component enters the myoplasm from the extracellular space (Fig. 9 B), whereas in the case of _1S it originates from the sarcoplasmic reticulum (Fig. 9 A, VDCR).

Both rapid synchronous activation by conformational coupling in _1S-expressing cells and rapid voltage activation of Ca²⁺ entry in _1C-expressing cells likely cause similar fast primary Ca²⁺ signals in the junctional gap that probably lead to similar phasic components of CICR (Fig. 7).

Comparison between _1C- and _1A-controlled Ca²⁺ fluxes
Strong differences had been found between _1C- and _1A-expressing dysgenic myotubes in their responsiveness to extracellular electrical stimulation: Whereas _1C-expressing cells responded frequently with a Ca²⁺ transient or contraction, _1A-expressing cells responded only rarely (Adams et al., 1994; Flucher et al., 2000). The similar size of the fluxes obtained in the two groups of cells in our voltage-clamp experiments seemed, therefore, surprising (Fig. 5). At closer look, however, differences could be detected. The Ca²⁺ input flux component in _1A cells that is sensitive to SR depletion by CPA and caffeine (Fig. 7 C, d) lacks the pronounced peak seen in the corresponding component of _1C cells (Fig. 7 B, d). In addition, the voltage-dependence of the Ca²⁺ signals in _1C cells was different. It was shifted to more negative potentials, with respect to the voltage-dependence of Ca²⁺ inward current activation (Fig. 4 B). This is a characteristic also found in heart cells and has been attributed to the stronger driving force for Ca²⁺ entry at more negative potentials leading to a higher gain of Ca²⁺-induced-Ca²⁺ release (Wier et al., 1994). It is a consequence of the local control of Ca²⁺-induced Ca²⁺ release that depends on local single-channel currents rather than on global Ca²⁺-current amplitudes (Cheng and Wang, 2002). In contrast, in _1A-expressing cells the voltage-dependence of Ca²⁺ signal and Ca²⁺ current was more similar (Figs. 3 and 4 C). These results point to the presence of a more efficient Ca²⁺-induced Ca²⁺-release component in _1C-expressing cells compared to _1A-expressing cells.

Because the transient CPA- and caffeine-sensitive flux component present in _1C but not in _1A cells occurs at the beginning of the depolarizing voltage step, it may be responsible for the much more frequent appearance of Ca²⁺ signals upon short extracellular stimuli in _1C-expressing myotubes (Flucher et al., 2000). As suggested by Flucher et al. (2000), a plausible reason is the specific targeting of _1C to the sarcolemma-SR junctions. Another possibility for the lower responsiveness of _1A-expressing myotubes would be a higher failure rate to elicit all-or-none action potentials, even though it is difficult to see how such a failure would come about.

Gain determinations
The amplification process in Ca²⁺-induced Ca²⁺ release has been studied in detail in cardiac myocytes and is usually quantified by calculating an EC coupling gain factor (for review, see Bers, 2001). Previous determinations of global gain derived from whole-cell measurements in heart cells were based on estimates of integral (total) Ca²⁺ (e.g., Shannon et al., 2000) or of the corresponding Ca²⁺ fluxes (e.g., Wier et al., 1994). The latter approach was also used in the present study and allowed a quantitative comparison of Ca²⁺ entry flux and total Ca²⁺ input flux to the myoplasm.

As a second approach to estimating amplification by Ca²⁺ release, we compared the change in intracellular Ca²⁺ caused by identical trigger pulses before and after depleting the SR of its stored Ca²⁺. In our experiments, the Ca²⁺ input flux controlled by the _1S-subunit was almost completely suppressed by the depletion procedure, demonstrating that it originated almost exclusively from SR Ca²⁺ release. In the presence of the other Ca_V channels tested, the Ca²⁺ current itself made a relatively large contribution to the total Ca²⁺ input flux, but not even in the case of _1A was secondary Ca²⁺ release completely absent.

Our experiments indicated a lower gain in _1C-transfected GLT cells (+30 mV) than in mature heart cells, for which values close to 10 have been estimated in a voltage range of comparable fractional activation (Wier et al., 1994). Reasons might be a higher coupling fidelity of RyR2 in cardiac myocytes compared to RyR1 in skeletal myotubes or a steeper Ca²⁺ gradient produced by the SERCA pump. In heart cells, gain was shown to exhibit a strong dependence on the concentration of Ca²⁺ in the SR lumen (Shannon et al., 2000). In the myotubes, a rather low rate of uptake, likely associated with establishing a smaller gradient, is indicated by the slow change of the signal at the end of a depolarization, which has been observed by us and others (García and Beam, 1994). However, strong differences in SR loading between the four constructs tested is unlikely, since a similar response on application of CPA/caffeine could be observed (see Fig. 6 B).

₁Aas(1592-clip) characteristics
Surprisingly, ₁Aas(1592-clip)-expressing cells did not show a fundamentally different pattern than _1A-type cells despite the observed differences in junctional targeting. Both Ca²⁺ inward current and Ca²⁺ input flux were larger than for _1A, but regarding gain they can at best be described as intermediate between _1A and _1C cells. The smaller gain at larger current density is reminiscent of the situation in heart cells where ß-adrenergic up-regulation of Ca²⁺ inward current does not increase the Ca²⁺ signal proportionally (Song et al., 2001). A fast CPA/caffeine-sensitive component was detectable in ₁Aas(1592-clip) (Fig. 7 D, d), but its amplitude was smaller than in _1C. Also, the voltage-dependence of Ca²⁺ inward current and Ca²⁺ signal (Figs. 3 and 4) resembled more _1A than _1C. The significantly larger Ca²⁺ input flux in ₁Aas(1592-clip) compared to _1A resulted mainly from the larger Ca²⁺ entry flux, not from an increase in gain.

From the present and previous results it seems that the C-terminal signal sequence of the L-type channel introduced into ₁Aas(1592-clip) improves the level of expression but may not establish full _1C-type Ca²⁺-induced Ca²⁺ release despite the junctional targeting. In dual immunostaining experiments for the localization of ₁-subunits and RyRs, a cell is labeled colocalized if regions of colocalization are detectable (Flucher et al., 2000). This criterion makes a quantitative comparison with electrophysiological measurements difficult, because an uncertain percentage of the Ca_V channels that participate in the functional signals may not be colocalized with RyR1. Moreover, the density of channels in the junction, a crucial determinant for establishing efficient cardiac type EC coupling, may well be considerably smaller in ₁Aas(1592-clip)-expressing cells than in _1C cells. The global Ca²⁺ current density was comparable, but the open probability of _1A in dysgenic myotubes is thought to be higher than that of _1C. This follows from the much larger whole-cell current measured per voltage-sensor charge movements (Adams et al., 1994). Longer openings provide more local Ca²⁺ per channel. However, it appears to be the amount of Ca²⁺ supplied immediately on opening of a Ca_V1.2 channel that triggers Ca²⁺ release (Song et al., 2001). Therefore, longer openings are not necessarily more effective than short openings and are unlikely to compensate for a lower channel density in the junction. Ca²⁺ channels that are not specifically targeted, such as the _1A channels, may generate a slowly rising junctional Ca²⁺ transient from diffusional delays, which may still cause a secondary efflux of Ca²⁺ from the SR (CICR in Fig. 7 C). However, this release is less synchronized, explaining the different shape of the CPA/caffeine-sensitive component in Fig. 7 C (d), which lacks a peak. ₁Aas(1592-clip) which is targeted to the junction may not reach a sufficient density there to provide the same local trigger flux of Ca²⁺ as _1C. This may be the reason why cells expressing this isoform show a behavior intermediate between Fig. 9, B and C.

	CONCLUSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
In summary, we compared, in the same cellular expression system, properties of interaction of different Ca_V channel ₁-subunits with the ryanodine receptor RyR1. The observations made here in voltage-clamped myotubes helped us to reconcile previous results from immunocytochemical localization studies and measurements of action-potential stimulated Ca²⁺ transients. In these studies, introducing the putative junctional targeting sequence into _1A had been shown to increase the response frequency approximately ninefold, but not nearly to the level of _1C-expressing cells (140-fold; see Flucher et al., 2000). We conclude that these differences result from different junctional channel densities and single-channel gating properties leading to characteristic local Ca²⁺ signals near the RyRs. To investigate the local functional properties of Ca²⁺ signaling of heterologously expressed channels in further detail will require more refined methods. A promising recent approach has been the recording of Ca²⁺ currents, i.e., sparklets and sparks from small junctional regions of patch-clamped cardiac myocytes (Wang et al., 2001).

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSION ACKNOWLEDGEMENTS REFERENCES

 
We thank Dr. F. Lehmann-Horn for his efforts as Research Training Network coordinator, E. Schoch for help in constructing setup components, and Dr. K. Föhr for providing software for the calculation of binding equilibria. We also thank W. Fritz and E. Schmid for help with the carbon-coating of coverslips and U. Pika-Hartlaub and S. Schäfer for excellent technical help with cell culture and solutions.

The work was supported by a research grant of the Deutsche Forschungsgemeinschaft (Me 713/10-3) to W.M., a training grant of the European Commission (HPRN-CT-2002-00331) to W.M. and B.E.F., and grants from the Austrian Science Fund and the Austrian National Bank (P16532-B05 and P16098-B11, to B.E.F. and M.G., respectively).

Submitted on August 13, 2004; accepted for publication December 22, 2004.

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