Ca2+ Release through Ryanodine Receptors Regulates Skeletal Muscle L-type Ca2+ Channel Expression*

Guillermo AvilaDagger, Kristen M. S. O'Connell, Linda A. Groom, and Robert T. Dirksen§

From the Department of Pharmacology and Physiology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642

Received for publication, October 23, 2000


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Skeletal muscle obtained from mice that lack the type 1 ryanodine receptor (RyR-1), termed dyspedic mice, exhibit a 2-fold reduction in the number of dihydropyridine binding sites (DHPRs) compared with skeletal muscle obtained from wild-type mice (Buck, E. D., Nguyen, H. T., Pessah, I. N., and Allen, P. D. (1997) J. Biol. Chem. 272, 7360-7367 and Fleig, A., Takeshima, H., and Penner, R. (1996) J. Physiol. (Lond.) 496, 339-345). To probe the role of RyR-1 in influencing L-type Ca2+ channel (L-channel) expression, we have monitored functional L-channel expression in the sarcolemma using the whole-cell patch clamp technique in normal, dyspedic, and RyR-1-expressing dyspedic myotubes. Our results indicate that dyspedic myotubes exhibit a 45% reduction in maximum immobilization-resistant charge movement (Qmax) and a 90% reduction in peak Ca2+ current density. Calcium current density was significantly increased in dyspedic myotubes 3 days after injection of cDNA encoding either wild-type RyR-1 or E4032A, a mutant RyR-1 that is unable to restore robust voltage-activated release of Ca2+ from the sarcoplasmic reticulum (SR) following expression in dyspedic myotubes (O'Brien, J. J., Allen, P. D., Beam, K., and Chen, S. R. W. (1999) Biophys. J. 76, A302 (abstr.)). The increase in L-current density 3 days after expression of either RyR-1 or E4032A occurred in the absence of a change in Qmax. However, Qmax was increased 85% 6 days after injection of dyspedic myotubes with cDNA encoding the wild-type RyR-1 but not E4032A. Because normal and dyspedic myotubes exhibited a similar density of T-type Ca2+ current (T-current), the presence of RyR-1 does not appear to cause a general overall increase in protein synthesis. Thus, long-term expression of L-channels in skeletal myotubes is promoted by Ca2+ released through RyRs occurring either spontaneously or during excitation-contraction coupling.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The skeletal muscle dihydropyridine receptor (DHPR)1 functions both as a slowly activating L-type Ca2+ channel (L-channel) and as a voltage sensor that controls the activity of the type 1 ryanodine receptor (RyR-1) present in the sarcoplasmic reticulum (SR). During excitation-contraction (EC) coupling, sarcolemmal depolarization (e.g. an action potential) induces voltage-driven conformational changes in the DHPR, which can be measured electrophysiologically as nonlinear capacitative currents, termed intramembrane charge movements or gating currents. These charge movements are thought to mechanically activate RyR-1 proteins during EC coupling and thus lead to a massive release of SR calcium (orthograde signal of EC coupling; see Ref. 1 for review).

Analysis of skeletal myotubes derived from RyR-1-knockout (dyspedic) mice has revealed that in addition to the orthograde signal of EC coupling (signal transmitted from the DHPR to the RyR-1), there is also a retrograde signal whereby RyR-1 promotes the calcium conducting activity of the skeletal L-channel (2, 3). This conclusion was inferred from the observation that despite a significant surface density of DHPRs, dyspedic myotubes exhibit a marked (~90%) reduction in L-current. Moreover, short-term (2-4 days) expression of RyR-1 in dyspedic myotubes considerably enhances L-current density in the absence of a change in intramembrane charge movement (2, 3). These observations indicate that RyR-1 promotes the L-channel activity of the skeletal muscle DHPR in a manner that is independent of L-channel expression (retrograde signal of EC coupling).

Dyspedic muscle exhibits a 25-50% reduction in total DHP binding (4, 5). Accordingly, we have reported that dyspedic myotubes possess a significant reduction in maximal intramembrane charge movement compared with normal myotubes (3). This apparent reduction in the number of functional DHPRs in the sarcolemma cannot completely account for the ~90% decrease in L-current density found in dyspedic myotubes. In fact, dyspedic myotubes exhibit a nearly 5-fold reduction in the current-to-charge and conductance-to-charge (Gmax/Qmax) ratios, compared with both normal and RyR-1-expressing dyspedic myotubes (3). These observations support the conclusion of Nakai et al. (2) that RyR-1 promotes the Ca2+ conducting activity of the skeletal muscle L-channel. However, the mechanism(s) underlying the different DHPR expression levels in normal and dyspedic muscle have yet to be investigated. Considering the functional effects of reintroduction of RyR-1 in dyspedic myotubes (i.e. restoration of both retrograde and orthograde signals of EC coupling), either Ca2+ influx (via L-channels) and/or SR Ca2+ release (via RyR-1) could play a critical role in the regulation of sarcolemmal DHPR expression.

Despite the central role that the DHPR plays in skeletal EC coupling, controversy exists with regard to the mechanisms that control the expression of L-channels and voltage sensors. In frog skeletal muscle, long-term blockade of skeletal L-channels increases the number of sarcolemmal L-channels and voltage sensors, suggesting that Ca2+ influx (via L-channels) inhibits DHPR expression (6). However, elevations in extracellular Ca2+ increase L-current density and intramembrane charge movement in cultured rat myoballs (7). In the present study, we have investigated the roles of Ca2+ influx and release on the regulation of DHPR expression (L-channel activity and charge movement), following expression in dyspedic myotubes of either wild-type RyR-1 or a mutant RyR-1 (E4032A) that preferentially restores the retrograde (i.e. L-current) signal of skeletal muscle EC coupling (8). Using electrophysiological criteria (Ca2+ current and charge movement magnitudes) as a measure of functional DHPR activity in the sarcolemma, our results demonstrate that functional DHPR expression is promoted by Ca2+ released through RyR-1 proteins and not via Ca2+ influx through sarcolemmal L-channels.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Preparation of Myotubes-- Myotubes were prepared from primary culture of normal and dyspedic muscle as described previously (2). Expression of wild-type RyR-1 and the E4032A point mutation in RyR-1 in dyspedic myotubes was achieved by nuclear microinjection (9) of the appropriate expression plasmid (0.5 µg/µl) 6-8 days after initial plating of myoblasts. The E4032A mutation in RyR-1 was constructed using a standard two-step site-directed mutagenesis strategy (10). The entire polymerase chain reaction-modified cDNA portion was ultimately confirmed by sequence analysis. Expressing myotubes were examined electrophysiologically 2-6 days following nuclear microinjection. In some experiments (Figs. 1, 2, and 5), expressing myotubes were identified by the development of green fluorescence 2-6 days after coinjection with a mixture of either RyR-1 or E4032A cDNAs (0.5 µg/µl) and a cDNA expression plasmid encoding an enhanced green fluorescence protein (0.1 µg/µl). Coinjection of GFP cDNA was omitted when cells were to be used for measurements of intracellular Ca2+ transients (Fig. 4). For these experiments, expression was established by the presence of either electrically evoked (8.0 V, 10-30 ms) contractile activity (RyR-1) or large, slowly activating L-type Ca2+ currents (E4032A).

Measurements of Ionic and Gating Currents-- The whole-cell variant of the patch clamp technique (11) was used to measure ionic and gating currents in both normal and dyspedic myotubes, as described previously (3). Inward L-currents were elicited by 200 ms test pulses of variable amplitude from a holding potential (HP) of -80 mV, following a prepulse protocol (12, 10) used to inactivate T-type Ca2+ channels (1 s to -20 mV followed by 50 ms to -50 mV). In some experiments, calcium currents were elicited in the absence of the prepulse protocol to investigate the T-channels activity (see Fig. 3). Peak L-currents were normalized to cell capacitance (pA/pF), plotted as a function of membrane potential (I-V curves), and fitted according to Equation 1,


<UP>I</UP>=<UP>G<SUB>max</SUB></UP>×(<UP>V<SUB>m</SUB></UP>−<UP>V<SUB>rev</SUB></UP>)<UP>/</UP>(<UP>1</UP>+<UP>exp</UP>[(<UP>V<SUB>G1/2</SUB></UP>−<UP>V<SUB>m</SUB></UP>)<UP>/</UP>k<SUB><UP>G</UP></SUB>]) (Eq. 1)
where Vrev is the extrapolated reversal potential of the L-current, Vm is the membrane potential during the test pulse, Gmax is the maximal L-channel conductance, VG 1/2 is the voltage for half-activation of Gmax, and kG is a slope factor.

Immobilization-resistant intramembrane charge movements were measured following the prepulse protocol and blockade of ionic Ca2+ currents by the addition of 0.5 mM Cd2+ + 0.2 mM La3+ to the extracellular recording solution (12). The amount of immobilization-resistant charge movement was estimated by integrating the transient of charge that moved outward after the onset of the test pulse (Qon) and subsequently normalized to cell capacitance (nC/µF). The magnitude of the maximum immobilization-resistant charge movement (Qmax) was estimated by fitting the Qon data according to Equation 2,
<UP>Q<SUB>on</SUB></UP>=<UP>Q<SUB>max</SUB>/</UP>(<UP>1</UP>+<UP>exp</UP>[(<UP>V<SUB>Q1/2</SUB></UP>−<UP>V<SUB>m</SUB></UP>)<UP>/</UP>k<SUB><UP>Q</UP></SUB>]) (Eq. 2)
where Vm, VQ 1/2, and kQ have their usual meanings with regard to charge movement.

Measurements of Intracellular Ca2+ Transients-- Changes in intracellular Ca2+ were recorded with Fluo-3 as described previously (13, 10). Briefly, the salt form of the dye was added to the internal recording solution (see below). After rupture of the cell membrane and entry into the whole-cell mode, a waiting period of ~5 min was used to allow the dye to diffuse into the cell interior. A 75 watt xenon bulb and high-speed DeltaRAM illuminator (Photon Technology Incorporated, Monmouth Junction, NJ) were used to excite the dye (480 ± 20 nm) present in a small rectangular region of the voltage-clamped myotube. A computer-controlled shutter was used to eliminate illumination during intervals between test pulses. Fluorescence emission was measured using a dichroic long-pass mirror centered at 505 nm, an emission filter centered at 535 ± 25 nm, and a photomultiplier detection system operating in analogue mode (analogue filter set at 0.5 ms) (Photon Technology Incorporated). The background fluorescence was measured and canceled by analogous subtraction. Fluorescence traces are expressed as Delta F/F, where F represents the baseline fluorescence immediately prior to depolarization, and Delta F represents the fluorescence change from baseline. Fluorescence amplitudes at the end of the test pulses were plotted as a function of the membrane potential and fitted according to Equation 3,


&Dgr;<UP>F/F</UP>=(&Dgr;<UP>F/F<SUB>max</SUB></UP>)<UP>/</UP>{<UP>1</UP>+<UP>exp</UP>[(<UP>V<SUB>F1/2</SUB></UP>−<UP>V<SUB>m</SUB></UP>)<UP>/</UP>k<SUB><UP>F</UP></SUB>]} (Eq. 3)
where Delta F/Fmax is the calculated maximal fluorescence change, VF 1/2 is the midpoint potential, and kF is the slope factor.

Recording Solutions-- Ionic and gating currents were recorded using a pipette solution containing (in mM): Cs-Aspartate (140), MgCl2 (5), Cs2EGTA (10), and HEPES (10), pH 7.4. The external solution contained (in mM): triethylammonium chloride (145), CaCl2 (10), tetrodotoxin (0.003), and HEPES (10), pH 7.4. For measurements of intracellular Ca2+ transients, the pipette solution contained (in mM): Cs-aspartate (145), CsCl (10), Cs2EGTA (0.1), MgCl2 (1.2), MgATP (5), K5Fluo-3 (0.2), HEPES (10), pH 7.4. The external calcium current recording solution was supplemented with 0.5 mM CdCl2 + 0.2 mM LaCl3 for measurements of intramembrane charge movement.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

We have previously shown that compared with normal myotubes, the maximum immobilization-resistant intramembrane charge movement (Qmax) is ~40% smaller in dyspedic myotubes (3). Moreover, Qmax remains unchanged even 2-4 days following reintroduction of RyR-1 into dyspedic myotubes (2, 3). We have tested whether long-term expression of RyR-1 in dyspedic myotubes restores Qmax to a value comparable with that of normal myotubes. Fig. 1 compares immobilization-resistant charge movements in dyspedic myotubes either 3 or 6 days after nuclear injection of RyR-1 cDNA (3d-RyR-1 and 6d-RyR-1, respectively). In agreement with previous reports, 3d-RyR-1 expressing dyspedic myotubes exhibited Qmax values similar to those of uninjected dyspedic myotubes (Fig. 1C and Table I). However, charge movements recorded from 6d-RyR-1-expressing dyspedic myotubes were significantly larger than those recorded from either uninjected dyspedic myotubes or 3d-RyR-1-expressing myotubes. On average, Qmax increased 85% 6 days after nuclear microinjection of dyspedic myotubes with RyR-1 cDNA (Fig. 1 and Table I). Fig. 1C shows the entire time course of the RyR-1-mediated increase in Qmax. A progressive increase in Qmax was found each day between 3 and 6 days following RyR-1 cDNA injection (gray bars). The protracted time course of this effect is consistent with the initiation of a slow process, such as activation of gene transcription and subsequent protein synthesis, rather than a simple redistribution of previously translated but sequestered voltage sensors. By contrast, Qmax values recorded from uninjected dyspedic myotubes during a parallel time did not exhibit a significant change in Qmax (white bars). Thus, the increase in DHPR expression (Qmax) requires long-term reintroduction of RyR-1 and therefore does not arise from a general increase in L-channel expression during prolonged culture. Interestingly, the 6d-RyR-1-expressing dyspedic myotubes exhibit Qmax values that were nearly identical to those obtained from normal myotubes (Table I; see also Ref. 3). Thus, the long-term presence of RyR-1 strongly influences the number of sarcolemmal DHPRs in skeletal myotubes.


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Fig. 1.   Immobilization-resistant charge movement is increased in dyspedic myotubes following long-term expression of the skeletal muscle ryanodine receptor (RyR-1). A, immobilization-resistant charge movements recorded from two RyR-1-expressing dyspedic myotubes. Dyspedic myotubes were injected with RyR-1 cDNA either 3 days (3d-RyR-1; empty squares, left) or 6 days (6d-RyR-1; filled squares, right) before electrophysiological recordings. B, voltage dependence of charge movements recorded as in A. The charge movement occurring during the test pulses (Qon) was integrated and plotted as a function of the membrane potential (Vm). Data were obtained from twenty 3-day and eighteen 6-day RyR-1-expressing dyspedic myotubes. The average values (± S.E.) for the parameters obtained by fitting each myotube within a group separately to Eq. 2 are given in Table I (Q-V). The solid lines were generated using Eq. 2 and the corresponding Q-V parameters given in Table I. C, time-dependent effect of RyR-1 on the magnitude of the maximum immobilization-resistant charge movement (Qmax). The Qmax values were estimated from dyspedic myotubes that were previously (3-6 days) injected with RyR-1 cDNA (gray bars). Qmax did not vary significantly (p > 0.1) for non-injected dyspedic myotubes investigated during the same period of time (white bars). Data were obtained from a minimum of six (dyspedic, 5 days) and a maximum of 18 (RyR-1, 6 days) myotubes for each condition.

                              
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Table I
Parameters of fitted I-V and Q-V curves
Values represent mean ± S.E. I-V and Q-V data are from the number of myotubes indicated in column n. The number of experiments used to calculate Gmax/Qmax are indicated in parenthesis. Values of Gmax and Qmax were obtained by fitting each myotube within a group separately to the appropriate equation (I-V, Eq. 1; Q-V, Eq. 2). RyR-1 and E4032A-expressing dyspedic myotubes were subject to patch-clamp experiments either three (3d) or six (6d) days after cDNA microinjection. Qdys (see Ref. 12) was not subtracted from Qmax in calculations of Gmax/Qmax as done previously (2), because Idys does not appear to be present in normal or dyspedic myotubes (3).

The ability of DHPRs to function efficiently as Ca2+ permeable L-channels is significantly enhanced by the presence of RyR-1 (retrograde signal of EC coupling; Refs. 2, 3). To investigate whether or not the additional sarcolemmal DHPRs observed upon prolonged reintroduction of RyR-1 in dyspedic myotubes are functionally coupled to RyR-1, we compared L-current magnitudes in 3d-RyR-1- and 6d-RyR-1-expressing dyspedic myotubes. Fig. 2A shows representative L-currents that were obtained from 3d-RyR-1 (left) and 6d-RyR-1 (right) expressing myotubes. As illustrated in Fig. 2, L-current density was significantly greater for 6d-RyR-1-expressing myotubes compared with 3d-RyR-1-expressing myotubes. On average, (Fig. 2B) peak L-current density increased from -8.5 ± 0.4 pA/pF (3 days; empty squares) to -12.5 ± 1.2 pA/pF (6 days; filled squares). This increase in L-current density arose primarily from a ~30% increase in the maximal conductance of L-channels (Gmax) and was not accompanied by significant alterations in the voltage-dependence of L-channel activation (VG 1/2 and kG, see also Table I). Normal myotubes and 6d-RyR-1-expressing dyspedic myotubes displayed nearly identical peak L-current densities (-12.5 ± 1.2 pA/pF Vs -12.9 ± 1.4 pA/pF) and both Gmax and Qmax values (Table I; see also Ref. 3).


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Fig. 2.   Long-term expression of RyR-1 increases peak L-type Ca2+ current density. A, representative L-currents recorded from either 3d-RyR-1 (left; empty squares) or 6d-RyR-1 (right; filled squares) expressing dyspedic myotubes. B, average peak L-current density recorded as in A and plotted as a function of membrane potential (Vm). Data were obtained from 14 3d-RyR-1- and 14 6d-RyR-1-expressing dyspedic myotubes. The average values for Gmax, VG 1/2, and kG obtained by fitting each myotube within a group separately to Eq. 1 are given in Table I. The solid lines were generated using Eq. 1, and the corresponding I-V parameters. C, 3d (n = 9) and 6d (n = 10) RyR-1-expressing dyspedic myotubes exhibited similar current-to-charge ratios. Current-to-charge ratios were calculated by dividing the peak calcium current at each membrane potential by the maximum immobilization-resistant charge movement (Qmax) that was determined for each myotube.

To estimate the absolute levels of retrograde EC coupling for 3d-RyR-1- and 6d-RyR-1-expressing dyspedic myotubes, we normalized the L-current amplitudes by their respective Qmax values, to obtain the current-to-charge ratio (3). Following this normalization procedure, current-to-charge ratios were found to be similar for 3d-RyR-1- and 6d-RyR-1-expressing myotubes at every membrane potential (Fig. 2C). In addition, both experimental conditions exhibited nearly identical maximal conductance-to-charge ratios (Gmax/Qmax), which were similar to the corresponding value obtained for normal myotubes (Table I; see also Ref. 3). Consequently, no significant differences were found in either the current-to-charge or conductance-to-charge (Gmax/Qmax) ratios for normal myotubes or either 3d-RyR-1- or 6d-RyR-1-expressing myotubes. These results strongly suggest that a similar proportion of DHPRs interact with ryanodine receptors (as judged by the absolute levels of retrograde coupling) under these three conditions. Thus, the additional sarcolemmal DHPRs observed following long-term RyR-1 reintroduction into dyspedic myotubes represent primarily functional or RyR-1-coupled DHPRs.

Normal myotubes express two main types of voltage-dependent calcium channels in the sarcolemma: L-channels and T-type calcium channels (T-channels). L-channels and T-channels exhibit distinct biophysical and pharmacological properties including thresholds of activation, rates of channel activation and inactivation, unitary conductance, and sensitivity to dihydropyridines (14, 15). To investigate whether the presence of RyR-1 also regulates T-channel expression, we compared T-currents recorded from normal and dyspedic myotubes (Fig. 3). T-channel activity was dissected by recording calcium currents in the absence (total ICa = T-current + L-current; Fig. 3, A-a, B-a, C-a) and presence of a conditioning prepulse (12) designed to inactivate T-channels (L-current only; Fig. 3, A-b, B-b, C-b). T-currents were then revealed following offline subtraction of L-currents from the total calcium current (Fig. 3, A-c, B-c, C-c).


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Fig. 3.   T-type Ca2+ current density is similar for normal and dyspedic myotubes. Representative calcium currents recorded from a normal (A) and a dyspedic (B) myotube in the absence (a) and presence (b) of a conditioning prepulse used to inactivate T-channels (see "Experimental Procedures"). Subtraction of currents obtained in a minus those obtained in b was used to isolate the rapidly inactivating T-type Ca2+ currents (c). Outward gating current transients were truncated for clarity. C, voltage dependence of calcium currents recorded as in A and B. Data were obtained from 17 normal (filled circles) and 22 dyspedic myotubes (empty circles).

Fig. 3C shows average current to voltage relationships of total ICa (a), L-current (b), and T-current (c). I-V curves were obtained from 17 normal (filled circles) and 22 dyspedic myotubes (empty circles). In normal myotubes, total ICa exhibits a prominent shoulder that divides the I-V curve at ~0 mV into the following two components: 1) a low-threshold component (more prominent at negative membrane potentials) and 2) a high threshold component (most clearly resolved at positive potentials). Application of the conditioning prepulse selectively eliminates the low-threshold component, without significantly affecting the high-threshold component (b, L-current). The low-threshold, rapidly inactivating component was then dissected by subtracting b from a (c, T-current). As reported previously (2, 4, 3), dyspedic myotubes exhibit very modest L-currents compared with those recorded from normal myotubes (Fig. 3, C-b). This arises from a 5-fold reduction in macroscopic channels conductance and an ~45% reduction in the number of L-channels in the sarcolemma. Interestingly, average T-channel current density was similar for both normal and dyspedic myotubes (Fig. 3, C-c). Apparently, only the number of sarcolemmal L-channels and not the number of T-channels are significantly influenced by the presence or absence of RyR-1. Thus, the RyR-1-mediated increase in L-channel expression does not represent a general overall increase in ion channel biosynthesis.

Introduction of RyR-1 into dyspedic myotubes restores the following two important Ca2+ homeostatic mechanisms: 1) an SR Ca2+ release pathway or skeletal-type EC coupling and 2) a Ca2+ influx pathway that is manifested as a ~10-fold increase in L-type current magnitude (2, 3). Because elevations in intracellular calcium concentration ([Ca2+]i) in response to activation of voltage- or ligand-gated Ca2+ channels are known to influence a variety of biochemical process in excitable cells (16), we investigated whether the RyR-1-mediated increase in DHPR expression requires restoration of the Ca2+ release pathway, Ca2+ influx pathway, or both.

Substitution of an alanine residue for a highly conserved glutamate residue in either RyR3 (E3885A) or RyR-1 (E4032A) dramatically reduces Ca2+ activation of the resulting SR Ca2+ release channel (17, 18). In addition, O'Brien et al. (8) found that expression of E4032A in dyspedic myotubes preferentially restores skeletal L-current activity (i.e. retrograde coupling), but not robust, voltage-activated SR Ca2+ release (i.e. orthograde coupling). We exploited the ability of E4032A to preferentially restore the retrograde coupling to determine the relative importance of the Ca2+ influx and release pathways on the RyR-1-mediated increase in DHPR expression. Fig. 4 illustrates that expression of the E4032A mutant in dyspedic myotubes fully restored the Ca2+ influx pathway (L-current or retrograde coupling) but not the Ca2+ release pathway (orthograde coupling). Fig. 4A shows L-currents (lower traces) and intracellular calcium transients (upper traces) recorded simultaneously from a normal myotube (left), a 3-day RyR-1-expressing dyspedic myotube (middle), and a 3-day E4032A-expressing dyspedic myotube (right). The 3d-RyR-1- and the 3d-E4032A-expressing dyspedic myotubes exhibited similar L-current densities, which are ~45% smaller than L-currents recorded from normal myotubes (Fig. 4A). As exemplified by the representative experiments shown in Fig. 4A, normal and 3d-RyR-1-expressing dyspedic myotubes displayed similar robust voltage-activated Ca2+ transients, whereas 3d-E4032A-expressing dyspedic myotubes exhibited only very modest voltage-activated Ca2+ transients. On average, the presence of the E4032A mutation caused an ~7-fold reduction in maximal SR Ca2+ release compared with wild-type RyR-1 (Fig. 4B). Interestingly, E4032A-expressing dyspedic myotubes exhibited a slight (~20 mV), but significant (p < 0.01) depolarizing shift in VF 1/2 compared with those obtained from either normal myotubes or RyR-1-expressing dyspedic myotubes. This observation is consistent with the notion that the E4032A mutation stabilizes a SR Ca2+ release channel closed state(s), thus causing the requirement of stronger depolarizations to activate the release channel. Nevertheless, Ca2+ transients recorded from normal myotubes, as well as dyspedic myotubes expressing either RyR-1 or E4032A, each exhibited a sigmoidal voltage-dependence demonstrating the presence of a skeletal-type (as opposed to a Ca2+ influx-dependent) EC coupling mechanism (Fig. 4C).


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Fig. 4.   Expression of the E4032A mutation in RyR-1 in dyspedic myotubes preferentially restores retrograde coupling. A, simultaneous recordings of L-currents (bottom traces) and calcium transients (top traces) obtained from a normal myotube (left), a 3-day RyR-1-expressing dyspedic myotube (3d-RyR-1; center) and a 3-day E4032A-expressing dyspedic myotube (3d-E4032A; right). L-currents and calcium transients were elicited by 30 ms test pulses to the indicated membrane potentials (mV). B, voltage-dependence of calcium transients as recorded in A. Calcium transient amplitudes were measured at the end of the test pulses (30 ms) and plotted versus test pulse amplitude (Vm). Experimental data were fitted according to Eq. 3. The following Delta F/Fmax, VF 1/2 (mV) and kF (mV) parameters were obtained: 1.6 ± 0.3, 6.0 ± 3.2, 6.3 ± 1.1 for normal (n = 7); 1.4 ± 0.2, 8.7 ± 2.1, 6.0 ± 0.8 for 3d-RyR-1 (n = 12); and 0.2 ± 0.1, 23.2 ± 4.2, 5.1 ± 1.5 for 3d-E4032A (n = 5) myotubes, respectively. C, calcium transient amplitudes (recorded as in B) were divided by their respective maximal fluorescence change (peak Delta F/Fmax) and plotted as a function of the membrane potential (Vm).

As shown in Fig. 4, the E4032A mutation in RyR-1 provides a powerful tool for dissecting the putative roles of calcium influx and calcium release on functional DHPR expression. To test if the E4032A mutant mimics the ability of wild-type RyR-1 to increase DHPR functional expression, we compared L-current densities (Fig. 5, A and B) and immobilization-resistant charge movements (Fig. 5, C and D) for 3-day E4032A- and 6-day E4032A-expressing dyspedic myotubes. Remarkably, no significant differences were found in the peak L-current density, Gmax, or Qmax values (Fig. 5, B and D; Table I). In addition, the voltage-dependence of charge movements and L-current activation were similar for 3-day E4032A- and 6-day E4032A-expressing dyspedic myotubes (Fig. 5 and Table I). Thus, despite the restoration of large L-currents (i.e. retrograde coupling), the E4032A mutant failed to increase the number of functional L-channels over a time interval during which wild-type RyR-1 expression caused a ~2-fold increase in DHPR expression. These data indicate that restoration of the Ca2+ release pathway, rather than the Ca2+ influx pathway, is required for the ability of RyR-1 to increase functional DHPR expression in skeletal myotubes.


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Fig. 5.   Long-term expression of the E4032A mutant in dyspedic myotubes fails to increase the number of functional L-channels. Representative family of L-currents (A) and immobilization-resistant charge movements (C) recorded from 3-day (3d-E4032A; left; empty triangles) and 6-day (6d-E4032A; right; filled triangles) E4032A-expressing dyspedic myotubes. B, voltage-dependence of L-currents recorded as in A, plotted versus membrane potential (Vm), and fitted by Eq. 1 (continuous line). Experimental data were obtained from 7 3d-E4032A (empty triangles) and 12 6d-E4032A (filled triangles) myotubes. The parameters obtained by fitting each 3d-E4032A- and 6d-E4032A-expressing dyspedic myotube separately to Eq. 1 were combined to generate the average values (± S.E.) shown in Table I (3-6d-E4032A). The smooth solid line though the data points was generated using Eq. 1, and the corresponding parameters give in Table I. The I-V curves for 3d-RyR-1- and 6d-RyR-1-expressing dyspedic myotubes (dotted lines) from Fig. 2 are illustrated for comparison. D, voltage-dependence of immobilization-resistant charge movements. Data were obtained from a total of ten 3-day (empty triangles) and twelve 6-day (filled triangles) E4032A-expressing dyspedic myotubes. The parameters obtained by fitting each 3d-E4032A and 6d-E4032A expressing dyspedic myotube separately to Eq. 2 were combined to generate the average values (± S.E.) shown in Table I (3-6d-E4032A). The smooth solid line though the data points was generated using Eq. 2 and the corresponding parameters give in Table I. The Q-V curves for 3d-RyR-1- and 6d-RyR-1-expressing dyspedic myotubes (dotted lines) from Fig. 1 are illustrated for comparison (dotted lines).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

In this study, we have demonstrated that long-term expression of RyR-1 in dyspedic myotubes increases functional DHPR expression in the sarcolemma (i.e. increased both peak L-current and maximal intramembrane charge movement). The influence of RyR-1 appears to be restricted to L-channel expression because T-current density was similar for both normal and dyspedic muscle. Interestingly, long-term expression of E4032A, a mutant RyR-1 that preferentially affects the orthograde signal of E-C coupling (i.e. fully restores L-channel activity but not SR Ca2+ release) failed to increase functional DHPR expression. These results suggests that Ca2+ release through SR Ca2+ release channels (but not Ca2+ influx through L-channels) influences a signaling pathway in myotubes that promotes the expression of functional L-type Ca2+ channels.

There is increasing evidence to suggest that the expression level of skeletal muscle DHPRs is subject to changes in intracellular Ca2+. In amphibian (frog) skeletal muscle, long-term blockade of skeletal L-channels increases DHP binding and charge movement, consistent with the notion that Ca2+ influx inhibits DHPR expression in frog skeletal muscle (6). However, opposing observations have been made using mammalian skeletal muscle preparations. Specifically, elevations in extracellular Ca2+ (7) increase L-type tail current density and intramembrane charge movement in mammalian (rat) cultured myotubes. Our results support the observations of Renganathan et al. (7), but appear to be in contrast to the postulated role of Ca2+ influx in frog skeletal muscle (6). However, numerous differences between DHPR function in amphibian and mammalian skeletal muscle have been identified (e.g. molecular determinants of slow activation, regulation by cAMP-dependent protein kinase A, presence of Qgamma , prepulse-induced kinetic acceleration; see O'Connell and Dirksen, Ref. 19 and Melzer et al., Ref. 1 for review). Thus, it is conceivable that the signaling pathways that modulate functional DHPR surface expression represent yet another fundamental difference between mammalian and amphibian skeletal muscle. Nevertheless, it will be interesting to determine whether SR Ca2+ release also influences functional DHPR expression in frog skeletal muscle.

The precise molecular mechanism(s) by which SR Ca2+ release increases the number of functional sarcolemmal DHPRs is currently unclear. Conceivably, alterations in myoplasmic Ca2+ homeostasis could alter pathways that control DHPR alpha 1-subunit (or a DHPR regulatory protein such as the beta -subunit) gene transcription (16), message stabilization (20), and/or degradation. Global elevations in resting intracellular Ca2+ levels have been reported to increase L-type Ca2+ channel expression in both cardiac (21) and skeletal muscle cells (7). However, Ca2+-mediated alterations in gene transcription may not necessarily require global changes in resting intracellular Ca2+ levels. Under certain conditions, activity-dependent gene transcription has been demonstrated to be activated by privileged local Ca2+ signaling mechanisms. For example, in hippocampal neurons, Ca2+ influx through neuronal L-type Ca2+ channels activates the nuclear translocation of specific Ca2+-dependent transcription factors, such as CREB (22, 23) and NF-ATc4 (24). A similar privileged role for Ca2+ influx through L-type Ca2+ channels in CREB activation has recently been observed in vascular smooth muscle (25). This elegant study demonstrated that depolarization-induced increases in Ca2+ influx through voltage-dependent Ca2+ channels promotes CREB phosphorylation (P-CREB) and results in increased c-fos mRNA levels in intact mouse cerebral arteries. Interestingly, Ca2+ release through RyRs in cerebral arteries reduced P-CREB and c-fos mRNA levels, presumably by causing membrane hyperpolarization following Ca2+ spark-mediated activation of Ca2+-sensitive K+ channels (25, 26). Thus, at least one mechanism for activity-dependent control of gene expression in excitable cells involves the privileged ability of Ca2+ influx through voltage-gated Ca2+ channels to fine-tune protein expression during a process referred to as excitation-transcription coupling (27).

Our results demonstrate that long-term expression of the skeletal muscle RyR-1 appears to modulate functional DHPR expression through a novel form of excitation-transcription coupling. Because SR Ca2+ release is an essential component of Ca2+ signaling in skeletal muscle cells, RyR-1 activity could serve a central role in regulating protein expression in skeletal muscle. Insulin-like growth factor-1 induces skeletal muscle hypertrophy through activation of the Ca2+-dependent transcription factor NF-ATc1 in myotubes (28, 29) and promotes L-channel gene expression in rat skeletal muscle (30). Our data provide strong evidence for a novel pathway that involves the privileged ability of Ca2+ released through RyR-1s, rather than influx through L-type Ca2+ channels, to strongly influence the degree of functional DHPR expression in cultured myotubes. Thus, it will be important for future studies to determine the precise downstream molecular components (e.g. NF-ATc1, CREB) that are activated by Ca2+ released through RyR-1s and if this pathway is recruited by growth factors such as insulin-like growth factor-1. Independent of the precise molecular mechanism, our data demonstrate the requirement for robust SR Ca2+ release in the up-regulation of the number of functional DHPRs and provides an explanation for the reduction in DHPR levels observed in both dyspedic muscle homogenates (4, 5) and cultured dyspedic myotubes (3).

    ACKNOWLEDGEMENTS

We thank Drs. Kurt G. Beam and Paul D. Allen for providing us access to the dyspedic mice used in this study as well as for their advice and continued support.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant AR44657 (to R. T. D.) and a Neuromuscular Disease research grant (to R. T. D.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger Recipient of Consejo Nacional de Ciencia y Technologia (CONACYT) Postdoctoral Fellowship 990236.

§ To whom correspondence should be addressed. Tel.: 716-275-4824; Fax: 716- 273-2652; E-mail: Robert_Dirksen@URMC.rochester.edu.

Published, JBC Papers in Press, January 22, 2001, DOI 10.1074/jbc.M009685200

    ABBREVIATIONS

The abbreviations used are: DHPR, dihydropyridine receptor; L-type Ca2+ channel, L-channel; EC, excitation-contraction; SR, sarcoplasmic reticulum; RyRs, ryanodine receptors; CREB, cAMP-response element-binding protein.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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