A Ca_V1.1 Ca²⁺ Channel Splice Variant with High Conductance and Voltage-Sensitivity Alters EC Coupling in Developing Skeletal Muscle

Abstract

The Ca²⁺ channel α_1S subunit (Ca_V1.1) is the voltage sensor in skeletal muscle excitation-contraction (EC) coupling. Upon membrane depolarization, this sensor rapidly triggers Ca²⁺ release from internal stores and conducts a slowly activating Ca²⁺ current. However, this Ca²⁺ current is not essential for skeletal muscle EC coupling. Here, we identified a Ca_V1.1 splice variant with greatly distinct current properties. The variant of the CACNA1S gene lacking exon 29 was expressed at low levels in differentiated human and mouse muscle, and up to 80% in myotubes. To test its biophysical properties, we deleted exon 29 in a green fluorescent protein (GFP)-tagged α_1S subunit and expressed it in dysgenic (α_1S-null) myotubes. GFP-α_1SΔ29 was correctly targeted into triads and supported skeletal muscle EC coupling. However, the Ca²⁺ currents through GFP-α_1SΔ29 showed a 30-mV left-shifted voltage dependence of activation and a substantially increased open probability, giving rise to an eightfold increased current density. This robust Ca²⁺ influx contributed substantially to the depolarization-induced Ca²⁺ transient that triggers contraction. Moreover, deletion of exon 29 accelerated current kinetics independent of the auxiliary α₂δ-1 subunit. Thus, characterizing the Ca_V1.1Δ29 splice variant revealed the structural bases underlying the specific gating properties of skeletal muscle Ca²⁺ channels, and it suggests the existence of a distinct mode of EC coupling in developing muscle.

Introduction

The voltage-gated Ca²⁺ channel Ca_V1.1 functions as a voltage sensor in skeletal muscle excitation-contraction (EC) coupling. It is located in triad junctions in close apposition to the Ca²⁺ release channel (type 1 ryanodine receptor (RyR1)) in the sarcoplasmic reticulum (SR). On depolarization of the surface membrane, Ca_V1.1 undergoes a conformational change that rapidly activates the Ca²⁺ release channel, presumably via protein-protein interactions. Ca²⁺ influx through the voltage-gated Ca²⁺ channel is not required for activation of skeletal muscle EC coupling. As a matter of fact, L-type Ca²⁺ currents through Ca_V1.1 activate very slowly and at more positive membrane potentials than EC coupling (for review, see Melzer et al. (1)). Therefore, it is unlikely that during a short skeletal muscle action potential Ca²⁺ channels contribute significant amounts of Ca²⁺ to the transients that trigger contraction.

Multiple splice variants greatly enrich the functional diversity of the CACNA1 gene family and the splicing patterns are conserved across different members of this gene family and across various species (2). Splicing has been extensively studied for the cardiac Ca_V1.2 encoded by CACNA1C (3–8) and for the neuronal Ca_V2.1 (CACNA1A) and Ca_V2.2 (CACNA1B) (9–11). In contrast, only one Ca_V1.1 splice variant has so far been described in rabbit skeletal muscle. Skipping of exon 29 (5) shortens the extracellular loop connecting transmembrane domains IVS3 and IVS4. This loop is a conserved splicing site of Ca_V1 α₁ subunits that has been shown to generate differentially distributed and functionally distinct channel variants. For example, the Ca_V1.3 and Ca_V2.2 IVS3–IVS4 splice variants are expressed in different regions of the cardiovascular system and the nervous system, respectively (11,12), and skipping of the corresponding exon in the smooth muscle variant of Ca_V1.2 resulted in a channel with increased voltage and drug sensitivity (13). For the skeletal muscle Ca_V1.1, the functional consequence of exon skipping in IVS3–IVS4 or the specific expression patterns of this splice variant have hitherto not been analyzed.

In this study, we describe Ca_V1.1 splice variants transcribed in human adult muscle and in muscle cell cultures. Because the variant lacking exon 29 (Ca_V1.1Δ29) was predicted to code for a functional channel and was abundantly expressed in myotubes, we performed a thorough analysis of its biophysical properties. Reconstitution of dysgenic (Ca_V1.1-null) myotubes with Ca_V1.1Δ29 fully restored skeletal muscle EC coupling, however, with Ca²⁺ currents eight times larger and activating at 30 mV less depolarizing potentials than Ca_V1.1 with exon 29. Thus, the analysis of the Ca_V1.1Δ29 splice variant identified the IVS3–IVS4 loop as the structural basis of the weak voltage sensitivity of the full-length skeletal muscle Ca²⁺ channels. Furthermore, our results indicate that in muscle cells expressing this newly characterized channel variant, Ca²⁺ influx contributes significantly to the cytoplasmic Ca²⁺ signal that triggers skeletal muscle contraction.

Materials and Methods

Detection of splice variants

Muscle biopsies (Vastus lateralis of five control subjects with their informed consent) were shredded and trypsinized for 30 min at 37°C. Satellite cells were isolated by filtration (nylon filter, 0.22 μm) and cultured in skeletal muscle growth medium (PromoCell, Heidelberg, Germany). After 10–14 days, the medium was changed to a differentiation medium (PromoCell). Mature myotubes (those with more than five nuclei) were obtained after an additional 1–2 weeks. Total RNA was extracted from homogenized mouse muscle tissues and cultured myotubes using TRIzol reagent (Gibco BRL, Gaithersburg, MD) according to the manufacturer's protocol. Reverse transcription polymerase chain reaction (RT-PCR) amplification was carried out with a one-step RT-PCR kit (Qiagen, Hilden, Germany) and overlapping primer pairs (see Table S3 in Supplementary Material). DNA fragments of splice variants were separated by 2% agarose gel electrophoresis and stained with ethidium bromide (0.7 mg/ml). Size and intensity of PCR bands were quantified using Scion Image 4.0.3.2 (Scion, Frederick, MD). DNA sequences of splice variants were confirmed sequencing (BIG Dye Terminator Cycle Sequencing Kits, Applied Biosystems, Foster City, CA).

Quantitative TaqMan PCR

Total RNA was isolated from differentiated myotubes from a skeletal muscle cell line derived from H2-KB-TSA58 mice (Immortomice (14,15)) using the RNeasy Mini kit (Qiagen) and reverse-transcribed (SuperScriptII, Invitrogen, Carlsbad, CA). The relative abundance of Ca_V1.1 and Ca_V1.1Δ29 mRNAs was assessed by TaqMan quantitative PCR (50 cycles) using a standard curve method based on PCR products of known concentrations (16). TaqMan gene expression assays, designed to span exon-exon boundaries (Table S3), were purchased from Applied Biosystems. cDNA concentrations of the three individual experiments were comparable, as revealed by the expression of seven different reference genes (β-actin, B2M, GAPDH, HPRT1, Tbp, Tfrc, and SDHA). Analyses were performed using the 7500 Fast System (Applied Biosystems).

Expression plasmids

Bases 3609–3666, corresponding to exon 29, were deleted from GFP-α_1S (17) by PCR splicing using a primer pair encompassing restriction sites upstream (XhoI) and downstream (BglII) of exon 29 (P1 and P2 (Table S3)), and two complementary primers each consisting of the end sequence of exon 28 joined to the start sequence of exon 30 (P3 and P4 (Table S3)). α₂δ-1 shRNA expression plasmid has been previously published (18).

Cell culture and transfections

Myotubes of the homozygous dysgenic (mdg/mdg) cell line GLT were cultured and transfected as previously described (19). Myotubes were analyzed 3–5 days after transfection.

Immunofluorescence and antibodies

Immunofluorescence analysis was performed as described (20), using the following antibodies: rabbit polyclonal anti-GFP (1:4000, Molecular Probes, Eugene, OR); mouse monoclonal anti-RyR (34-C, 1:1000, Alexis Biochememicals, Lausen, Switzerland); secondary goat-anti-mouse Alexa-594 and goat-anti-rabbit Alexa-488 (1:4000, Molecular Probes). Images were captured on a Zeiss Axiophot microscope with a cooled CCD camera and METAVUE image-processing software (Universal Imaging, West Chester, PA).

Electrophysiology and fluorescent Ca²⁺ measurements

Ca²⁺ currents were recorded with the ruptured whole-cell patch-clamp technique in voltage-clamp mode. The patch pipettes (borosilicate glass, Harvard Apparatus, Holliston, MA) had resistance of 1.5–3 MΩ when filled with (mM) 145 Cs-aspartate, 2 MgCl₂, 10 HEPES, 0.1 Cs-EGTA, 2 Mg-ATP, and 0.2 Fluo-4, with pentapotassium salt to record Ca²⁺ transients (pH 7.4 with CsOH). The extracellular bath solution contained (mM) 10 CaCl₂, 145 tetraethylammoniumchloride, 10 HEPES (pH 7.4 with tetra-ethylammoniumhydroxide). Recording of gating charge movement was performed in the presence of 0.5 mM Cd²⁺ and 0.2 mM La³⁺ (Cd²⁺/La³⁺) to block the inward Ca²⁺ currents. All recordings were made with an Axopatch 200A amplifier (Axon Instruments, Foster City, CA). Data acquisition and command potentials were controlled by pClamp software (version 8.0, Axon Instruments).

The current-voltage dependence was fitted according to

$$I=G_{\max }\times \left(V-V_{\text{rev}}\right)/\left(1+\exp \left(-\left(V-V_{1/2}\right)/k\right)\right),$$ (1)

where G_max is the maximum conductance of the L-type Ca²⁺ channels, V_rev is the extrapolated reversal potential of the Ca²⁺ current, V_1/2 is the potential for half-maximal conductance, and k is the slope.

The voltage dependence of the Ca²⁺ conductance, “On” gating charge movement, and Ca²⁺ transients were fitted according to a Boltzmann distribution:

$$G=G_{\max }/\left(1+\exp \left(-\left(V-V_{1/2}\right)/k\right)\right)\text{.}$$ (2)

The kinetic properties of the Ca²⁺ current activation were determined by fitting the rising phase of the maximum sweep (+40 or +50 mV for GFP-α_1S and +10 or +20 mV for GFP-α_1SΔ29 and GFP-α_1SΔ29 + α₂δ-1 siRNA) with a single- or double-exponential function:

$$I=A_{\text{fast}}\times \left(\exp \left(-t/{\tau }_{\text{fast}}\right)\right)+A_{\text{slow}}\times \left(\exp \left(-t/{\tau }_{\text{slow}}\right)\right)+C\text{or}$$

$$I=A_{\text{mono}}\times \left(\exp \left(-t/{\tau }_{\text{mono}}\right)\right)+C,$$ (3)

where I is the current; A_fast, A_slow, and A_mono are the individual current amplitudes; and τ_fast, τ_slow, and τ_mono are the specific time constants of the current amplitudes.

Results

Human and mouse skeletal muscles express transcripts of multiple Ca_V1.1 splice variants

To determine the genetic variability of the skeletal muscle voltage-gated Ca²⁺ channel, we performed an RT-PCR screen for Ca_V1.1 splice variants on RNA from human and mouse muscle and cultured myotubes. We identified nine rare variants coding for nonsense proteins, each of which made up <3% of transcripts in myotubes and muscle (Table S1 and Fig. S1). Eight of these produced frame shifts resulting in premature stops, precluding formation of functional channels. The ninth, a deletion of exon 19, lacks almost an entire transmembrane segment and would thus invert the transmembrane orientation of the channel distal to IIIS1. Most likely, all of these transcripts are subject to nonsense-mediated decay, a process that affects transcripts with premature termination codons (21).

Further, we identified three C-terminal Ca_V1.1 variants, all of which code for truncated but putatively functional proteins (Table S2). Deletion of exons 39–40 results in a premature stop after only three residues. Alternative 5′ splicing of exon 40 results in the in-frame deletion of 26 residues in positions 1599–1625. The insertion of intron 43 generates a protein with 61 residues at the C-terminus. Because the distal end of the channel has previously been shown to exert an autoinhibitory effect on gating charge movements and Ca²⁺ currents (22), all three C-terminal variants are expected to result in a gain of function similar to an artificial truncation at position 1698. Transcripts of all three C-terminal variants were expressed in low abundance.

Finally, the RT-PCR screen revealed an in-frame deletion of exon 29 (Δ29) (Fig. 1 A) that encodes a 1854-amino-acid protein lacking 19 residues in the extracellular loop between transmembrane segments IVS3 and IVS4 (Fig. 1 B). This Ca_V1.1 variant had previously been identified in mouse ovaries, in the BC3H1 cell line (5), and in rat osteosarcoma cells (23). Quantifying the RT-PCR bands of the full-length and splice-variant Ca²⁺ channels showed that Ca_V1.1Δ29 was expressed at low levels in differentiated muscles from man and mouse (2–10% of transcripts), but was abundant in primary human myotubes and in the C₂C₁₂ mouse muscle cell line (60–80% of transcripts (Fig. 1 C)). Taqman RT-PCR, using assays spanning the borders between exons 28–29 and exons 28–30 (Table S3), confirmed the high expression levels of Ca_V1.1Δ29 in an additional skeletal muscle cell line. Of the Ca_V1 transcripts in RNA from myotubes derived from the Immortomouse, 77 ± 3% lacked exon 29 (Fig. 1 D). Together, these data show that Ca_V1.1Δ29 is the predominant Ca_V1.1 isoform in skeletal myotubes. Analogous splice variants in other L-type Ca²⁺ channel α₁ subunits code for functional channels. Thus, it is likely also that Ca_V1.1Δ29 is functional and thus may be of physiological relevance.

Figure 1.

Detection of GFP-α_1SΔ29 in human and mouse myotubes. (A) Sequence of the boundary between exons 28 and 30. (B) Location of exon 29 in a domain model of Ca_V1.1. (C) Full-length (upper band) and Ca_V1.1Δ29 (lower band) detected by RT-PCR amplification of exons 26–30 in RNA prepared from mouse muscle, C₂C₁₂ myotubes, human muscle, and human primary myotubes. (D) The fraction of Ca_V1.1 transcripts with and without exon 29 measured with quantitative RT-PCR in mRNA from mouse myotubes. Error bars represent the mean ± SE, p << 0.001.

Ca_V1.1Δ29 is localized in the EC-coupling apparatus

To analyze the physiological properties of Ca_V1.1Δ29, we deleted the sequence corresponding to amino acids 1203–1222 in the GFP-tagged Ca_V1.1 α_1S subunit (17) and expressed the resulting construct, GFP-α_1SΔ29, in dysgenic (α_1S-null) myotubes. Reconstitution of dysgenic myotubes with GFP-α_1S restores normal triad targeting of the GFP-α_1S subunit (Fig. 2, left column) as well as skeletal muscle EC coupling (20). Double-immunofluorescence analysis of GFP-α_1SΔ29-transfected myotubes showed that, like the wild-type GFP-α_1S, GFP-α_1SΔ29 was localized in a clustered pattern that was colocalized with clusters of RyR1 (Fig. 2, right column). This labeling pattern is typical for triad proteins and indicates that GFP-α_1SΔ29 is correctly targeted into junctions between the SR and t-tubules or the plasma membrane, and thus can function in EC coupling (20).

Figure 2.

GFP-α_1SΔ29 is targeted into junctions between the SR and t-tubules or the plasma membrane. Dysgenic myotubes expressing full-length GFP-α_1S (left) or the GFP-α_1SΔ29 splice variant (right) were double-immunolabeled with anti-RyR (upper) and anti-GFP (middle). Clusters of GFP-α_1S and GFP-α_1SΔ29 colocalized with the RyR1 (lower, yellow clusters in color overlay) indicate the correct targeting of both Ca²⁺ channel variants into t-tubule/SR or plasma membrane/SR junctions. Scale bar, 10 μm.

Ca_V1.1Δ29 has distinct current properties

Whole-cell patch-clamp recordings in dysgenic myotubes reconstituted with GFP-α_1SΔ29 revealed greatly augmented Ca²⁺ currents (Fig. 3 A). Mean peak current density was increased eightfold from 1.9 ± 0.2 pA/pF in GFP-α_1S-expressing myotubes to 14.8 ± 1.2 pA/pF in myotubes expressing GFP-α_1SΔ29 (Fig. 3 B and Table 1). Moreover, the current-voltage curves showed a greatly increased voltage sensitivity. The half-maximal activation was shifted from 39.1 ± 1.3 mV for the wild-type to 9.3 ± 1.0 mV for the GFP-α_1SΔ29 channel (Fig. 3 C). Because at less positive voltages the driving force for Ca²⁺ currents is larger, the left-shifted voltage dependence of activation accounted in part for the observed increase in current density.

Figure 3.

GFP-α_1SΔ29 has increased current density, voltage sensitivity of activation, and open probability (P_o). (A) Representative whole-cell currents from myotubes expressing either GFP-α_1S or GFP-α_1SΔ29. (B) I/V curves show that the peak current density of GFP-α_1SΔ29 is increased. (C) Voltage dependence of activation is shifted toward more negative potentials for GFP-α_1SΔ29 compared to full-length GFP-α_1S. (D and E) Analysis of the “On” gating charges (Q_on) while currents are blocked with Cd²⁺/ La³⁺ shows that deletion of exon 29 did not alter the expression of functional channels in the membrane. (F and G) The amplitudes of the tail currents, recorded at the reversal potential were plotted against Q_on. The increased slope of the linear regression indicates that the channel P_o is considerably increased in GFP-α_1SΔ29 compared to GFP-α_1S. Error bars indicate the SE.

Table 1.

Properties of Ca²⁺ currents from α_1S and α_1S-ΔE29

	Parameters	α1S	α1S-ΔE29	Significance (p-value)
Current properties	Ipeak (pA/pF)	−1.9 ± 0.2	−14.8 ± 1.2	<<0.001
	Gmax (nS/nF)	79.1 ± 8.4	255.1 ± 17.8	<<0.001
	V1/2 (mv)	39.1 ± 1.3	9.3 ± 1.0	<<0.001
	kact (mv)	7.2 ± 0.4	4.8 ± 0.3	<<0.001
	Vrev (mv)	82.7 ± 2.1	83.1 ± 0.8	0.81
	n	26	33	—
Kinetics	Time to peak (ms)	84.7 ± 6.3	39.8 ± 4.5	<<0.001
	% Inactivation	4.7 ± 1.2	23.8 ± 2.0	<<0.001
	n	26	33	—
	Aslow contribution	62%	65%	0.24
	Afast contribution	38%	35%	0.24
	Aslow (pA/pF)	1.4 ± 0.3	7.9 ± 1.3	<<0.001
	Afast (pA/pF)	0.9 ± 0.1	3.9 ± 0.9	0.001
	τslow (ms)	28.7 ± 3.3	14.1 ± 2.5	0.003
	τfast (ms)	5 ± 0.5	3.9 ± 0.6	0.15
	n	21	18	—
	Amono (pA/pF)	—	20.1 ± 3.2	—
	τmono (ms)	—	4.3 ± 0.2	—
	n	—	15	—
Charge movement	QON(nC/μF)	4.1 ± 0.9	3.9 ± 0.5	0.90
	Slope	19.8 ± 2.2	15.5 ± 0.9	0.07
	V1/2 (mv)	14.3 ± 5.6	7.8 ± 2.6	0.28
	n	8	10	—
QON vs. ITail	ITail (pA/pF)	−5.7 ± 0.8	−40.3 ± 7.3	<0.001
	Slope	0.38 ± 0.07	3.59 ± 0.89	—
	n	10	12

All data are presented as mean ± SE.

To determine whether enhanced membrane expression of Ca²⁺ channels also contributed to the increase in current density, we analyzed the charge movements upon channel activation in the presence of Cd²⁺/La³⁺ to block the Ca²⁺ conductance (Fig. 3 D). The integral of the immobilization-resistant “On” gating charge movement (Q_on) is a measure of functional Ca²⁺ channels in the membrane (24). In myotubes expressing GFP-α_1S or GFP-α_1SΔ29, Q_on was not significantly different at any voltage (Fig. 3 E), indicating that membrane expression of functional Ca²⁺ channels was not changed in the variant lacking exon 29. Alternatively, altered single-channel conductance could be responsible for the increased current density. Therefore, we analyzed the tail currents at the end of 200-ms test pulses to near the reversal potential, where a maximal number of channels should be activated (Fig. 3 F), and we plotted the peak current density of the tail currents against the integral of Q_on (25). The slope of the linear regression was increased from 0.38 ± 0.07 (GFP-α_1S) to 3.59 ± 0.89 (GFP-α_1SΔ29) (Fig. 3 G). This indicated that deletion of exon 29 caused a significant increase in the relative open probability (P_o) (Table 1). Together, electrophysiological analyses demonstrated that compared to the full-length isoform, the Ca_V1.1Δ29 splice variant has a greatly increased voltage sensitivity and open probability, resulting in an eightfold increase of the whole-cell Ca²⁺ currents.

Ca_V1.1Δ29 has accelerated current kinetics

Another property of the Ca²⁺ current that was altered by the deletion of exon 29 was the speed of activation and inactivation. This is evident in the representative current traces shown in Fig. 3 A. The average time to peak measured in the maximum current traces was significantly reduced from 84.7 ± 6.3 ms (GFP-α_1S) to 39.8 ± 4.5 ms (GFP-α_1SΔ29) (Fig. 4 A). In a similar way, the fractional inactivation at the end of the 200-ms test pulse increased from 4.7 ± 1.2% to 23.82 ± 2.0% (Fig. 4 B).

Figure 4.

Deletion of exon 29 accelerates Ca²⁺ current kinetics. (A and B) Currents recorded from myotubes expressing GFP-α_1SΔ29 exhibit a significantly shorter time to peak (A) and an increased fractional inactivation during a 200-ms pulse (B). The rising phase of Ca²⁺ currents was fitted by a double-exponential function and the amplitudes and time constants of the two components were calculated. Neither the ratio between fast and slow components (C) nor the time constant of the fast component (D, solid bars) was affected by the deletion of exon 29. The time constant of the slow component was significantly faster in GFP-α_1SΔ29 compared to GFP-α_1S (D, hatched bars). Error bars represent the mean ± SE.

To elucidate the mechanism of Ca²⁺-current acceleration we further analyzed the activation phase by fitting the maximum current traces. Kinetic analysis of GFP-α_1SΔ29-expressing myotubes using a double-exponential function (26) showed that the majority of the currents consisted of a fast- and a slow-activating component. Whereas the relative abundance of both components (Fig. 4 C), and the time constant of the fast component, were equal for GFP-α_1S and GFP-α_1SΔ29, the time constant of the slow component was significantly faster (14.1 ± 2.5 ms) compared to that of GFP-α_1S (28.7 ± 3.3 ms) (n = 18) (Fig. 4 D and Table 1). In addition, activation kinetics of 15 out of 33 currents were best described by a single-exponential function with a time constant equal to that of τ_fast in the currents with two activation components (Table 1). Whether this resulted from an even greater acceleration of the slow component or from an almost complete loss of its contribution to the whole-cell current cannot be discerned.

Ca_V1.1Δ29 interacts with the α₂δ-1 subunit

Previously, we reported that depletion of the Ca²⁺ channel α₂δ-1 subunit with siRNA also accelerated Ca²⁺ current kinetics (18). Therefore, we examined whether GFP-α_1SΔ29 currents were affected by depletion of α₂δ-1. Fig. 5 A shows that coexpressing dysgenic myotubes with GFP-α_1SΔ29 and a plasmid coding for α₂δ-1 shRNA further accelerated activation and inactivation of Ca²⁺ currents compared to those expressing GFP-α_1SΔ29 alone. Knock-down of α₂δ-1 showed a 42% decrease in time to peak to 22.9 ± 0.1 ms (Fig. 5 B and Table 2), and a 22% increase in the fractional inactivation to 30.6 ± 0.2% (Fig. 5 C). Moreover, as previously shown for GFP-α_1S, it is also the case with GFP-α_1SΔ29 that the rising phase of all currents in α₂δ-1-depleted myotubes was best fitted by a single-exponential function (Fig. 5 D). This indicates that GFP-α_1SΔ29 still interacted with the α₂δ-1 subunit in skeletal myotubes. Furthermore, immunocytochemical analysis demonstrated that the deletion of exon 29 did not affect triad targeting of the α₂δ-1 subunit (Fig. S2). Thus, the mechanisms by which the IVS3–IVS4 loop and the α₂δ-1 subunit regulate activation kinetics are independent of each other.

Figure 5.

Deletion of exon 29 does not affect the interaction of α₁ with the α₂δ-1 subunit. The α₂δ-1 subunit was depleted with shRNA in myotubes expressing GFP-α_1SΔ29. This resulted in a further acceleration of activation and inactivation kinetics as seen in the sample recordings (A). The time to peak was further reduced (B), and the percentage of inactivation was increased (C). Kinetic analysis of the activation phase revealed only one component of activation (D, white bar) with a time constant equal to τ_fast in GFP-α_1SΔ29 controls (E, gray bars), indicating the loss of the slow-activating component. Error bars represent the mean ± SE.

Table 2.

Properties of Ca²⁺ currents from α_1S-ΔE29 and α_1S-ΔE29 + α₂δ-1 siRNA

	Parameters	α1S-ΔE29	α1S-ΔE29 + α2δ-1 siRNA	Significance (p-value)
Current properties	Ipeak (pA/pF)	−14.8 ± 1.2	−14.3 ± 2.1	0.84
	Gmax (nS/nF)	255.1 ± 17.8	249.5 ± 28.8	0.89
	V1/2 (mv)	9.3 ± 1.0	8.9 ± 1.3	0.88
	kact (mv)	4.8 ± 0.3	4.5 ± 0.3	0.70
	Vrev (mv)	83.1 ± 0.8	79.7 ± 1.7	0.05
	n	33	6	—
Kinetics	Time to peak (ms)	39.8 ± 4.5	22.87 ± 2.1	0.12
	% Inactivation	23.8 ± 2.0	30.6 ± 2.0	0.17
	n	33	6	—
	Aslow contribution	65%	0%	—
	Afast contribution	35%	100%	—
	Aslow (pA/pF)	7.9 ± 1.3	—	—
	Afast (pA/pF)	3.9 ± 0.9	—	—
	τslow (ms)	14.1 ± 2.5	—	—
	τfast (ms)	3.9 ± 0.6	—	—
	n	18	6
	Amono (pA/pF)	20.1 ± 3.2	14.1 ± 2.38	0.28
	τmono (ms)	4.3 ± 0.2	5.3 ± 0.42	0.07
	n	15	6	—

All data are presented as mean ± SE.

Ca_V1.1Δ29 supports skeletal muscle EC coupling with a substantial component of Ca²⁺ influx

Ca_V1.1 Ca²⁺ currents activate slowly and at much higher membrane potentials than depolarization-induced Ca²⁺ release from the SR. As a consequence, Ca²⁺ influx through the Ca_V1.1 Ca²⁺ channel does not essentially contribute to skeletal muscle EC coupling (1). However, the current properties of GFP-α_1SΔ29 described above resemble much more those of the cardiac Ca_V1.2, which activates EC coupling by Ca²⁺-induced Ca²⁺ release (27). Therefore, it was important to determine whether GFP-α_1SΔ29 can trigger EC coupling in skeletal myotubes, and if so, by which mechanism. To this end, we measured Ca²⁺ transients during patch-clamp experiments using the fluorescent Ca²⁺ indicator Fluo-4. Fig. 6 A shows that during a depolarizing pulse, GFP-α_1SΔ29-expressing myotubes give rise to Ca²⁺ transients, indicating that GFP-α_1SΔ29 can activate EC coupling. Because of its independence of Ca²⁺ influx, skeletal muscle EC coupling remains fully activated at test pulses to near the reversal potential. Therefore, in myotubes expressing GFP-α_1S, the voltage-dependence curve of Ca²⁺ transients has the characteristic sigmoidal shape (Fig. 6 B, squares). The voltage-dependence curve of transients in GFP-α_1SΔ29 expressing myotubes peaks earlier and at a higher level than that of GFP-α_1S before it declines to the same level as that of GFP-α_1S (Fig. 6 B (gray stars) and Table 3). The fact that at the reversal potential GFP-α_1SΔ29 still activates solid Ca²⁺ transients demonstrates that it supports the skeletal muscle EC coupling mechanism. The early-activating additional component is likely caused by Ca²⁺ entering the myotubes from the outside through the Ca_V1.1Δ29 channel. Indeed, blocking Ca²⁺ currents with Cd²⁺/La³⁺ abolished the early peak of the voltage-dependence curve so that it exactly matched that of the full-length GFP-α_1S isoform (Fig. 6 B, open stars).

Figure 6.

GFP-α_1SΔ29 supports skeletal muscle type EC coupling with an additional component of Ca²⁺ influx. Depolarization-induced Ca²⁺ transients were recorded in dysgenic myotubes reconstituted with GFP-α_1S (A, upper) or GFP-α_1SΔ29 (A, lower). The voltage dependence of activation was not altered by the deletion of exon 29, but Ca²⁺ transients were augmented by a component that declined at voltages near the reversal potential and could be inhibited by blocking Ca²⁺ currents with Cd²⁺/La³⁺ (B). Error bars represent the mean ± SE.

Table 3.

Properties of Ca²⁺ transients

	Parameters	α1S-WT	α1S-ΔE29	Significance (p-value)
Transients before Cd2+/La3+	ΔF/F	0.63 ± 0.07	0.81 ± 0.10	0.44
	V1/2 (mv)	4.8 ± 1.1	1.0 ± 1.0	0.02
	kact (mv)	7.1 ± 0.16	6.2 ± 0.25	0.005
	n	15	16	—
Transients after Cd2+/La3+	ΔF/F	0.64 ±0.06	0.55 ± 0.08	0.33
	V1/2 (mv)	6.8 ± 1.4	8.1 ± 1.7	0.55
	kact (mv)	8.4 ± 0.3	9.4 ± 0.9	0.27
	n	15	13	—

All data are presented as mean ± SE.

Discussion

Here, we report the first functional characterization of a splice variant of the skeletal muscle voltage-gated Ca²⁺ channel, and the extent to which the channel properties of Ca_V1.1Δ29 differ from those of the classical full-length Ca_V1.1 is amazing. Ca_V1.1Δ29 is normally targeted into the triads and supports skeletal muscle type EC coupling, but the lack of exon 29 causes drastically increased voltage sensitivity and open probability of the channel. Alternative splicing of the extracellular loop between transmembrane segments 3 and 4 in the fourth homologous repeat has also been described in other L-type and non-L-type Ca²⁺ channels (5,7,9–11,13,23). It is interesting to note that in all these cases, changes in the length of the IVS3–IVS4 linker were accompanied by changes in the voltage dependence of activation. In Ca_V1.2, 12 possible splicing combinations exist in this domain, four of which have been functionally characterized. The IVS3–IVS4 loop of variant A was shortened by 13 amino acids and the potential of half-maximal activation (V_1/2) was left-shifted by −9.5 mV; that of variant B was 6 residues shorter, with V_1/2 shifted by −4.9 mV; and that of variant C was 11 residues shorter, with V_1/2 shifted by −6.8 mV. In variant D, a 12-amino-acid sequence was exchanged without changing the total length of the IVS3–IVS4 loop, and the voltage sensitivity remained unaltered (7). Here, we demonstrate that the lack of exon 29 in Ca_V1.1Δ29, which shortens the IVS3–IVS4 loop by 19 amino acids, leads to a −29.8 mV shift of the voltage dependence of activation. Together, these observations suggest an inverse correlation of the length of the IVS3–IVS4 loop and the voltage sensitivity of Ca²⁺-channel splice variants. From a mechanistic perspective, it is conceivable that a shorter IVS3–IVS4 loop pulls the positively charged voltage sensor IVS4 toward the extracellular side of the membrane and thus facilitates its transition into the activated state upon depolarization. This would result in an increased voltage sensitivity and an increased dwell time in the open state.

The distinguishing characteristics of the skeletal muscle Ca²⁺ current are its extremely slow activation kinetics and activation at very positive potentials. Using skeletal/cardiac muscle Ca_V1 chimeras, a sequence including transmembrane domain S3 and the S3–S4 loop of the first repeat was shown to determine the slow gating mode (28). The results of this study demonstrate the importance of a separate domain, the IVS3–IVS4 loop, for the characteristic voltage dependence of activation of Ca_V1.1. Indeed, Nakai et al. (28) reported that in their chimeras, the slow and fast kinetics did not correlate with low and high voltage sensitivity, respectively. Our study provides independent evidence that voltage sensitivity and activation kinetics are determined by separate mechanisms. We have shown previously that the auxiliary α₂δ-1 subunit is an important determinant of the slow activation of Ca_V1.1, and that shRNA depletion of α₂δ-1 accelerated current activation by increasing the population of fast-activating channels at the expense of slow-activating channels (18). The voltage sensitivity of activation was not altered by α₂δ-1 siRNA treatment. Although the removal of exon 29 also accelerated activation kinetics, this resulted from a reduced time constant of the slow-activating component. Moreover, Ca_V1.1Δ29 channels were still sensitive to depletion of α₂δ-1, which caused an additional increase in activation kinetics. Thus, two separate sequences in the corresponding domains of the first and fourth repeats of Ca_V1.1 are responsible for the characteristic kinetics and voltage dependence of activation, respectively. Whereas IS3 and the IS3–IS4 loop appear to cooperate with the α₂δ-1 subunit in determining the activation kinetics, the length of the IVS3–IVS4 loop determines the voltage sensitivity of Ca_V1.1.

It is interesting that the deletion of exon 29 only affected the voltage sensitivity of the Ca²⁺ current, but not that of the “On” gating charges and Ca²⁺ transients, which activate at similarly low potentials in Ca_V1.1 and Ca_V1.1Δ29. Thus, the long IVS3–IVS4 loop, and with it the voltage sensor of repeat 4, is rate-limiting for activation of the Ca²⁺ current but not for activation of SR Ca²⁺ release. Removing exon 29 was sufficient to render the voltage sensor in the fourth repeat as sensitive to depolarization as those responsible for EC coupling. Based on elegant electrophysiological experiments in frog muscle fibers, Feldmeyer et al. (29) (reviewed in Melzer et al. (1)) proposed a model according to which Ca_V1.1 possesses three fast- and one slow-activating voltage sensors, which are responsible for the activation of EC coupling and Ca²⁺ currents, respectively. A similar model would readily explain the differential voltage sensitivity of EC coupling and the Ca²⁺ currents. If the rapid gating of two or three voltage sensors is sufficient for activating SR Ca²⁺ release, but the gating of all four voltage sensors is necessary for activation of the Ca²⁺ current, a single less responsive voltage sensor would be enough to delay current activation. However, since the voltage sensors of repeats I and IV appear to be involved in controlling the kinetics and voltage sensitivity of the current, respectively, the two other voltage sensors, those of repeats II and III, which flank the cytoplasmic loop that interacts with the RyR1, may be sufficient for the rapid activation of EC coupling. If, however, the short IVS3–IVS4 loop would hold the voltage sensor of repeat IV in a constitutively activated position, a reduction of Q_on by one-fourth would have been expected in Ca_V1.1Δ29. This was not observed. Therefore, the shorter IVS3–IVS4 loop appears to enhance the coupling between the voltage-dependent step and the final voltage-independent step of channel gating.

We further demonstrate that transcripts of the Ca_V1.1 splice variant lacking exon 29 (Ca_V1.1Δ29) are expressed in human and mouse muscles and that Ca_V1.1Δ29 is the predominant isoform in human and mouse cultured myotubes. If this is so, Ca²⁺ currents in normal myotubes like the C₂C₁₂ cell line, which express a mix of Ca_V1.1 and Ca_V1.1Δ29, should have an intermediate voltage sensitivity compared to dysgenic myotubes expressing either GFP-α_1S or GFP-α_1SΔ29. Indeed, this was observed in two previous studies by Schuhmeier et al. (30,31), who determined the voltage sensitivity of Ca²⁺ currents and Ca²⁺ transients in reconstituted dysgenic myotubes and in C₂C₁₂ myotubes. Consistent with the data presented here, in dysgenic myotubes transfected with GFP-α_1S, half-maximal activation of Ca²⁺ currents occurred at ∼30 mV higher potentials than activation of Ca²⁺ transients. In contrast, this difference was only ∼15 mV in C₂C₁₂ cells. Thus, the current properties of cultured myotubes most likely reflect the properties of a mixed population of Ca_V1.1 splice variants.

Finally, Ca_V1.1Δ29 was correctly incorporated into triad junctions and supported skeletal muscle type EC coupling, although the highly increased voltage sensitivity and amplitude of Ca²⁺ currents altered the EC coupling properties. Dysgenic myotubes exclusively expressing the GFP-α_1SΔ29 isoform showed depolarization-dependent Ca²⁺ transients, which did not require Ca²⁺ influx through the channel. In addition, the rapidly activating Ca²⁺ current contributed significant amounts of Ca²⁺ to the cytoplasmic Ca²⁺ transients. Under physiological conditions, this component will be diminished by the presence of the full-length Ca_V1.1. Nevertheless, the additional Ca²⁺ influx is expected to increase the force of contraction and it creates the need for increased export of Ca²⁺ from the cell to maintain equilibrium. The observation that Ca_V1.1Δ29 transcripts were highly expressed in cultured myotubes from humans and mice suggests a physiological role of this splice variant in developing and regenerating muscle. It is plausible that myotubes with an incompletely differentiated SR Ca²⁺ storage and release apparatus require more Ca²⁺ influx and rely part on a cardiac-like EC coupling mechanism. Moreover, given the great variety of muscle types in our body it is intriguing to consider the possibility that Ca_V1.1Δ29 may also play a role in some type of differentiated muscle. Finally, Ca_V1.1 mutants affecting Ca²⁺ current properties have been linked to human disease, but the effects of these disease mutants on EC coupling were small (32,33). In light of the existence of a splice variant with dramatically different current properties and its possible function during development, the effects of these disease mutations on this splice variant, as well as the possible pathological consequences, need to be investigated.

Supplementary Material

Three tables and two figures are available at www.biophys.org/biophysj/supplemental/S0006-3495(08)00036-2.