A malignant hyperthermia–inducing mutation in RYR1 (R163C): alterations in Ca²⁺ entry, release, and retrograde signaling to the DHPR

In skeletal muscle, excitation–contraction coupling (ECC) is initiated by the activation of the L-type Ca²⁺ channel or 1,4-dihydropyridine receptor (DHPR). The DHPR in turn activates Ca²⁺ release from the SR Ca²⁺ release channel (RYR type 1 [RYR1]). The communication between the two channels is rapid and does not require Ca²⁺ entry through the DHPR, and it is believed that there is a physical interaction between the two proteins (Armstrong et al., 1972; Tanabe et al., 1990; Dirksen and Beam, 1999). In addition to the “orthograde” signal that triggers gating of RYR1, a “retrograde” signal from RYR1 to the DHPR was revealed by the observation that L-type currents of dyspedic (RYR1-null) myotubes were substantially smaller than L-type currents of wild-type (WT) myotubes, despite similar surface membrane expression of the L-type channel (Nakai et al., 1996).

Malignant hyperthermia (MH) is a fulminant pharmacogenetic life-threatening syndrome, which occurs when susceptible individuals are exposed to triggering factors, which include halogenated inhalation anesthetics and/or depolarizing muscle relaxants like succinylcholine (López et al., 1985, 1988; Nelson, 2001, 2002; Treves et al., 2005). The syndrome is associated with massive increases in intracellular Ca²⁺ ([Ca²⁺]_i) in response to exposure to the triggering agent (López et al., 1988). ECC dysfunction is believed to be the underlying cause of MH susceptibility because of its linkage in humans to more than 122 mutations within 19q13, the gene that codes for RYR1 (Robinson et al., 2006), or two characterized mutations within 1q31-32, the gene that codes Ca_V1.1 (Monnier et al., 1997; Jurkat-Rott et al., 2000). MH-susceptible (MHS) pigs and mice possessing a missense mutation (R614C [pigs] and R163C or Y522S [knock-in mice]) in RYR1 develop fulminant episodes when exposed to triggering agents and heat stress (López et al., 1988; Chelu et al., 2006; Yang et al., 2006). In addition, there is evidence that the R163C mutation not only influences RYR1 channel properties (Yang et al., 2003, 2006), but also potentiates depolarization-induced Ca²⁺ entry (Cherednichenko et al., 2004, 2008; Yang et al., 2007b). However, exactly how MHS mutations potentiate depolarization-induced Ca²⁺ entry and modify important aspects of orthograde and retrograde signaling between DHPR and RYR1 is poorly understood.

In the present study, we have examined how an MH mutation in RYR1 (R163C) affects the voltage dependence, time course, and extracellular Ca²⁺ dependence of the Ca²⁺ transient during prolonged depolarization induced by high extracellular K⁺. In doing so, we demonstrate that the R163C mutation alters retrograde signaling between RYR1 and the DHPR, as manifested by delayed inactivation of the DHPR voltage sensor, and at this level of depolarization enhances sarcolemmal Ca²⁺ entry.

Primary myoblast cell lines were generated from the hind limb and forelimb muscles of neonatal heterozygous (Het) and homozygous (Hom) F2 R163C C57Bl6/129svJ mice and their WT littermates (Rando and Blau, 1994; Yang et al., 2006). Myoblasts were differentiated into myotubes by withdrawal of growth factors as described previously (Yang et al., 2003). Animals were housed, fed, and sacrificed according to the standards set by the Harvard Medical School Institutional Animal Care and Use Committee.

Differentiated WT and R163C myotubes were loaded with 5 µM Fluo-4-AM (Invitrogen) at 37°C for 30 min in mammalian Ringer’s solution as described previously (Yang et al., 2006). The myotubes were then washed several times with mammalian Ringer’s solution and transferred to a microscope (Nikon). Fluorescence emission was measured at 516 nm with a long-pass filter using a 40×, 1.3 NA objective lens. Data were collected with a 12-bit digital-intensified CCD at 23 fps (Mega12; Stanford Photonics) from regions consisting of 8–12 individual cells per well and analyzed using QED software (QED Imaging). The magnitude of integrated Ca²⁺ transient (defined by the average area under the Ca²⁺ transient during a 3-min depolarization elicited by elevating [K⁺] in the extracellular medium) was compared among genotypes.

Single-barrel capillary tubing (1.5 mm OD; TW150F-4; WPI) containing an internal filament was used in the construction of microelectrodes for measuring the resting membrane potential. The pipettes were backfilled with 3 M KCl (resistance of ∼15 MΩ) and used to impale individual WT and R163C myotubes to measure membrane potential (at room temperature) before and during the time that myotubes were incubated in 60 mM K⁺ for 180 s. Results were discarded if tip resistance was altered by cell membrane penetration or from myotubes with resting potential values less negative than −60 mV.

WT, R163C Het, or Hom R163C myotubes were exposed to high external potassium ([K⁺]_e) solutions in ascending or descending concentrations (10, 20, 40, and 60 mM KCl) for 180 s for each concentration separated by 180 s of rest. The concentration of 60 mM K⁺ was chosen for the remaining experiments because we have found that this concentration of K⁺ elicits a maximum Ca²⁺ response in skeletal muscle myotubes (Yang et al., 2007a). WT and Hom R163C myotubes were exposed to 20 mM caffeine for 10 s, and 180 s later to 60 mM [K⁺]_e for 180 s. During the 180-s period that the cell was depolarized, a second 10-s application of 20 mM caffeine was applied at intervals ranging from 20 to 200 s after the beginning of the 60-mM [K⁺]_e application. WT and R163C myotubes were exposed to 60 mM [K⁺]_e in the presence of 1.8 × 10⁻³ M Ca²⁺ (normal Ca²⁺) or 8.7 × 10⁻⁶ M Ca²⁺/5 mM Mg²⁺ (low Ca²⁺) solutions. The exposure to the low Ca²⁺ solution was done for 10 s before depolarization with 60 mM [K⁺]_e. WT and Hom R163C myotubes were exposed either to 10 µM nifedipine or 20 µM SKF-96365 for 10 s before depolarization with 60 mM [K⁺]_e. This time was chosen to minimize any effect on the amplitude of the Ca²⁺ transient that we observed with longer incubation times. WT and R163C Het and Hom myotubes were exposed to 60 mM [K⁺]_e first with normal Ca²⁺ solution, allowed to recover for 10 min, then with low Ca²⁺ solution, allowed to recover for 10 min, and then finally with low Ca²⁺ solution supplemented with 2 mM [Ni²⁺]_e; the responses were compared. WT and R163C Het and Hom myotubes were exposed to 60 mM [K⁺]_e for 10 s, and then the Ca²⁺ transient was interrupted by replacing the high [K⁺]_e solution with normal mammalian Ringer’s solution, followed by a second application of 60 mM [K⁺]_e 3, 5, or 10 s later.

Differentiated primary WT or R163C Hom myotubes were loaded with 5 mM Fura-2 AM (Invitrogen) for 30 min at 37°C and washed four times for 10 min, each time in imaging buffer. After a baseline was established, the external solution was switched to a Ca²⁺-free buffer containing a final concentration of 500 µM Mn²⁺ (in the presence of 5 mM external Mg²⁺) with or without the addition of 2 mM Ni²⁺, and then stimulated with 60 mM KCl. The cells were continuously imaged before and after KCl stimulation at an excitation wavelength of 360 ± 3 nm (the isosbestic point of Fura-2), and fluorescence intensity was monitored with a 510-nm long-pass filter through a 40× quartz objective using an intensified CCD camera (see above). Data were collected from 3–10 individual cells simultaneously at 15 frames per second. The upward deflection that is seen in all raw traces shown indicates the application of KCl and is an artifact caused by intracellular Ca²⁺ release because it is not possible to be precisely at the isosbestic point of Fura-2 using any filter system.

The composition of standard Ringer’s solution was (in mM): 125 NaCl, 5 KCl, 2 CaCl₂, 1.2 MgSO₄, 5 glucose, and 25 HEPES, pH 7.4. Pilot experiments were conducted with Het and Hom R163C and WT myotubes exposed to [K⁺]_e solutions in which the [K⁺] × [Cl⁻] was either maintained constant or not maintained constant. No significant difference was observed in the amplitude and kinetics of the Ca²⁺ transients elicited by the high K⁺ solutions under either condition; therefore, we did not maintain the [K⁺] × [Cl⁻] in the solutions used experimentally (Hodgkin and Horowicz, 1959). The NaCl concentrations were adjusted to maintain a total ionic strength [Na⁺] + [K⁺] constant at 130 mM. Caffeine was prepared by adding the desired amount directly into the Ringer’s solution or the corresponding high [K⁺]_e solutions.

All values are expressed as mean ± SEM, with the numbers in parentheses indicating the number of myotubes tested. Statistical analysis was performed using paired t test, unpaired t test, or one-way analysis of variance coupled with Tukey’s t test for multiple measurements to determine significance (P < 0.05). Rates of decay were calculated using the formula y(t) = ae^−kt.

Fig. 1 shows representative Ca²⁺ transients elicited from WT and R163C Het and Hom myotubes by a random order titration of [K⁺]_e (10, 20, 40, and 60 mM). The K⁺-evoked Ca²⁺ transients from the myotubes of all three genotypes had a magnitude and time course that was dependent on [K⁺]_e. The threshold [K⁺]_e needed to elicit Ca²⁺ transients was 10 mM for Het and Hom R163C myotubes, whereas it was 20 mM in WT cells (P < 0.05, Tukey’s multiple tests; Fig. 1). Independent of the [K⁺]_e concentration, the integrated magnitude of the Ca²⁺ transients showed the rank order as Hom >> Het > WT. WT and R163C myotubes respond to prolonged [K⁺]_e exposures with an initial brief Ca²⁺ peak followed by a spontaneous exponential decay. This spontaneous decay was seen in all myotubes tested, despite the fact that depolarization of the plasma membrane was maintained throughout the exposure. At the maximum [K⁺]_e concentration tested (60 mM), the average integrals of the Ca²⁺ transients were 1.8 and 3.6 times greater in Het and Hom R163C myotubes in relation to WT myotubes, and the peak rates of Ca²⁺ transient decay were 0.107 ± 0.004 s⁻¹ (n = 11), 0.068 ± 0.004 s⁻¹ (n = 13), and 0.037 ± 0.003 s⁻¹ (n = 17) in WT and R163C Het and Hom myotubes, respectively (P < 0.001).

To make certain that membrane depolarization evoked by 60 mM [K⁺]_e was maintained at similar potentials among the myotubes used in the study, the membrane potential was recorded from each genotype before and during the 180-s depolarization. There was no significant difference (P > 0.05, nonpaired t test) in resting membrane potentials or potentials obtained during the 60-mM [K⁺]_e challenge, and the level of depolarization was sustained for the duration of the [K⁺]_e test, regardless of the myotube’s genotype (Table I).

To test the hypothesis that the spontaneous decay of the K⁺-evoked Ca²⁺ transients was the result of SR Ca²⁺ depletion or inactivation of RYR1 by high myoplasmic Ca²⁺ concentrations, we elicited SR Ca²⁺ release with a 10-s application of 20 mM caffeine before, during, and after exposure to 60 mM [K⁺]_e. Fig. 2 shows the response to caffeine at different intervals (20, 40, 80, 120, 160, and 200 s) after initiating a 180-s challenge to 60 mM [K⁺]_e. At each interval, caffeine produced a robust release of Ca²⁺ whose amplitude was not significantly different (P > 0.1) than that measured before the [K⁺]_e challenge, regardless of genotype. Thus, decay of the [K⁺]_e-triggered Ca²⁺ transient in WT and Hom R163C myotubes does not appear to be related to either depletion of SR Ca²⁺ stores (Hodgkin and Horowicz, 1960b) or inactivation of RYR1 (Smith et al., 1985; Suarez-Isla et al., 1986; Simon et al., 1991; Jong et al., 1995).

It is well established that a relatively short exposure to a low [Ca²⁺]_e medium has no effect on the peak K⁺ contracture tension; however, it greatly shortens the contracture time course, reducing the duration of the plateau and increasing the relaxation rate (Bianchi and Shanes, 1959; Frank, 1960; Caputo and Gimenez, 1967). Recently, the L-type current has been identified as the major means of Ca²⁺ entry during prolonged depolarizations or repetitive activity (Bannister and Beam, 2009; Bannister et al., 2009). Such Ca²⁺ entry is enhanced in both Het and Hom R163C myotubes in response to [K⁺]_e depolarization and electrical pulse trains (Yang et al., 2007a; Cherednichenko et al., 2008). Here, we have examined the impact of the R163C mutation on external Ca²⁺ effects in more detail. Exposure of WT and Het and Hom R163C myotubes to low Ca²⁺ solution (8.7 × 10⁻⁶ M/5 mM Mg²⁺) did not modify initial Ca²⁺ peak, but it dramatically altered the time course of the Ca²⁺ transients, making their duration shorter and their rate of decay faster in all three genotypes (Fig. 3). The integral of the Ca²⁺ transient was reduced by 62 ± 5.8% (n = 9) in WT, 74 ± 7% (n = 14) in Het R163C, and by 80 ± 9.3% (n = 12) in Hom R163C in low [Ca²⁺]_e solution, and the rate of Ca²⁺ decay was increased from 0.107 ± 0.004 s⁻¹ (n = 11) to 0.233 ± 0.006 s⁻¹ (n = 9), from 0.068 ± 0.004 s⁻¹(n = 13) to 0.144 ± 0.004 s⁻¹ (n = 14), and from 0.037 ± 0.003 s⁻¹ (n = 17) to 0.137 ± 0.005 s⁻¹ (n = 12) in WT, Het, and Hom, respectively. The changes in the transient integral and rate of decay were fully reversible in a third depolarization for all genotypes, when [Ca²⁺]_e was restored in the bath solution (unpublished data).

Because removal of external Ca²⁺ would be expected not only to eliminate Ca²⁺ entry, but also to accelerate inactivation of the voltage sensor for ECC (Rios and Brum, 1987), we also examined the effect of two organic Ca²⁺ entry blockers, nifedipine and SKF-96365 (Almers and Palade, 1981; Rivet et al., 1989; Leung and Kwan, 1999; Araya et al., 2003; Bannister et al., 2009), in the presence of Ca²⁺ replete external solutions.

Exposure to 10 µM nifedipine in the presence of normal external Ca²⁺ concentrations did not affect Ca²⁺ transient amplitude, but it accelerated the mean rate of decay from 0.107 ± 0.004 s⁻¹ (n = 11) to 0.135 ± 0.005 s⁻¹ (n = 11) in WT (∼30%), and from 0.037 ± 0.003 s⁻¹ (n = 17) to 0.117 ± 0.002 s⁻¹ (n = 16) in Hom R163C (∼70%) myotubes (P < 0.001; Fig. 4 A). Exposure of WT and Hom R163C myotubes to 20 µM SKF-96365 did not lower Ca²⁺ transient amplitude, but it accelerated the mean rate of decay from 0.107 ± 0.004 s⁻¹ (n = 11) to 0.146 ± 0.005 s⁻¹ (n = 9) in WT myotubes (∼27%), and from 0.037 ± 0.003 s⁻¹ (n = 17) to 0.072 ± 0.004 s⁻¹ (n = 13) in Hom R163C (∼49%) myotubes (P < 0.001; Fig. 4 B). To test whether the effects of nifedipine or SKF96365 on the decay rate of the Ca²⁺ transients could be explained by their ability to partially depolarize the myotubes’ membrane at rest, or affect a change in the magnitude of the depolarization induced by 60 mM [K⁺]_e, membrane potentials were measured directly under these experimental conditions. Neither nifedipine nor SKF-96365 modified the resting membrane potential or the ability of 60 mM [K⁺]_e to depolarize the myotubes to the expected membrane potential (unpublished data).

Although both nifedipine and SKF-96365 accelerated Ca²⁺ transient decay, the decay rates remained different between WT and Hom myotubes, providing further support for the hypothesis that the R163C mutation affects the rate at which the DHPR transitions into the ECC-inactivated state. To further test this hypothesis, we examined the effect of replacement of external Ca²⁺ by Ni²⁺. Previous investigations in amphibian muscle have shown that when Ni²⁺ is used to replace Ca²⁺ in the extracellular medium, the normal time course of K⁺ contractures is maintained (Caputo, 1981; Lorković and Rüdel, 1983), despite the fact that Ni²⁺ blocks Ca²⁺ current conducted by amphibian skeletal L-type channels (Almers and Palade, 1981; Obejero-Paz et al., 2008). Fig. 5 shows superimposed averaged records from experiments in which WT and R163C Het and Hom myotubes were exposed to 60 mM [K⁺]_e under three different experimental conditions: in the presence of (1) 1.8 × 10⁻³ M Ca²⁺, (2) 8.7 × 10⁻⁶ M Ca²⁺, and (3) 8.7 × 10⁻⁶ M Ca²⁺ supplemented with 2 × 10⁻³ M Ni²⁺ (see Materials and methods for details). As shown above, the removal of Ca²⁺ significantly accelerated the rate of decay compared with the response in normal Ca²⁺ solutions. Importantly, the addition of Ni²⁺ to low [Ca²⁺]_e solutions also resulted in prolongation of the K⁺-evoked Ca²⁺ transients in WT, Het, and Hom myotubes, although it did not restore the rate of decay to control levels. Under all three conditions, the decay rate was significantly slower in Hom and Het myotubes compared with WT. If the decay rates of the Ca²⁺ transients in the presence of Ni²⁺ are taken to indicate the normal course of SR Ca²⁺ release without any influence from Ca²⁺ entry, the fraction of the total Ca²⁺ release that is presumably due to the inactivation of the DHPR caused by the lack of extracellular Ca²⁺ was 35.7% of the area under the curve in WT, 44.1% in Het R163C, and 64.3% in Hom R163C myotubes. These results suggest that the shortening of the Ca²⁺ transient in low Ca²⁺ solution was a combined consequence of accelerated inactivation of the DHPR voltage sensor and decreased depolarization-induced Ca²⁺ entry (see Table II for decay rates). Furthermore, these data show that the lack of extracellular Ca²⁺ affects the inactivation of the DHPR ECC voltage sensor in the Het and Hom R163C myotubes to a much greater extent than in WT myotubes.

We directly measured the differences in cation entry in response to membrane depolarization using the Mn²⁺ quench technique in Hom R163C and WT myotubes. Fig. 6 A shows representative Fura-2 emission traces from Hom R163C and WT myotubes before and after stimulation with 60 mM KCl in the presence and absence of Ni²⁺. As summarized in Fig. 6 B, these data show the average rate of Mn²⁺ quench measured during the linear phase of quench after KCl exposure was significantly greater in Hom R163C myotubes than in WT myotubes (P < 0.01). Furthermore, cation entry was almost completely abolished in both groups of myotubes by the addition of 5 mM Ni²⁺.

Depolarization converts resting DHPR voltage sensors into their active state, after which they spontaneously change conformation and transition into their inactive state. The amount of SR Ca²⁺ release in response to depolarization is proportional to the degree that the voltage sensors are in their active state (Dulhunty and Gage, 1988; Dulhunty, 1992), when the T-tubule is depolarized. WT and R163C Het and Hom myotubes were initially exposed to 60 mM [K⁺]_e for 10 s (Fig. 7, Int₁), and then returned to normal mammalian Ringer’s solution for 3, 5, or 10 s (Fig. 7, Δt), and then reexposed to 60 mM [K⁺]_e for 10 s (Fig. 7, Int₂). We observed that washout of the elevated K⁺ solution with normal Ringer’s solution immediately stopped the Ca²⁺ transient in all three genotypes, an effect that can be attributed to the repolarization of the plasma membrane. However, the amplitude of the Ca²⁺ transient triggered by reexposure to high [K⁺]_e was highly genotype dependent (Fig. 7, Int₂). The integral of the Ca²⁺ transient during lnt₂ in Hom R163C myotubes was not significantly different from the integral during Int₁ for all interpulse intervals (Fig. 7; P > 0.05, Tukey’s multiple tests). In Het R163C myotubes, lnt₂ was significantly smaller than Int₁ at 3 s, but not significantly different from Int₁ at 5 and 10 s (Fig. 7). In WT myotubes, on the other hand, Int₂ was significantly lower than Int₁ at all three intervals tested (i.e., 3, 5, and 10 s; P < 0.001). These results indicate that the DHPR voltage sensor inactivates more slowly in MHS myotubes than in WT.

There is general agreement that membrane depolarization elicited by high [K⁺]_e triggers intracellular Ca²⁺ transients in amphibian (Blinks et al., 1978; López et al., 1983; Snowdowne, 1985; Caputo and Bolaños, 1994) and mammalian skeletal muscle (López et al., 2005). Exposure of WT and R163C myotubes to prolonged high [K⁺]_e triggered Ca²⁺ transients characterized by a peak Ca²⁺ transient that varied with [K⁺]_e and subsided over time, despite the fact that the membrane remained depolarized.

As has been shown previously, the duration and the peak amplitude of a K⁺ contracture (Hodgkin and Horowicz, 1960b) or the corresponding Ca²⁺ transient (Caputo and Bolaños, 1994) depend on the [K⁺]_e and hence on membrane potential. The results of the present study confirm these essential features in all three genotypes studied (Fig. 1). Thus, the peak amplitude of the Ca²⁺ transient was smaller and the duration of the Ca²⁺ transient was greatly prolonged at low [K⁺]_e. Although the peak amplitude of the Ca²⁺ transient was enhanced with increasing [K⁺]_e, the duration of the transient decreases. The inverse relationship between [K⁺]_e and transient duration is related to the rate of inactivation of the voltage sensor of the DHPR (Caputo and Bolaños, 1994).

In agreement with previous work on dyspedic myotubes expressing RYR1 with any of seven MH-causing mutations (Yang et al., 2003, 2007a), we found in the present work that elevated K⁺ produced Ca²⁺ transients in primary R163C myotubes, which activated at lower membrane potentials and decayed more slowly than in WT myotubes. Thus, these alterations of Ca²⁺ handling may be a common consequence of many MH mutations of RYR1. These observations are in agreement with those reported by Gallant and co-workers (Gallant et al., 1982; Gallant and Lentz, 1992), in which the mechanical threshold for [K⁺]_e was lower in MHS than in WT pig skeletal muscle fibers.

Interestingly, the rate of decay of the Ca²⁺ transient elicited by [K⁺]_e was slower in Hom and Het R163C compared with WT myotubes. Our data show that the decay in the peak of the Ca²⁺ transient during membrane depolarization observed in WT and R163C myotubes is related neither to SR Ca²⁺ depletion (Hodgkin and Horowicz, 1960b) nor to inactivation of RYR1 due to the high Ca²⁺ present in the myoplasm (Smith et al., 1985; Suarez-Isla et al., 1986; Simon et al., 1991; Jong et al., 1995) because the application of 20 mM caffeine was able to induce a robust Ca²⁺ release in all genotypes at all time points during the decay phase of the Ca²⁺ transient during the prolonged K⁺ depolarization (Fig. 2). The most probable explanation for this observation is that an alteration in the conformation of the RYR1 tetramer caused by even one MHS allele affects both orthograde and retrograde coupling between the DHPR and RYR1 (Nakai et al., 1996, 1998). Specifically, our data demonstrate at least two retrograde effects of the R163C mutation on DHPRs that alter the properties of ECC. The first is that the threshold for activation of Ca²⁺ release induced by [K⁺]_e is lowered by the RYR1 MH mutation (this change is confirmed electrophysiologically in the companion paper [see Bannister et al. in this issue]). Second, the R163C RyR1 mutation slows the rate of inactivation of the DHPR voltage sensor for ECC (Figs. 1 and 7). Additionally, our data suggest that the R163C mutation increases Ca²⁺ entry during prolonged depolarization.

Previous reports have shown that skeletal muscle fibers can contract after electrical stimulation or under whole cell voltage clamp with short step depolarizations in low extracellular Ca²⁺ concentrations (Armstrong et al., 1972; Bolaños et al., 1986). However, the importance of extracellular Ca²⁺ both in the presence (Lüttgau and Spiecker, 1979; Caputo, 1981; Cota and Stefani, 1981) and absence of Ca²⁺ buffers (Caputo and Gimenez, 1967; Lüttgau and Spiecker, 1979) for maintenance of muscle K⁺ contractures has also been well demonstrated. Our data provide evidence that external Ca²⁺ plays a significant physiological role for the maintenance of Ca²⁺ transients induced by elevated [K⁺]_e in WT and R163C MHS myotubes (Fig. 3). In fact, the duration of the Ca²⁺ transient was shortened (increased rate of decay of peak Ca²⁺) in nominally Ca²⁺-free external solution or in the presence of Ca²⁺ entry blockers (Figs. 3 and 4) regardless of genotype. We have ruled out the possibility that the increased rate of decay in the presence of low [Ca²⁺]_e might be a consequence of reduced SR Ca²⁺ content resulting from prolonged membrane depolarization (Hodgkin and Horowicz, 1960a,b; Caputo and Gimenez, 1967), as the addition of 5 mM Mg²⁺ to the external solutions prevents this depolarization (unpublished data; see Materials and methods). Furthermore, if resting SR Ca²⁺ content was reduced by the lower [Ca²⁺]_e, the peak amplitude (early phase) of the Ca²⁺ transient during long depolarization would have also been reduced. Because a reduction in peak transient amplitude was not observed in our experiments on intact myotubes, it appears that SR Ca²⁺ stores were not significantly reduced by exposure to lower [Ca²⁺]_e. This also argues against the alternative explanation that the slower rate decay in R163C compared with WT could have been due to changes in electrical properties of the myotubes during high [K⁺]_e exposure, as was reported in amphibian muscle fibers (Stefani and Chiarandini, 1973). If this were the case, the peak transient amplitude would also be reduced (Fig. 1). The most likely explanation is that the main alteration caused by low [Ca²⁺]_e is an inactivation of the voltage sensor and a reduction in SR Ca²⁺ release with no change in the rate of Ca²⁺ removal from the myoplasm (Brum et al., 1988). The suggested mechanism for this is the existence of a high affinity Ca²⁺ binding site or sites on the external side of the DHPR (Ebata et al., 1990), which modulate its inactivation process (Rios and Brum, 1987). In addition, the removal of external Ca²⁺ compromises the Ca²⁺ entry, which sustains the time course of the Ca²⁺ transients in WT and R163C myotubes, induced by [K⁺]_e depolarization and long trains of electrical stimuli (Cherednichenko et al., 2004, 2008). This combined explanation is directly supported by our result that when Ni²⁺ was added to the low Ca²⁺ solution, the rate of decay of the Ca²⁺ transient was significantly retarded but not completely restored (Fig. 5). It is important to point out that the sarcolemma is not permeable to Ni²⁺, and when it is present it blocks the inward Ca²⁺ current during depolarization of amphibian skeletal muscle fibers (Palade and Almers, 1985). In this regard, we have found that 2 mM Ni²⁺ blocks the L-type Ca²⁺ current in both WT and R163C myotubes (unpublished data) and divalent cation entry during depolarization (Fig. 6). Interestingly, we found the contribution of extracellular Ca²⁺ to be more substantial in both Hom and Het R163C myotubes than in WT.

Exposure of WT and R163C myotubes to the L-type Ca²⁺ channel antagonist nifedipine or the nonspecific cation channel blocker SKF-96365 also accelerated the rate of decline of Ca²⁺ transients (Fig. 4) in the presence of extracellular Ca²⁺. The fact that we did not observe a significant reduction in the peak Ca²⁺ transient amplitude in either WT or R163C myotubes shows that peak Ca²⁺ transient amplitude is largely controlled by SR Ca²⁺ release, and the brief exposure to both drugs prevented any secondary effects of these drugs. Although both drugs are known to block Ca²⁺ entry via the DHPR (Bannister et al., 2009), nifedipine caused a more rapid decline in the Ca²⁺ transient because it also alters the inactivation of the DHPR voltage sensor.

The general model for ECC suggests that Ca²⁺ release in skeletal muscle is controlled by a voltage sensor, which is an integral function of the DHPR that initially activates and subsequently inactivates to tightly control SR Ca²⁺ release. Furthermore, it is known that the time course of K⁺-triggered contractures is determined by the onset and time course of the voltage sensor inactivation process (Caputo, 1976; Bolaños et al., 1986), which is initiated immediately after activation begins (Caputo, 1972a,b). From the experiments in which Ni²⁺ was substituted for Ca²⁺, it was shown that depolarization-induced Ca²⁺ entry is enhanced in Het and Hom R163C, but the majority of the difference in the rate of decay of the Ca²⁺ transient in Het and Hom R163C myotubes is due to prolonged SR Ca²⁺ release caused by either a delay in the onset or a reduction in the rate of DHPR voltage sensor inactivation.

The double K⁺ application protocol (Fig. 7) confirmed this idea by demonstrating that the second Ca²⁺ transient of the pair was significantly smaller in WT myotubes than in R163C myotubes with all delays. Furthermore, the rate of decay of the Ca²⁺ transient was slower in Hom than in Het R163C myotubes, suggesting that the conformation of R163C has a significant impact through retrograde signaling from RYR1 to the DHPR on the voltage sensor inactivation process, in addition to the retrograde signals that control L-type Ca²⁺ current. In the companion paper (Bannister et al., 2010), we have examined the effects of the R163C mutation in RYR1 on the biophysical properties of the skeletal muscle L-type Ca²⁺ channel.

This work was supported by a fellowship grant from the Fondation pour la Recherche Médicale (SPE20040901554 to E. Estève), the Muscular Dystrophy Association (MDA4155 to R.A. Bannister and MDA4319 to K.G. Beam), and the National Institutes of Health (NIH P01AR052534 to P.D. Allen and I.N. Pessah, and R01AR44750 to K.G. Beam).

Kenneth C. Holmes served as editor.