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Cardiac-type EC-Coupling in Dysgenic Myotubes Restored with Ca²⁺ Channel Subunit Isoforms _1C and _1D Does not Correlate with Current Density

Nicole Kasielke ^*, Gerald J. Obermair , Gerlinde Kugler ^*, Manfred Grabner ^* and Bernhard E. Flucher ^* 

^* Department of Biochemical Pharmacology and Department of Physiology, University of Innsbruck, A-6020 Innsbruck, Austria

Correspondence: Address reprint requests to Dr. Bernhard E. Flucher, Dept. of Physiology, University of Innsbruck, Fritz-Pregl-Str. 3, A-6020 Innsbruck, Austria. Tel.: +43-512-507-3787; Fax: +43-512-507-2836; E-mail: bernhard.e.flucher{at}uibk.ac.at.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Ca²⁺-induced Ca²⁺-release (CICR)—the mechanism of cardiac excitation-contraction (EC) coupling—also contributes to skeletal muscle contraction; however, its properties are still poorly understood. CICR in skeletal muscle can be induced independently of direct, calcium-independent activation of sarcoplasmic reticulum Ca²⁺ release, by reconstituting dysgenic myotubes with the cardiac Ca²⁺ channel _1C (Ca_V1.2) subunit. Ca²⁺ influx through _1C provides the trigger for opening the sarcoplasmic reticulum Ca²⁺ release channels. Here we show that also the Ca²⁺ channel _1D isoform (Ca_V1.3) can restore cardiac-type EC-coupling. GFP-_1D expressed in dysgenic myotubes is correctly targeted into the triad junctions and generates action potential-induced Ca²⁺ transients with the same efficiency as GFP-_1C despite threefold smaller Ca²⁺ currents. In contrast, GFP-_1A, which generates large currents but is not targeted into triads, rarely restores action potential-induced Ca²⁺ transients. Thus, cardiac-type EC-coupling in skeletal myotubes depends primarily on the correct targeting of the voltage-gated Ca²⁺ channels and less on their current size. Combined patch-clamp/fluo-4 Ca²⁺ recordings revealed that the induction of Ca²⁺ transients and their maximal amplitudes are independent of the different current densities of GFP-_1C and GFP-_1D. These properties of cardiac-type EC-coupling in dysgenic myotubes are consistent with a CICR mechanism under the control of local Ca²⁺ gradients in the triad junctions.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Excitation-contraction (EC) coupling in muscle depends on the close interaction of a voltage-gated Ca²⁺ channel in the t-tubule or the plasma membrane with a Ca²⁺ release channel (ryanodine receptor, RyR) in the sarcoplasmic reticulum (SR). In cardiac EC-coupling this interaction is mediated by Ca²⁺ entering the cell through a voltage-gated Ca²⁺ channel that in turn activates Ca²⁺ release from the SR. In skeletal muscle, SR Ca²⁺ release is activated in the absence of Ca²⁺ influx, presumably by a physical interaction of the two Ca²⁺ channels. Under experimental conditions, the skeletal muscle Ca²⁺ release channel (RyR1) can also be activated by µM concentrations of cytoplasmic free Ca²⁺ (Nagasaki and Kasai, 1983; Smith et al., 1986). Whether Ca²⁺-induced Ca²⁺-release (CICR) in skeletal muscle occurs under physiological conditions, and if so, to what extent it contributes to skeletal muscle EC-coupling is still not resolved.

Reconstitution of Ca²⁺ channel null-mutant skeletal myotubes with wild-type and mutant heterologous Ca²⁺ channels has been a potent tool for analyzing the mechanism of EC-coupling in an intact muscle system. Normal function can be restored in dysgenic myotubes, which lack the skeletal muscle Ca²⁺ channel _1S subunit, by heterologous expression of _1S (Tanabe et al., 1988). When dysgenic myotubes are reconstituted with the cardiac Ca²⁺ channel _1C subunit, "cardiac-type" EC-coupling is restored, which is characterized by its dependence on Ca²⁺ influx (Tanabe et al., 1990). Presumably the skeletal muscle Ca²⁺ release channel is activated by trigger Ca²⁺ entering the myotube through _1C. However, not all examined Ca²⁺ channel isoforms that produced Ca²⁺ currents when expressed in dysgenic myotubes also triggered CICR from the SR. The neuronal isoform _1A, which is not targeted into triads of dysgenic myotubes, only very rarely displayed evoked contractions (Adams et al., 1994) or Ca²⁺ transients in response to electrically induced action potentials (action potential-induced Ca²⁺ transients) (Flucher et al., 2000). Apparently the correct subcellular distribution of the channel is important for the activation of cardiac-type EC-coupling in skeletal myotubes. Skeletal-type EC-coupling properties could be conferred onto the cardiac _1C subunit by replacing a short sequence of the cytoplasmic loop connecting the homologous repeats II and III with that of _1S (Nakai et al., 1998). Thus, the chimera CSk53 was created, which combines skeletal-type activation of EC-coupling with cardiac current properties.

If CICR participates in normal skeletal muscle EC-coupling, it is unlikely that the trigger Ca²⁺ comes from the Ca²⁺ currents across the sarcolemma, because the slow activation of the skeletal Ca²⁺ current lags behind activation of SR calcium release (Garcia et al., 1989; Feldmeyer et al., 1990). Instead, trigger Ca²⁺ must be provided by Ca²⁺ released from the SR by the population of the skeletal muscle RyR1 that is under the direct control of the voltage-sensor _1S subunit (reviewed in Rios and Stern, 1997). In other words, Ca²⁺ released through the RyR1 provides a positive feedback resulting in release of more Ca²⁺ from the same or adjacent RyRs. A recent report by O'Brien et al. (2002) indicates that, whereas CICR appears not to be necessary for activation of EC-coupling in skeletal muscle, SR Ca²⁺ release is reduced by fivefold when Ca²⁺ activation of RyR1 is impeded. This indicates a significant contribution of CICR to skeletal muscle EC-coupling.

Here we studied the properties of CICR in skeletal myotubes separate from skeletal-type activation of Ca²⁺ release by expressing nonskeletal ₁ subunits with distinct current characteristics and with distinct targeting characteristics in dysgenic myotubes. Analyzing the capability of different types of Ca²⁺ channels to generate action potential-induced and voltage-clamp-induced Ca²⁺ transients indicates that in the absence of direct skeletal coupling, Ca²⁺ currents of similar size as those of the skeletal _1S are sufficient to trigger CICR. Moreover, the results indicate that activation of SR Ca²⁺ release seems to depend less on the magnitude of the whole-cell current than on the correct localization of the channels in the triads.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Cell culture and transfections
Myotubes of the homozygous dysgenic (mdg/mdg) cell line GLT were cultured as described in Powell et al. (1996). At the onset of myoblast fusion (2 days after addition of differentiation medium), GLT cultures were transfected using FuGene transfection reagent (Roche, Basel, Switzerland). Cultures were analyzed 3–5 days after transfection.

₁ subunit constructs
The following GFP-tagged Ca²⁺ channel constructs have been used: _1S, _1A, _1C (Grabner et al., 1998), _1D (Koschak et al., 2001), and _1G (Monteil et al., 2000). GFP-CSk53: The BamHI-EcoRV fragment (nt 1265–4351; _1C numbering) of clone CSk53 (Nakai et al., 1998), where part of the II-III loop coding _1C cDNA sequence (nt 2549–2690) was replaced by _1S sequence (nt 2156–2297), was ligated into the corresponding restriction enzyme sites of clone GFP-_1C (Grabner et al., 1998) resulting in an N-terminally GFP-tagged CSk53.

GFP and immunofluorescence labeling
Differentiated GLT cultures were fixed and immunostained as previously described (Flucher et al., 1994; Flucher et al., 2000). For double-immunofluorescence labeling we used an affinity-purified anti-GFP antibody (Molecular Probes, Eugene, OR) at a final dilution of 1:4000 and the affinity-purified antibody 162 (Giannini et al., 1995) against RyR1 at a dilution of 1:5000. Alexa-488- and Alexa-594-conjugated secondary antibodies (Molecular Probes) at dilutions of 1:4000 were used to achieve a wide separation of the fluorescence signals. Alexa-488 was usually used with the anti-GFP antibody so that the antibody label and the intrinsic GFP signal were both recorded in the green channel. Controls, for example multiple combinations of primary and secondary antibodies and the omission of primary antibodies, were routinely performed.

Images of the double labeling experiments were recorded on an Axiophot microscope (Carl Zeiss, Jena, Germany) using a cooled CCD camera and Meta View image processing software (Universal Imaging Corporation, West Chester, PA). Quantitative analysis of the labeling patterns was performed by systematically screening the cover glasses for transfected myotubes using a 63x objective. The labeling pattern in transfected myotubes with more than two nuclei was classified as "clustered" when punctuate ₁ subunit fluorescence was co-localized with a similar RyR1 label (Fig. 1). The counts were obtained from several samples of at least three different experiments.

View larger version (77K): [in this window] [in a new window]   FIGURE 1  The L-type Ca2+ channel subunit GFP-1D is targeted into skeletal muscle triads of dysgenic myotubes. Dysgenic myotubes transfected with GFP-1D were double immunofluorescence labeled with antibodies against GFP (top) and against the skeletal muscle RyR1 (middle). GFP-1D is distributed in clusters co-localized with RyR1 (examples indicated by arrows), which indicates a localization of GFP-1D in junctions of the SR with the plasma membrane or the t-tubules (triads). The color overlay (bottom) shows the co-localization of GFP-1D clusters (green) and RyR1 clusters (red) as yellow foci. The inset shows a fourfold enlarged area. Bar, 10 µm.

Leak currents were digitally subtracted by a P/4 prepulse protocol. Recordings were low-pass Bessel filtered at 1 kHz and sampled at 5 kHz. Currents were determined with 200 ms depolarizing steps from a holding potential of -80 mV to test potentials between -40 and +80 mV in 10 mV increments. Test pulses were preceded by a 1-s prepulse to -30 mV to inactivate endogenous T-type Ca²⁺ currents (Adams et al., 1990). Current recordings of cells transfected with the T-type Ca²⁺ channel _1G subunit were measured with 200 ms depolarizing steps from a holding potential of -90 mV to test potentials between -80 mV and +80 mV in 10 mV increments. Ca²⁺ current densities were normalized by linear cell capacitance and expressed in pA/pF.

Action potential–induced Ca²⁺ transients were recorded in cultures loaded for 45–60 min at room temperature with 5 µM fluo-4-AM plus 0.1% Pluronic F-127 (Molecular Probes) in HEPES and bicarbonate-buffered DME, as previously described (Flucher et al., 1993; Powell et al., 1996). Action potentials were elicited by passing 1 ms pulses of 30 V across the 19-mm incubation chamber. 0.5 mM Cd²⁺ and 0.1 mM La³⁺ were added to block Ca²⁺ influx and therefore allow discrimination between CICR and skeletal-type EC coupling. Application of 6 mM caffeine to the bath solution resulted in rapid Ca²⁺ release and thus proved the capacity of SR Ca²⁺ release.

Statistics
Data were expressed as mean ± SE where n is the number of cells examined. Data analysis, unpaired Student's t-test, and one-way-ANOVA analysis followed by the Tukey post test were performed with Clampfit 8.0 (Axon instruments, Union City, CA, USA) and GraphPad Prism software (GraphPad Inc., San Diego, CA, USA). P < 0.05 was considered statistically significant.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
It has previously been demonstrated that heterologous expression of different muscle ₁ subunit isoforms in dysgenic myotubes restores EC-coupling by two distinct mechanisms. Whereas the skeletal muscle _1S triggers SR Ca²⁺ release independently of Ca²⁺ influx through the L-type Ca²⁺ channel, the cardiac _1C depends on Ca²⁺ influx for activation of EC-coupling, presumably by CICR (Garcia et al., 1994). In contrast to the two muscle isoforms _1S and _1C, the neuronal non-L-type _1A subunit was not capable of restoring EC-coupling in dysgenic myotubes, although _1A generated sizable Ca²⁺ currents (Adams et al., 1994; Flucher et al., 2000). Here we expressed _1S, _1C, _1A, and for the first time the L-type Ca²⁺ channel ₁ subunit isoform _1D in dysgenic myotubes to compare their current properties and targeting characteristics with their ability to restore EC-coupling.

GFP-_1D is targeted into triads of dysgenic myotubes
Immunofluorescence analysis (Fig. 1) shows that a fusion protein of GFP attached to the N-terminus of _1D (GFP-_1D) is efficiently expressed in myotubes of the dysgenic cell line GLT and is distributed in a clustered pattern. Double labeling of GFP-_1D and RyR1 demonstrated that GFP-_1D clusters were co-localized with clusters of RyR1, thus identifying the clusters as t-tubule/SR or plasma membrane/SR junctions, all of which will from here on be called triads. GFP-_1D/RyR1 coclustering was found in 62% of 454 analyzed GFP-_1D-expressing myotubes, which was similar to the degree of clustering obtained with the native _1S or with _1C (see also Fig. 4 C). Myotubes in which GFP-_1D was expressed in a nonclustered pattern typically showed ER/SR labeling (not shown), presumably because these myotubes were still very immature or due to unbalanced expression of the heterologous channel and endogenous muscle proteins (Flucher et al., 1994). Localizing GFP-_1D (Fig. 1, green) in clusters coincident with the RyR (Fig. 1, red), makes _1D the third member of the L-type Ca²⁺ channel family that, when expressed in skeletal muscle cells, is targeted into triads.

View larger version (14K): [in this window] [in a new window]   FIGURE 4  The ability of Ca2+ channel isoforms to restore EC-coupling compared to their current densities and targeting properties. (A) Myotubes responding to field stimulation with Ca2+ transients (see Fig. 3) were counted in cultures transfected with GFP-1D, GFP-1C, GFP-1A, and GFP-1S. Counts from seven experiments are expressed in percent (mean ± SE) with the values of GFP-1D normalized to 100%. All three L-type channels (GFP-1D, GFP-1C, GFP-1S) restored EC-coupling with similar efficiency, whereas with GFP-1A only a single responding cell was observed. (B) Average peak current densities shown for the same channel isoforms as above (see also Table 1). Although the nonskeletal 1 isoforms activate action potential-induced Ca2+ transients by CICR, no correlation of current density and restoration of EC-coupling exists (cf. A and B). (C) Fraction of myotubes in which the channel isoforms were targeted into the triads (see Fig. 1). The triad targeting properties of the Ca2+ channel isoforms exactly correspond to their ability to restore EC-coupling (cf. A and C).

View larger version (30K): [in this window] [in a new window]   FIGURE 2  Current properties of GFP-1D expressed in dysgenic myotubes compared to those of GFP-1S and GFP-1C. (A) Voltage dependence of peak Ca2+ current densities (pA/pF; mean ± SE) of GFP-1D (; n = 42), GFP-1S (; n = 33), and GFP-1C (•; n = 28). Maximal current density of GFP-1D is similar to that of GFP-1S and about four times lower than that of GFP-1C. (B) Normalized I/V curves show that GFP-1D activates at 13 mV more negative potentials than GFP-1C and at 37 mV more negative potentials than GFP-1S (for values of half maximal activation, see Table 1). (C) Representative whole-cell current recordings of GFP-1D, GFP-1S, and GFP-1C expressed in dysgenic myotubes. Cells were held at -80 mV and after a prepulse to inactivate low-voltage activated currents test pulses to increasing voltages from -40 to +80 mV were applied; currents were recorded in 10 mM Ca2+. GFP-1D activates faster than GFP-1S.

GFP-_1D restores Ca²⁺ currents in dysgenic myotubes

View larger version (32K): [in this window] [in a new window]   FIGURE 3  Restoration of action potential-induced Ca2+ transients by GFP-1D in dysgenic myotubes. Field stimulation at low frequency (30 V, 1 ms, 0.3 Hz/0.5 Hz) evoked rapid Ca2+ transients in myotubes expressing GFP-1D, GFP-1C, or GFP-1S. Bath application of 0.5 mM Cd2+/0.1 mM La3+ immediately stopped the activation of transients in myotubes expressing GFP-1D (A) and GFP-1C (B), indicating that the transients were dependent on Ca2+ influx through the L-type channels. Addition of 6 mM caffeine (A, right trace) triggered a strong Ca2+ transient, proving that SR Ca2+ release was still functional during the Cd2+/La3+ block. (C) Persistent Ca2+ transients after addition of Cd2+/La3+ in myotubes expressing GFP-1S characterizes skeletal-type EC-coupling, which is independent of Ca2+ influx. In contrast, GFP-1D restored action potential-induced Ca2+ transients with cardiac characteristics. Marks underneath the traces indicate the approximate times of stimulation; bars indicate periods of Cd2+/La3+ and caffeine application.

GFP-_1D restores cardiac-type EC-coupling in dysgenic myotubes

Cultures transfected with GFP-_1D responded to field stimulation with brief Ca²⁺ transients (Fig. 3). The transients were characterized by a rapid upstroke that terminated abruptly, indicative of a strong voltage-dependence of both, the activation and the deactivation of the EC-coupling mechanism. The average amplitudes of the transients reached a (F/F)_max of 0.46 ± 0.04, which was not significantly different from the amplitudes recorded with GFP-_1C or GFP-_1S (Table 2). The speed of the upstroke and the decay of the transients were similar between GFP-_1D and GFP-_1C; however, the average values obtained with these channel isoforms were significantly slower than those of GFP-_1S. Unless the expression of nonskeletal channels adversely affects the overall differentiation of the myotubes, for which there is no independent evidence based on the morphology of the myotubes and on the expression of proteins seen with immunocytochemistry, this difference in the time course of the transients probably reflects the different modes of EC-coupling (see below). But most important, despite the differences observed in the whole-cell current properties (Fig. 2), time-to-peak, amplitudes, and time constant of the decline of the Ca²⁺ transients were not significantly different (P>0.05) between GFP-_1D and GFP-_1C (Table 2).

Restoration of cardiac-type EC-coupling depends on triad targeting but not on the magnitude of the current density
In the field stimulation experiments GFP-_1D was as efficient as GFP-_1C in restoring action potential-induced Ca²⁺ transients in dysgenic myotubes. In Fig. 4 A we compare the frequencies at which action potential-induced Ca²⁺ transients were observed in cultures transfected with GFP-_1D, GFP-_1C, GFP-_1A, and GFP-_1S. The number of responding cells in cultures transfected with GFP-_1D was not significantly different from those of GFP-_1C and GFP-_1S. As previously shown (Flucher et al., 2000), GFP-_1A very rarely produced action potential-induced Ca²⁺ transients, probably due to the fact that this channel is not targeted into the triads. Out of six culture dishes transfected with GFP-_1A, only a single myotube in one dish responded with an action potential-induced Ca²⁺ transient; therefore, the bar is not visible at this scale (Fig. 4 A). Comparing the capability of each of these channel isoforms to restore EC-coupling with the average densities of their Ca²⁺ currents shows no correlation of the two parameters (cf. Fig. 4, A and B). For GFP-_1S, which directly activates SR Ca²⁺ release, such a correlation would not have been expected. However, for the constructs which elicit cardiac-type EC-coupling—presumably by CICR—a correlation of current density and their ability to activate action potential-induced Ca²⁺ transients was to be expected. Most interestingly, GFP-_1D, with a current density of less than a third of that of GFP-_1C, activated EC-coupling just as efficiently as the cardiac channel. In contrast to current density, restoration of EC-coupling correlated very well with the triad targeting properties of the individual channel isoforms (cf. Fig. 4, A and C). Those constructs which are targeted into the triads (GFP-_1S, GFP-_1C, GFP-_1D) all activated EC-coupling with similar efficiency. However GFP-_1A, which generated the largest current density of all examined channel isoforms (see also Table 1) but is not targeted into triads, hardly ever activated action potential-induced Ca²⁺ transients in dysgenic myotubes. Thus, for the activation of cardiac-type EC-coupling, the spatial arrangement of the Ca²⁺ channels opposite the RyR appears to be more important than the pure size of the Ca²⁺ currents.

Ca²⁺ transients activated by GFP-_1C and GFP-_1D are qualitatively and quantitatively different from those activated by GFP-_1S
To further characterize the dependence of cardiac-type EC-coupling on Ca²⁺ currents we performed combined patch-clamp and fluo-4 Ca²⁺ recording experiments in dysgenic myotubes expressing different ₁ subunit isoforms. This approach enables us to simultaneously monitor the whole-cell Ca²⁺ influx and intracellular Ca²⁺ transients in the same myotubes, and it allows us to analyze the voltage-dependence of Ca²⁺ transients. But these experiments differ from the analysis of action potential-induced Ca²⁺ transients in several important aspects: 1), Sampling—whereas with field stimulation we see and record only those myotubes that have achieved excitability and a good degree of EC-coupling (probably the peak of the population of expressing cells), with the patch-clamp approach we get a random sample of differently well developed and expressing myotubes; 2), Stimulation—instead of a 1-ms extracellular current pulse, which stimulates an action potential lasting only a few milliseconds, in the patch-clamp experiments we depolarize the cells for 200 ms giving rise to unnaturally long periods of Ca²⁺ influx; and 3), Intracellular milieu—while the myotube is being loaded with the Ca²⁺ indicator through the patch pipette, the cells are dialyzed and Ca²⁺ buffers and the entire intracellular milieu are altered. Thus, in this set of experiments we trade in better control and quantitative analysis for less physiological conditions. This is reflected in the shape of the Ca²⁺ transients in response to the 200-ms depolarization (Fig. 5 C), which does not resemble that of action potential-induced Ca²⁺ transients (Fig. 3).

View larger version (26K): [in this window] [in a new window]   FIGURE 5  Simultaneous recording of whole-cell Ca2+ currents and Ca2+ transients in dysgenic myotubes transfected with skeletal and nonskeletal 1 subunit isoforms. (A, B) Voltage-dependence of Ca2+ transients (A) and of Ca2+ currents (B). Ca2+ transients of GFP-1S (; n = 34) have a sigmoidal voltage relationship that does not follow the I/V curve. Transient-to-voltage curves for GFP-1C (•; n = 10), GFP-1D (; n = 9), and GFP-1A (; n = 11) are bell-shaped and follow their I/V curves with respect to activation properties but not with respect to amplitudes. Whereas current densities for GFP-1D are significantly smaller than those of GFP-1C (P = 0.0011) or GFP-1A (P = 0.014), their maximal amplitudes of Ca2+ transients are not significantly different from each other (P > 0.05). The dotted line shows the shift in the GFP-1A I/V curve after removing one exceptionally high recording from the analysis. But transients of all nonskeletal isoforms are significantly smaller (P < 0.001) than that of GFP-1S. (C) Representative examples of Ca2+ transients recorded with fluo-4 in parallel to whole-cell currents show the difference between skeletal and nonskeletal 1 isoforms. (D) A scatter plot of the maximal peak current densities versus peak F/F values for GFP-1S (), GFP-1C (•), and GFP-1D () shows that skeletal and nonskeletal Ca2+ signals clearly separate into two distinct groups. Superthreshold Ca2+ transients of GFP-1C and GFP-1D are confined to a narrow window of amplitudes over a wide range of current densities. (E) I/V curves of GFP-1C (•) and GFP-1D () (both the same as in B) compared to the average I/V curves of those currents for each group, which did not evoke Ca2+ transients (, GFP-1C, n = 5; GFP-1D, n = 16). For both channels, currents without transients were relatively smaller than those that evoked transients, but the absolute values differed significantly.

Unexpectedly, also GFP-_1A, which in the field stimulation experiments restored action potential-induced Ca²⁺ transients very rarely (Fig.4 A), frequently produced cardiac-type Ca²⁺ transients in the combined patch-clamp/fluo-4 Ca²⁺ recording experiments. Apparently the mechanism that limited activation of cardiac-type EC-coupling in response to action potential-induced Ca²⁺ transients to the correctly targeted channel isoforms broke down under the conditions of the patch-clamp experiments. But even in this case, the average maximal amplitudes of Ca²⁺ transients were similar to those of GFP-_1C and GFP-_1D (Fig. 5). Whereas the transient-to-voltage relationship for GFP-_1A is somewhat higher than that of GFP-_1C and GFP-_1D (Table 1), this difference is not statistically significant (P > 0.05) and mainly reflects a single recording with an exceptionally high current and transient. If this recording is excluded from the analysis, the transients of GFP-_1A are also in the exact same range as GFP-_1D and GFP-_1C (dotted curve in Fig. 5 A). At this point we have no explanation for the occurrence of such exceptionally strong responders. However, one needs to remember that also in the field stimulation experiments occasional responders were observed with GFP-_1A (see above; Adams et al., 1994; Flucher et al., 2000).

Analysis of the combined patch-clamp/fluo-4 Ca²⁺ recording experiments revealed striking differences between the skeletal and nonskeletal ₁ subunits expressed in dysgenic myotubes. Plotting the amplitudes of the Ca²⁺ transients versus the peak current densities for all individual experiments (Fig. 5 D) shows that dysgenic myotubes reconstituted with GFP-_1S produce Ca²⁺ transients of greatly variable size (F/F values up to 2.60) with no correlation to current densities. Ca²⁺ transients are observed even in myotubes with no detectable currents, and some of the myotubes with the highest current densities showed very small transients. In contrast, GFP-_1C and GFP-_1D display a completely different picture. Data points of both channels spread out over a wide range of current densities but within a relatively narrow window of Ca²⁺ transient amplitudes (F/F values between 0.15 and 0.42). The maximal transient amplitudes of both nonskeletal channel isoforms do not reach those of the skeletal GFP-_1S.

Furthermore, a fraction of GFP-_1C- and GFP-_1D-expressing myotubes showed Ca²⁺ currents but no detectable Ca²⁺ transient (33% and 64% of myotubes expressing GFP-_1C and GFP-_1D, respectively). This was never the case with GFP-_1S. In Fig. 5 E a second set of I/V curves is shown for GFP-_1C and GFP-_1D. In addition to the I/V curve of those myotubes responding with both currents and transients (same as in Fig. 5 B), I/V curves were calculated from those recordings, in which Ca²⁺ currents were not accompanied with Ca²⁺ transients. For both channel isoforms the currents not associated with Ca²⁺ transients were significantly smaller (_1D: P = 0.0001; _1C: P = 0.0180) than those associated with transients. Thus, in both cases the larger currents activated transients whereas the smaller currents did not. This could be interpreted as the existence of a threshold for triggering the Ca²⁺ signal. However, the whole-cell current densities necessary to activate Ca²⁺ transients differed greatly between the two channel isoforms. The amplitudes of Ca²⁺ currents of GFP-_1D that were associated with Ca²⁺ transients were almost identical to those of GFP-_1C that were not associated with transients (Fig. 5 E). Thus, whereas current amplitudes matter for the activation of Ca²⁺ release in skeletal myotubes, the absolute values of whole-cell current densities do not reliably describe the activation threshold in the triad.

The two distinct groups of Ca²⁺ transient amplitudes do not depend on the skeletal versus cardiac EC-coupling mechanisms
The data presented above show that GFP-_1C and GFP-_1D expressed in dysgenic myotubes give rise to Ca²⁺ transients with an average maximal amplitude at about one third of that activated by GFP-_1S (Table 1). Therefore it was reasonable to assume that the difference in skeletal versus cardiac EC-coupling mechanisms (direct or Ca²⁺-dependent) is responsible for this striking difference. This hypothesis was tested with a channel chimera, GFP-CSk53, which consists of GFP-_1C with a 46-amino-acid sequence in the II-III cytoplasmic loop replaced by the corresponding sequence of _1S (Nakai et al., 1998). This sequence is sufficient to confer skeletal-type EC-coupling properties onto the cardiac channel. Action potential-induced Ca²⁺ transients persist after application of Cd²⁺/La³⁺ to block Ca²⁺ currents (Fig. 6 E). This observation is in agreement with earlier reports showing that GFP-CSk53 activated EC-coupling by the skeletal Ca²⁺-independent mechanism (Nakai et al., 1998). Fig. 6, A–D shows the voltage-dependence of Ca²⁺ transients and currents of GFP-CSk53 compared to those of GFP-_1S and GFP-_1C. Peak current densities similar to those of GFP-_1C (Fig. 6 B; Table 1) demonstrate the normal functional expression of the channel chimera. But surprisingly, on first sight the voltage relationship of the transients of GFP-CSk53 resembled that of the cardiac and not of the skeletal isoforms in both amplitude and shape (Fig. 6 A). However, the analysis of individual recordings from the combined patch-clamp/fluo-4 Ca²⁺ experiments showed that the cardiac-like decline of the transients at voltages above +20 mV was preferentially seen in those recordings where the baseline shifted upward during the duration of the experiment. Dividing the recordings into those with baseline shifts below and those above 10% (Fig. 6 C) revealed that the records with little or no shift displayed the typical skeletal characteristics of the voltage-dependence of the Ca²⁺ transients, i.e., no decline at higher test potentials. The same analysis performed with the recordings of GFP-_1C showed typical cardiac characteristics in both groups (Fig. 6 D). Therefore, the apparent decline of the amplitudes of Ca²⁺ transients at positive potentials was an experimental artifact that could be avoided by excluding the data of recordings with a baseline shift greater than 10%.

View larger version (22K): [in this window] [in a new window]   FIGURE 6  Comparison of the cardiac-skeletal chimera CSk53 with 1C and 1S. The voltage-dependences of Ca2+ transients (A) and of peak current densities (B) of CSk53 (, n = 18) resemble in size and shape those of GFP-1C (•) but not those of GFP-1S (). The bell-shaped (cardiac-type) Ca2+ transient-to-voltage curve of CSk53 is in disagreement with skeletal-type EC-coupling properties as indicated by the insensitivity of action potential-induced Ca2+ transients to Cd2+/La3+ block (E). (C) Dividing the CSk53 records into two groups based on the occurrence and magnitude of a fluorescence baseline shift (above and below 10%) shows that the stable recordings (baseline shift < 10%; , n =7; > 10%, , n = 11) display a sigmoidal transient-to-voltage relationship. (D) In contrast, the transient-to-voltage curves of 1C are bell-shaped regardless of the size of a baseline shift (baseline shift < 10%, C, n = 5; >10%, •, n = 5). Therefore, the bell-shaped appearance of the curve for CSk53 in A is due to this artifact, and the true character of the CSk53 transients is skeletal. (F) A scatter plot of maximal peak current densities versus F/F values emphasizes the difference between Ca2+ transients of CSk53 and GFP-1S. Whereas current densities for CSk53 vary between 1.5 and 40 pA/pF, the amplitudes of the transients do not exceed F/F values of 0.6.

Cardiac-type and skeletal-type Ca²⁺ transients can be individually activated in the same myotube
Finally, we wanted to exclude the possibility that the relatively low Ca²⁺ transients in myotubes transfected with nonskeletal muscle ₁ subunit isoforms and the chimera CSk53 were simply due to a reduced Ca²⁺ release capacity of those myotubes. Therefore we examined whether the same myotube responded with different size Ca²⁺ transients when activated by the native GFP-_1S or by a nonskeletal channel. The low-voltage-activated Ca²⁺ channel GFP-_1G (Monteil et al., 2000) also gave rise to Ca²⁺ transients in dysgenic myotubes. These activated at potentials of -60 mV and peaked at about -40 mV, both at 40 mV more negative potentials than transients in GFP-_1S-expressing myotubes (Fig. 7 A). Similar to the Ca²⁺ transients of all other examined nonskeletal ₁ subunits, GFP-_1G transients reached maximal amplitudes that were substantially smaller than those of GFP-_1S. When coexpressed with GFP-_1S, Ca²⁺ transients activated by GFP-_1G could be well separated from transients activated by the high-voltage-activated GFP-_1S (Fig. 7 C). The transient-to-voltage curve of cotransfected myotubes displayed a shoulder at -30 mV and continued to rise at -10 mV (Fig. 7 A). Subtracting the transient-to-voltage curve of GFP-_1G alone from that of the cotransfection experiments eliminated the shoulder, resulting in a curve with typical skeletal characteristics (Fig. 7 B). Thus, we conclude that the shoulder represents the contribution of GFP-_1G. The height of this shoulder is near F/F values of 0.3, which corresponds to transients observed with GFP-_1C, GFP-_1D, GFP-_1A, and GFP-CSk53, and it is only about one third of the maximal Ca²⁺ signal activated by GFP-_1S in the same cells at more positive voltages. Thus, in our experiments Ca²⁺ transients activated by channels other than the native GFP-_1S peak at amplitudes significantly below that activated by GFP-_1S, even in the same myotube.

View larger version (24K): [in this window] [in a new window]   FIGURE 7  Cardiac-type and skeletal-type EC-coupling activated independently in the same myotube. (A) Voltage-dependence of Ca2+ transients in dysgenic myotubes transfected with GFP-1G alone (; n = 9) and in combination with GFP-1S (; n = 9). The low-voltage activated Ca2+ channel GFP-1G generates cardiac-type Ca2+ transients activated at negative voltages with maximal amplitudes (F/F) not significantly different from those of GFP-1C and GFP-1D (cf. Fig. 5). The I/V curve of cotransfected myotubes displays a shoulder at negative potentials and continues to rise at positive potentials to levels similar to those achieved with GFP-1S alone (; data from Fig. 5 A). Maintained transients at +80 V near the reversal potential characterize this component as skeletal. (B) Subtraction of the I/V curve of GFP-1G from that of the GFP-1G/GFP-1S-cotransfected myotubes results in a curve (•) resembling that of GFP-1S (). (C) Examples of Ca2+ transients and currents from a cotransfected cell depolarized to -10 mV (left traces) and to +80 mV (right traces). Ca2+-dependent (cardiac-type) and Ca2+-independent (skeletal-type) transients can be activated independently from each other in the same cell.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

GFP-_1D, together with the native _1S and the cardiac _1C, is now the third member of the voltage-gated Ca²⁺ channel family that has been demonstrated to become normally targeted into the skeletal muscle triads. Apparently all the members of the L-type subclass so far examined possess the triad targeting signals, whereas the neuronal, non-L-type _1A does not (Flucher et al., 2000). We have previously shown that an important triad targeting signal resides in the C-terminus of _1S and can be conferred onto GFP-_1A by replacing a short C-terminal sequence with the corresponding sequence of _1S (Flucher et al., 2000). Similarly the C-terminus of GFP-_1D was able to confer triad targeting properties onto GFP-_1A (Flucher and Grabner, unpublished results). Thus, triad targeting signals could be a conserved property of the L-type Ca²⁺ channel family. Alternatively, since _1S, _1C, and _1D isoforms are all expressed in various muscle tissues whereas _1A is not, the triad targeting property could be a muscle-specific feature.

Ca²⁺ currents generated by GFP-_1D expressed in dysgenic myotubes showed the typical characteristics previously reported for _1D in native cells (Zidanic and Fuchs, 1995) and of GFP-_1D expressed in mammalian heterologous expression systems (Koschak et al., 2001). GFP-_1D currents activated at 13 mV more negative potentials compared to GFP-_1C. The current density of GFP-_1D in dysgenic myotubes was less than one third of that of GFP-_1C. Nevertheless, both channels restored action potential-induced Ca²⁺ transients in dysgenic myotubes equally well and as efficiently as the native GFP-_1S. The fact that these Ca²⁺ transients were blocked by Cd²⁺/La³⁺ demonstrated that the mechanism by which GFP-_1D activated action potential-induced Ca²⁺ transients was CICR, with the trigger Ca²⁺ provided by the L-type Ca²⁺ current. Therefore it was surprising that the considerable difference in the size of the currents of GFP-_1D and GFP-_1C was not reflected in the size of the transients or in the rate at which action potential-induced Ca²⁺ transients were observed. Thus, activation of CICR is not strongly dependent on the total influx of Ca²⁺ manifested in the whole-cell current densities. This conclusion is further supported by data on _1A collected by us and others (Adams et al., 1994; Flucher et al., 2000; and present study). While _1A expressed in dysgenic myotubes produces high current densities the size of GFP-_1C or larger, it restores contractions or action potential-induced Ca²⁺ transients very rarely.

Whereas the ability of the examined ₁ subunits to trigger Ca²⁺ transients does not correlate with the magnitude of their Ca²⁺ currents, it correlates well with their triad targeting properties. All channel isoforms that are efficiently targeted into the triads (GFP-_1S, GFP-_1C, GFP-_1D) also restore action potential-induced Ca²⁺ transients. On the other hand, GFP-_1A fails to do either. This suggests that activation of EC-coupling by CICR is dependent on local Ca²⁺ concentrations achieved in the restricted space of the triad in the moment of depolarization by an action potential. In the triad, even the small GFP-_1D currents—which are of similar size as those of _1S—attain the concentration necessary for activating SR Ca²⁺ release. However, the Ca²⁺ activation threshold of Ca²⁺ release is not reached during fivefold larger Ca²⁺ currents generated by the nontargeted GFP-_1A channel isoform. A local activation mechanism for CICR is also consistent with the observation that the termination of the Ca²⁺ release during action potential-induced Ca²⁺ transients is under the tight control of the membrane potential. At least in the dysgenic system, CICR is not a regenerative process that is self-sustained by Ca²⁺ released through the RyRs alone. In that case transients would be expected to continue to rise after repolarization and the deactivation of voltage-gated Ca²⁺ channels. On the contrary, the abrupt termination of the transients upon repolarization argues that the sustenance of the CICR requires the continued influx of Ca²⁺ through a voltage-gated channel. This tight control of Ca²⁺ activation of the Ca²⁺ release channels by voltage-gated Ca²⁺ currents in dysgenic myotubes reconstituted with GFP-_1C or GFP-_1D is reminiscent of EC coupling in cardiac myocytes (Cannell et al., 1987; Stern et al., 1999; Bers, 2000) and can only be envisioned in the restricted space of the triad junction, where _1C or _1D come within nanometers of the Ca²⁺ release channel (Rios and Stern, 1997).

A role of local Ca²⁺ gradients in the activation of CICR in skeletal myotubes is further corroborated by the observation that the lower limits of current densities required for triggering Ca²⁺ transients differ between GFP-_1C and GFP-_1D. In both cases failure of activation of Ca²⁺ transients was preferentially observed in myotubes with low current densities, whereas the myotubes with higher current densities usually produced Ca²⁺ transients. However, the current density required for the activation of Ca²⁺ transients was not constant. It varied between individual myotubes, and it was clearly different between GFP-_1C and GFP-_1D. The existence of a threshold of current density below which no Ca²⁺ release would be stimulated is expected for CICR. But this threshold should be the same in myotubes expressing GFP-_1C and GFP-_1D. Thus, we have to conclude that the whole-cell current densities only incompletely reflected the actual Ca²⁺ concentration at the activation site. Either the geometry of the channels in the triad is such that the spatiotemporal Ca²⁺ gradients required for Ca²⁺ activation of the RyR are similar for GFP-_1C and GFP-_1D even though their channel properties differ or, alternatively, the fractions of channels inside and outside the triad might differ for the two ₁ isoforms, so that there is no linear relationship between whole-cell current densities and current densities in the triads.

GFP-_1A, which is not targeted into triads and usually fails to trigger action potential-induced Ca²⁺ transients, generated Ca²⁺ transients in the combined patch-clamp and fluo-4 Ca²⁺ recording experiments. On first sight, this result appears to contradict the hypothesis that triggering cardiac-type EC-coupling by CICR depends on the co-localization of the t-tubular and SR Ca²⁺ channels in the triads. However, if activation of CICR in the triads depends on local Ca²⁺ gradients, these could easily break down during the 200-ms depolarization of the patch-clamp experiments. During a brief action potential locally high Ca²⁺ concentrations near the mouth of a channel may activate SR Ca²⁺ release in the case of channels concentrated in the triad junction, whereas similarly high Ca²⁺ concentrations generated by _1A channels diffusely located outside the triads may dissipate in the highly buffered cytoplasm before reaching the RyRs in the triad. However, during continued activation over the period of 200 ms, Ca²⁺ entering the myotube through _1A may flood the cell to a degree that spatial gradients can no longer be maintained and CICR is triggered even by a channel positioned outside the triad. Interfering with the Ca²⁺ buffering milieu in the patched myotubes may further contribute to this inconsistent behavior under the different experimental conditions. The different shapes of Ca²⁺ transients produced by the field stimulation and in the combined patch-clamp recordings support this explanation (see also Garcia et al., 1994). The fact that in these experiments Ca²⁺ transients do not decline immediately after repolarization suggests that in contrast to action potential-induced Ca²⁺ transients, CICR stimulated by long depolarization contains a regenerative component, which sustains Ca²⁺ release even in the absence of influx. Also the slow decline of the Ca²⁺ transients indicates that the cytoplasmic Ca²⁺ buffering capacity has been exceeded under these experimental conditions. Applying shorter test pulses (10 ms) partially reversed this problem, and Ca²⁺ transients looked more like those of the action potential-induced Ca²⁺ transients (not shown). However, analyzing the voltage-dependence of current and transient activation and a reliable comparison of current densities of channels with different activation kinetics necessitated long depolarization. Therefore, even though analysis of Ca²⁺ signals under voltage-clamp control is common practice and has contributed significantly to our current understanding of EC-coupling (e.g., Baylor et al., 1983; Brum et al., 1987; Melzer et al., 1984; Beurg et al., 1997; Dietze et al., 1998; Jurkat-Rott et al., 1998; Grabner et al., 1999; Wilkens et al., 2001), it has to be kept in mind that spatiotemporal Ca²⁺ gradients, which appear to be critical for activation of CICR under physiological conditions, probably break down during prolonged patch-clamp recordings.

A trivial alternative explanation for the different size, shape, and behavior of Ca²⁺ transients obtained with patch-clamp stimulation would be that the transients recorded from nonskeletal Ca²⁺ channels in the patch-clamp mode are not CICR at all, but simply reflect the Ca²⁺ influx through these channels. This would explain the reduced amplitude compared to that of the skeletal transients as well as the fact that GFP-_1A, which failed to restore action potential-induced Ca²⁺ transients, generates a Ca²⁺ signal under patch-clamp conditions. Attempts to test this possibility by pharmacologically isolating a possible Ca²⁺ influx signal from that of CICR did not resolve the issue either, because even concentrations between 10 µM and 40 µM ryanodine (Lipp et al., 2002) or 10 µM and 160 µM ruthenium red (Xu et al., 1999) in the patch pipette did not completely block Ca²⁺ release in control myotubes transfected with GFP-_1S. However, important evidence from this and previous studies (Tanabe et al., 1990; Garcia et al., 1994; Garcia and Beam, 1994) argues against the possibility that these Ca²⁺ transients arise exclusively from Ca²⁺ influx: first, the poor correlation of current densities with the occurrence of associated Ca²⁺ transients in individual myotubes; i.e., myotubes with current densities as small as 3 pA/pF showed transients whereas other myotubes with much larger currents showed no measurable transients (Fig. 5 D) (see also Garcia et al., 1994); second, the poor correlation of current densities with the amplitudes of Ca²⁺ transient for GFP-_1C and GFP-_1D, i.e., whereas their current densities differed by almost threefold, GFP-_1C and GFP-_1D produced Ca²⁺ transients of similar magnitude (Fig. 5, A and B); and finally, Ca²⁺ transients triggered by the cardiac-skeletal chimera GFP-CSk53 were not significantly larger than those of GFP-_1C, as would be expected from a construct that triggers skeletal-type Ca²⁺ release in addition to a fluo-4 signal reflecting pure Ca²⁺ influx. If we had merely recorded the influx of Ca