Malignant hyperthermia (MH) is an inherited pharmacogenetic disorder caused by mutations in the skeletal muscle ryanodine receptor (RyR1) and the dihydropyridine receptor (DHPR) α1S-subunit. We characterized the effects of an MH mutation in the DHPR cytoplasmic III-IV loop of α1S (R1086H) on DHPR-RyR1 coupling after reconstitution in dysgenic (α1S null) myotubes. Compared with wild-type α1S, caffeine-activated Ca2+ release occurred at approximately fivefold lower concentrations in nonexpressing and R1086H-expressing myotubes. Although maximal voltage-gated Ca2+ release was similar in α1S- and R1086H-expressing myotubes, the voltage dependence of Ca2+ release was shifted ∼5 mV to more negative potentials in R1086H-expressing myotubes. Our results demonstrate that α1S functions as a negative allosteric modulator of release channel activation by caffeine/voltage and that the R1086H MH mutation in the intracellular III-IV linker disrupts this negative regulatory influence. Moreover, a low caffeine concentration (2 mM) caused a similar shift in voltage dependence of Ca2+ release in α1S- and R1086H-expressing myotubes. Compared with α1S-expressing myotubes, maximal L channel conductance (Gmax) was reduced in R1086H-expressing myotubes (α1S 130 ± 10.2, R1086H 88 ± 6.8 nS/nF; P < 0.05). The decrease in Gmax did not result from a change in retrograde coupling with RyR1 as maximal conductance-charge movement ratio (Gmax/Qmax) was similar in α1S- and R1086H-expressing myotubes and a similar decrease in Gmax was observed for an analogous mutation engineered into the cardiac L channel (R1217H). In addition, both R1086H and R1217H DHPRs targeted normally and colocalized with RyR1 in sarcoplasmic reticulum (SR)-sarcolemmal junctions. These results indicate that the R1086H MH mutation in α1S enhances RyR1 sensitivity to activation by both endogenous (voltage sensor) and exogenous (caffeine) activators.
- excitation-contraction coupling
- calcium channel
- muscle disease
excitation-contraction (EC) coupling in skeletal muscle involves a mechanical signaling interaction between sarcolemmal dihydropyridine receptors (DHPRs) and ryanodine receptors (RyRs) located in the sarcoplasmic reticulum (SR) (for review, see Ref. 36). During EC coupling, sarcolemmal DHPRs convert an electrical impulse in the sarcolemma (i.e., an action potential) into a chemical signal (i.e., release of SR Ca2+ through activated RyRs), which in turn activates the contractile apparatus, ultimately resulting in contraction of the muscle fiber. The process by which DHPRs mediate this electrochemical conversion is referred to as the orthograde signal of EC coupling. The DHPR-RyR mechanical signaling interaction in skeletal muscle is bidirectional because the Ca2+ conducting activity and biophysical properties of the sarcolemmal DHPR are strongly influenced by its interaction with the skeletal muscle isoform (type 1) of the ryanodine receptor (RyR1) (2, 24, 40), a process referred to as the retrograde signal of skeletal muscle EC coupling (for review, see Ref. 10)
Malignant hyperthermia (MH) is an autosomal dominant pharmacogenetic syndrome that until recently was one of the main causes of death due to anesthesia. MH-susceptible individuals respond to potent inhalation anesthetics (e.g., halothane) and depolarizing skeletal muscle relaxants (e.g., succinylcholine) with skeletal muscle rigidity, hypermetabolism, lactic acidosis, hypoxia, and tachycardia and consequently with a dramatic rise of body temperature (27, 37). If not immediately reversed (via hyperventilation with 100% O2 and administration of dantrolene), the attack can rapidly lead to severe tissue damage and eventually death. Standardized in vitro contracture tests have been developed to detect MH susceptibility in suspected individuals. These tests determine the sensitivity of biopsied muscle to contractures induced by applications of caffeine and halothane. If contractures occur in the presence of normally subthreshold concentrations of caffeine and halothane, a diagnosis of MH susceptibility is made. In some cases, MH is associated with central core disease (CCD), a rare congenital myopathy of autosomal dominant inheritance in which affected individuals present with infantile hypotonia and delayed attainment of motor milestones (for review, see Ref. 11).
The RyR1 gene on chromosome 19q13.1 (MHS-1) clearly represents a primary molecular locus for both MH and CCD in humans, as mutations in the RyR1 gene have been linked to >50% of all MH families and most CCD families. Currently, >60 deletions and missense mutations in the RyR1 gene have been causally linked to MH and/or CCD. However, evidence also indicates that other MH-susceptible gene loci exist. For example, linkage of MH has also been made to chromosomes 17 (MHS-2), 7 (MHS-3), 3 (MHS-4), 1 (MHS-5), and 5 (MHS-6) (27, 35). However, MHS-5 mutations to a highly conserved arginine residue in the III-IV linker in the α1S-subunit of the skeletal muscle DHPR (R1086H and R1086C) represent the only specific MH-causing mutations that have so far been identified in a protein other than RyR1 (27, 39). The finding that MH is caused by mutations in both the skeletal muscle DHPR and RyR1, two key proteins of muscle EC coupling, strongly suggests that the pathogenesis of MH stems from a dysfunction in muscle EC coupling and that if mutations are to be found in other proteins they are likely to involve other members of the triad complex.
However, the effects of MH mutations in the DHPR α1S-subunit on the sensitivity of the intracellular Ca2+ release mechanism to activation and bidirectional DHPR-RyR1 signaling in skeletal muscle are unknown. Toward this end, we compared the Ca2+ channel activity and junctional targeting of wild-type and R1086H mutant DHPRs, as well as the sensitivity of SR Ca2+ release to activation by caffeine and the voltage sensor (orthograde coupling) after expression in skeletal myotubes derived from dysgenic mice, which lack a functional gene for α1S. Our results demonstrate that, consistent with the diagnostic MH disease phenotype, the sensitivity of RyR1 to activation by both membrane depolarization and pharmacological agents (e.g., caffeine) is significantly enhanced in R1086H-expressing dysgenic myotubes. In addition, our results indicate that the intracellular III-IV linker of the DHPR functions as a key negative allosteric modulator of SR Ca2+ release channel activation.
MATERIALS AND METHODS
Construction of mutant DHPR cDNAs.
The wild-type clone GFP-α1S was generated by fusing the α1S-subunit cDNA in frame to the COOH terminus of the coding region of a modified green fluorescent protein (GFP) contained in a proprietary mammalian expression vector (23). We previously showed (23) that the GFP tag does not alter the functional properties of α1S. To construct the R1086H point mutation in GFP-α1S [nucleotide numbers (nt) are given in parentheses and asterisks indicate restriction enzyme (RE) sites introduced by PCR with proofreading Pfu Turbo DNA polymerase (Stratagene)], a silent AflII* RE site (nt 3278) together with a base substitution (nt 3256) that created a triplet coding for histidine instead of arginine were introduced via the antisense primer by PCR. The XhoI-AflII* fragment (nt 2654–3278) produced by this PCR step was coligated with the PCR-generated AflII*-BglII fragment (nt 3278–4488) into the corresponding XhoI/BglII RE sites of plasmid GFP-α1S. To construct the R1217H point mutation in a GFP-tagged cardiac DHPR (GFP-α1C), two base substitutions (nt 3650, 3651) that created a triplet coding for His1217 were introduced by using the “gene SOEing” technique (26). The ApaI-EcoRV fragment (nt 3491–4351) generated by this fusion PCR was coligated with the AflII-ApaI fragment (nt 2690–3491) of α1C into the corresponding AflII/EcoRV RE sites of plasmid GFP-α1C. For simplicity, we refer in the text to these constructs as R1086H and R1217H, respectively. A GFP-free version of α1S(R1086H) was produced by eliminating the GFP cDNA by peripheral PstI RE sites before the cloning steps described above and was used in indo 1 experiments in intact myotubes (see Fig. 1). All sequences generated and modified by PCR were checked for integrity by sequence analysis (MWG Biotech, Ebersberg, Germany).
Cell culture and transfection.
Myotubes generated from the homozygous dysgenic (mdg/mdg) mouse cell line GLT were cultured as described previously (44). GLT cultures were transfected with GFP-tagged constructs at the onset of myoblast fusion (2–4 days after addition of differentiation medium) by using the liposomal transfection reagent FuGene according to the manufacturer’s protocol (Roche Diagnostics, Mannheim, Germany). Three to four days after transfection, myotubes were either fixed for immunocytochemistry analysis (see Fig. 5) or were used in patch-clamp experiments by identifying expressing cells via GFP fluorescence (see Figs. 2–4).
Measurements of electrically elicited and caffeine-induced Ca2+ release in intact myotubes.
For these experiments, cultures of dysgenic myotubes were generated from primary myoblasts as described previously (15). Dysgenic breeding mice were housed in a pathogen-free area at Brigham and Women’s Hospital. Dysgenic myoblasts were obtained from newborn dysgenic mice after rapid decapitation and spinal cord destruction following procedures that were reviewed and approved by the Harvard Medical School Institutional Animal Care and Use Committee. To maximize detection of Ca2+ release at threshold concentrations, caffeine response curves were monitored in intact indo 1-AM-loaded dysgenic myotubes expressing GFP-free variants of wild-type α1S and R1086H. Expression of wild-type α1S and R1086H DHPRs was achieved by nuclear microinjection of cDNAs encoding CD8 (0.2 μg/μl) and the appropriate DHPR expression plasmid (0.2 μg/μl) 4–6 days after initial plating of myoblasts (3). DHPR/CD8-expressing myotubes were subsequently identified 48–72 h after injection by incubation with CD8 antibody-coated beads (Dynabeads; Dynal ASA, Oslo, Norway). We previously found (3, 4) that coexpression of CD8 does not influence measurements of resting Ca2+, SR Ca2+ content, or orthograde/retrograde DHPR-RyR1 coupling.
Because caffeine fluorescence quenching of single-wavelength Ca2+ dyes (e.g., fluo 4) could obscure detection of Ca2+ release at threshold concentrations, caffeine response curves were obtained with indo 1, a ratiometric Ca2+ dye in which caffeine produces comparable quench of both emission wavelengths and thus is without effect on the resulting indo 1 ratio (43). Indo 1-loaded myotubes were bathed in a normal rodent Ringer solution [containing (in mM) 145 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, and 10 HEPES, pH 7.4] and excited at 350 nm with a DeltaRam illumination system (Photon Technology, Princeton, NJ). Fluorescence emission at 405 (F405) and 485 (F485) nm was collected with a photomultiplier detection system, and the results are presented as the ratio (R) of F405 to F485. Caffeine concentrations (0.1–30 mM) prepared in Ringer solution were applied directly to individual myotubes with a rapid perfusion system (Warner Instrument, Hamden, CT) that permits fast, local application of agonist as well as rapid washout with control solution (4, 5). During each experiment, myotubes were continuously perfused with either control or agonist-containing Ringer solution. Multiple myotubes were measured per dish, and bulk gravity perfusion (∼10 ml) with control Ringer solution was used to wash the dish between each measurement. Peak intracellular Ca2+ changes in response to agonist application are expressed as ΔR (Ragonist − Rbaseline). At the beginning of each experiment, individual cells were electrically stimulated three times (8 V for 20 ms at 0.2 Hz for 15 s) with a stimulating electrode (an extracellular pipette filled with 200 mM NaCl) placed close to the cell of interest. Myotubes were then sequentially exposed to 60-s applications of different concentrations of caffeine (0.1, 0.3, 0.5, 0.7, 1.0, 3.0, 10, and 30 mM), each step followed by a 60-s wash with Ringer solution. Data were analyzed with FeliX (Photon Technology, Princeton, NJ) and SigmaPlot 8.0 (SPSS, Chicago, IL) software packages. Caffeine concentration-response curves were fitted according to an equation of the following general form: (1) where Dmax is the maximal response, [D] is the caffeine concentration, nH is the Hill coefficient, and EC50 is the concentration of caffeine at which the response is half-maximal.
Simultaneous measurements of voltage-gated Ca2+ currents and transients in whole cell patch-clamp experiments.
The whole cell patch-clamp technique was used to simultaneously measure L-type Ca2+ currents (L currents) and Ca2+ transients in expressing myotubes as described previously (4, 5). L currents were monitored from myotubes bathed in an external solution that contained (in mM) 145 tetraethylammonium (TEA)-Cl, 10 CaCl2, and 10 HEPES (pH 7.4 with TEA-OH). Patch pipettes were pulled from borosilicate glass (Harvard Apparatus), fire-polished (Microforge MF-830, Narishige), and had resistances of 1.8–3 MΩ when filled with an internal recording solution consisting of (in mM) 145 Cs-aspartate, 2 MgCl2, 10 HEPES, 0.1 Cs-EGTA, 2 Mg-ATP, and 0.2 mM pentapotassium fluo 4 (Molecular Probes, Eugene, OR) (pH 7.4 with CsOH). Depolarization-induced intracellular Ca2+ transients were monitored from a small rectangular region of the myotube with a photomultiplier detection system (Photon Technology International, South Brunswick, NJ). In some experiments, voltage-gated Ca2+ release was also monitored in the presence of a low concentration (2 mM) of extracellular caffeine (see Fig. 3).
L currents were elicited with 200-ms depolarizing steps from a holding potential of −80 mV to test potentials between −50 and +80 mV in 10-mV increments at 0.2 Hz. Each test pulse was preceded by a 1-s prepulse to −30 mV, followed by a 50-ms interval pulse to −50 mV to inactivate endogenous Na+ and T-type Ca2+ channels (1). Linear leak and capacitative currents were digitally subtracted with a P/4 prepulse protocol (2). Recordings were low-pass Bessel filtered at 2 kHz and sampled at 5 kHz. Ca2+ currents were normalized by linear cell capacitance (expressed in pA/pF). Immobilization-resistant intramembrane charge movements were measured after the prepulse protocol and blockade of ionic Ca2+ currents by the addition of 0.5 mM Cd2+ and 0.1 mM La3+ to the extracellular recording solution. The amount of immobilization-resistant charge movement was quantified 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) as described previously (2).
The L channel conductance (G) for each test voltage (V) was obtained with the following equation: (2) where ICa is the peak Ca2+ current during the test pulse, Vrev is the extrapolated reversal potential, and V is the membrane potential.
The voltage dependence of G, Qon, and intracellular Ca2+ release (ΔF/F) was fitted according to a Boltzmann distribution: (3) where Y is G, Qon, or ΔF/F; Ymax is Gmax, Qmax, or (ΔF/F)max; V1/2 is the potential at which Y = Ymax/2; and k is a slope factor.
All recordings were made at room temperature (∼23°C), and data are reported as means ± SE. Only currents with a maximal voltage error <5 mV due to series resistance were analyzed [average was 1.8 ± 0.2 (n = 45) and 0.98 ± 0.1 (n = 42) mV for α1S and R1086H, respectively] with Clampfit 8.0 (Axon Instruments, Foster City, CA) and SigmaPlot 8.0 (SPSS) software. Statistical significance (P < 0.05) was determined by unpaired Student’s t-test.
Differentiated GLT cultures were fixed and immunostained as previously described (19) with a mouse monoclonal anti-GFP antibody (Molecular Probes) at a dilution of 1:4,000 and the affinity-purified antibody 162 against RyR1 at a dilution of 1:5,000 (22). Alexa 488-conjugated secondary antibodies were used with the anti-GFP antibodies so that the antibody label and the intrinsic GFP signal were both recorded in the green channel. Alexa 594-conjugated antibodies were used in double-labeling experiments to achieve a wide separation of the excitation and emission bands. Controls, such as the omission of primary antibodies and incubation with inappropriate antibodies, were routinely performed. Images were recorded on a Zeiss Axiophot microscope with a cooled charge-coupled device camera and MetaView image-processing software (Universal Imaging, West Chester, PA). Quantitative analysis of the labeling patterns was performed by systematically screening coverslips of transfected myotubes with a ×63 objective. Labeling patterns of expressing multinuclear myotubes were classified as either “clustered” or “endoplasmic reticulum (ER)/SR staining.”
Ca2+ release channel sensitivity to activation by caffeine is modulated by Arg1086 of the α1S-subunit of skeletal muscle DHPR.
Increased sensitivity to activation by caffeine is used as a primary diagnostic measure for MH susceptibility. Moreover, a similar increase in caffeine sensitivity of Ca2+ release is also observed after expression of MH mutations in RyR1 in either HEK-293 cells (47) or skeletal myotubes derived from RyR1-knockout (dyspedic) mice (48). Therefore, we determined whether the sensitivity of SR Ca2+ release to activation by caffeine was also increased by the R1086H MH mutation in the α1S-subunit of the skeletal muscle DHPR (Fig. 1). For these experiments, dysgenic myotubes, which lack a functional α1S gene (8), were microinjected with cDNA for CD8 (an expression marker) and either wild-type α1S or R1086H. Functional expression of skeletal muscle DHPRs in injected myotubes was confirmed in each myotube by the presence of electrically evoked (8 V for 20 ms at 0.2 Hz for 15 s) Ca2+ transients, which are absent in noninjected dysgenic myotubes. This brief electrical train of three stimuli (Fig. 1A) elicited Ca2+ release events of similar magnitude and kinetics in both wild-type α1S- and R1086H-expressing myotubes. However, compared with noninjected and α1S-expressing myotubes, R1086H-expressing myotubes exhibited a significantly (P < 0.01) higher average resting indo 1 ratio [resting ratios were 0.62 ± 0.02 (n = 29), 0.63 ± 0.03 (n = 18), and 0.73 ± 0.02 (n = 25), for noninjected, α1S, and R1086H, respectively].
Immediately after the electrical train of stimuli, myotubes were exposed to 60-s sequential applications (Fig. 1A) of increasing concentrations of caffeine (0.1, 0.3, 0.5, 0.7, 1.0, 3.0, 10, and 30 mM), with each application followed by a 60-s wash with control Ringer solution. The data in Fig. 1 indicate that maximal caffeine-induced Ca2+ release is significantly (P < 0.01) reduced in R1086H-expressing myotubes. Specifically, maximal caffeine-induced changes in indo 1 fluorescence ratio were 0.56 ± 0.01 (n = 18) and 0.45 ± 0.03 (n = 25) for wild-type- and R1086H-expressing myotubes, respectively. Because the magnitude of caffeine-induced Ca2+ release is a measure of releasable SR Ca2+ load (7), the data in Fig. 1 indicate that expression of the R1086H MH mutation causes a modest reduction in RyR1-releasable SR Ca2+ content. In addition, similar to those observed previously (16), caffeine concentration-response relationships were very steep in any given wild-type α1S-expressing dysgenic myotube, presumably because of the presence of a strong component of Ca2+-induced Ca2+ release (CICR). Nevertheless, caffeine-induced Ca2+ release in α1S-expressing myotubes typically required concentrations of ≥1 mM (Fig. 1A, left). The threshold for caffeine-induced Ca2+ release was approximately fivefold lower (0.1–0.3 mM) in R1086H-expressing dysgenic myotubes (Fig. 1A, right). Interestingly, the sensitivity of caffeine-induced Ca2+ release was similar for uninjected dysgenic myotubes and R1086H-expressing myotubes (Fig. 1B).
Effects of R1086H mutation on L channel activity and voltage-gated Ca2+ release.
Skeletal muscle cells obtained from the porcine model of MH (homozygous for a R615C mutation in RyR1) exhibit a lower threshold for contraction (20) that arises from a selective hyperpolarizing shift in the voltage dependence of SR Ca2+ release that occurs in the absence of a significant change in the voltage dependence of L-type Ca2+ currents (9). We used the whole cell patch-clamp technique in conjunction with a Ca2+-sensitive dye to determine whether the R1086H MH mutation in α1S causes a similar increase in the sensitivity of the SR Ca2+ release mechanism to activation by voltage (Fig. 2). Expression of R1086H in dysgenic myotubes restored voltage-gated SR Ca2+ release that exhibited a maximal magnitude similar to that attributable to wild-type α1S. However, similar to that observed for myotubes obtained from MHS pigs, the threshold potential for Ca2+ release in R1086H-expressing myotubes was 5–10 mV more hyperpolarized than that of wild-type α1S-expressing myotubes (Fig. 2, A and B). However, in stark contrast to that of porcine MHS myotubes, the magnitude and voltage dependence of L channel conductance in R1086H-expressing myotubes were markedly reduced compared with those of wild-type α1S-expressing myotubes (Fig. 2C). Specifically, maximal L-channel conductance (Gmax) of R1086H-expressing myotubes was reduced 33%, the voltage for one-half activation of Gmax was shifted 6 mV to more positive potentials, and the extrapolated reversal potential was shifted to more negative potentials by ∼7 mV (Table 1).
Data presented in Figs. 1 and 2 indicate that the R1086H MH mutation in α1S enhances the sensitivity of the Ca2+ release mechanism to activation by both pharmacological (i.e., caffeine) and endogenous (voltage sensor) activators of RyR1. To test whether these two effects on release channel activation are additive in nature, we compared the magnitude and voltage dependence of Ca2+ release in the absence and presence of a low concentration (2 mM) of caffeine (Fig. 3). Representative L currents and voltage-gated Ca2+ transients from wild-type α1S- and R1086H-expressing myotubes in the presence of 2 mM caffeine are shown in Fig. 3A. Consistent with previous observations in normal myotubes (6), L-current density in both α1S- and R1086H-expressing myotubes was significantly reduced in the presence of near-threshold concentrations of caffeine (Table 1). As suggested previously, the mechanism for the caffeine-induced reduction in Gmax might reflect an increase in junctional Ca2+ that promotes Ca2+-dependent dephosphorylation of the DHPR and a subsequent reduction in the number of L channels available for activation (6). Interestingly, L-current density, but not the maximal magnitude of Ca2+ release [(ΔF/F)max], was further reduced in R1086H-expressing myotubes. Additionally, the threshold of voltage-gated Ca2+ release (as well as VF1/2) occurred at more hyperpolarized potentials (5–10 mV) in the presence of 2 mM caffeine (Fig. 3B and Table 1). Thus the R1086H MH mutation and threshold concentrations of caffeine produced additive effects on release channel sensitivity to activation by the voltage sensor.
Mechanism of reduced L-channel activity in R1086H-expressing myotubes.
We previously showed that reduced L-current density in dyspedic myotubes, which lack a functional RyR1 gene, arises from a decrease in both the Ca2+-conducting activity (40) and surface expression (2, 5) of the DHPR, as inferred from the maximal magnitude of immobilization-resistant intramembrane charge movement (Qmax). The influence of RyR1 on DHPR function is referred to as retrograde coupling (40). To determine whether the observed reduction in Gmax involves altered retrograde coupling between RyR1 and R1086H DHPRs, we compared L-channel current density and Qmax in wild-type α1S- and R1086H-expressing myotubes. For these experiments, L-current density was first measured in the presence of 10 mM extracellular Ca2+. Intramembrane charge movements (gating currents) were then measured after blockade of ionic Ca2+ currents by the extracellular addition of 0.5 mM Cd2+ and 0.1 mM La3+ (Fig. 4). Figure 4A illustrates representative gating currents elicited by 20-ms depolarizations to the indicated potentials, and Fig. 4B shows the average (±SE) voltage dependence of charge movement for dysgenic myotubes expressing either wild-type α1S or R1086H. Our results show that the voltage dependence of charge movement as well as Qmax is comparable in α1S- and R1086H-expressing myotubes [Qmax was 4.6 ± 0.5 (n = 15) and 3.9 ± 0.5 (n = 16) nC/μF for α1S and R1086H, respectively; P = 0.26, unpaired Student’s t-test]. Gmax values calculated from these same experiments were 119 ± 15.8 and 81 ± 13.7 nS/nF for wild-type α1S- and R1086H-expressing myotubes, respectively (P = 0.08). Gmax-to-Qmax ratios calculated from individual experiments were 26.2 ± 3.2 and 23.5 ± 3.3 nS/pC for wild-type α1S- and R1086H-expressing myotubes, respectively, and were not significantly different (P = 0.56).
Therefore, our data suggest that the decrease in Gmax for R1086H-expressing myotubes does not result from a selective reduction in either DHPR surface expression or aberrant retrograde coupling with RyR1, but more likely involves a general effect of the mutation on L-channel Ca2+ conductance. This is supported by investigations on the function and targeting of the analogous mutation (R1217H) engineered into the α1-subunit of the cardiac L channel (α1C). Cardiac DHPRs do not directly couple to RyRs (for review, see Ref. 10) and thus cannot support retrograde coupling from the RyR to the DHPR. Cardiac Ca2+ currents (Fig. 5A) and maximal L-channel conductance of GFP-α1C(R1217H)-expressing dysgenic myotubes were also significantly (P < 0.001) smaller than those attributable to wild-type α1C [Gmax was 383 ± 32 (n = 23) and 236 ± 25 (n = 24) nS/nF for wild-type α1C and R1217H, respectively]. Our observation that the R1217H mutation in α1C produced an effect on Gmax similar to that observed for the analogous mutation in α1S further supports the conclusion that the decrease in L-channel conductance observed in R1086H-expressing myotubes does not involve alterations in retrograde coupling with RyR1.
Similar to a recent report (28), α1C proteins expressed in dysgenic myotubes do not directly couple to RyR1 but nevertheless activate a small but significant SR Ca2+ release via a CICR mechanism. However, we found that neither the magnitude nor the voltage dependence of CICR attributable to α1C expression is altered by the R1217H mutation (Table 1). These data provide strong evidence that, although substitution of histamine for a highly conserved arginine residue in the intracellular III-IV linker imparts a similar effect on L-channel conductance mediated by both α1S and α1C, the mutation selectively sensitizes the voltage-gated release mechanism triggered by α1S.
Analogous to their skeletal muscle counterparts, both wild-type α1C and R1217H targeted primarily to clusters that overlapped with discrete RyR1 clusters (Fig. 5B). This colocalization indicates correct junctional targeting. This is particularly evident in the pseudocolor overlay images (Fig. 5B), in which green and red indicate α1 and RyR1, respectively, and yellow represents sites of colocalization of both channels. DHPR-RyR1 clusters were observed in 81% (n = 150) and 82% (n = 300) of myotubes transfected with α1C and R1217H, respectively. Comparable values of DHPR-RyR1 coclustering were found in myotubes transfected with α1S (81%; n = 150) and R1086H (78%; n = 450), thus confirming that DHPR junctional targeting is not affected by mutation of the conserved arginine residue in the DHPR III-IV loop.
MH is a pharmacogenetic disorder of skeletal muscle EC coupling characterized by increased release channel sensitivity to activation by triggering agents (e.g., halothane and caffeine) (27, 39). In many cases, MH is caused by point mutations/deletions in the gene encoding RyR1. Significant strides in understanding the pathogenic mechanisms of MH were initially made with a porcine model for MH that is caused by a single RyR1 point mutation (R615C). SR Ca2+ release channels isolated from skeletal muscle of MH pigs exhibit higher rates of CICR (14, 38), an increased sensitivity to activation by caffeine (18, 25, 45), halothane (42), 4-chloro-m-cresol (25, 45), and t-tubule depolarization (13, 29), and reduced inhibition by high concentrations of both Ca2+ (17) and Mg2+ (30). These abnormalities, which may be potentiated by inhalation anesthetics and depolarizing skeletal muscle relaxants, would be anticipated to result in hypersensitive or overactive SR Ca2+ release channels (11). In addition, in response to sarcolemmal depolarization, muscle fibers and myotubes obtained from the porcine model of MH exhibit lower thresholds for Ca2+ release (9) and contraction (20, 21). Thus the underlying pathogenesis of MH susceptibility is widely believed to arise from uncontrolled Ca2+ release that results from an overall increase in the sensitivity of the Ca2+ release mechanism to a broad range of activating stimuli (including caffeine, halothane, 4-chloro-m-cresol, Ca2+, and voltage sensors).
Consistent with this notion, our finding that voltage-gated Ca2+ release activates at more negative voltages in R1086H-expressing myotubes compared with that observed for α1S-expressing myotubes indicates that the R1086H mutation also enhances the sensitivity of the release mechanism to activation by the voltage sensor. This effect is similar to (but smaller than) that reported for porcine myotubes homozygous for the R615C MH mutation (9) and dyspedic myotubes expressing MH/CCD mutations in RyR1 (3). However, quantitative comparisons between our results and those in Ref. 9 should be made with caution because the measurements were made under very different conditions (native R615C RyR1 mutation in pig myotubes vs. transient expression of the R1086H MH mutant rabbit DHPR in dysgenic myotubes). Nevertheless, it is interesting to note that the shift in VF1/2 of R1086H-expressing myotubes is considerably less than that observed for the more severe MH/CCD mutants located in the NH2-terminal region of RyR1 (3). This observation supports the notion that mutations that result in certain forms of CCD may lead to a greater destabilization of the release channel closed state and an even greater shift in the voltage sensitivity of release compared with that produced by more benign, MH-selective mutations.
Our results revealed that the threshold for caffeine-induced Ca2+ release is approximately fivefold lower in R1086H-expressing myotubes compared with that of α1S-expressing dysgenic myotubes (Fig. 1). Surprisingly, we also found that the threshold for Ca2+ release in noninjected dysgenic myotubes also occurs at approximately fivefold lower caffeine concentrations compared with that of α1S-expressing dysgenic myotubes (Fig. 1B). Thus, given that the R1086H mutation increases release channel sensitivity to activation by both caffeine (Fig. 1) and voltage (Fig. 2), our data suggest that the intracellular III-IV linker of the DHPR acts as a negative regulatory module for release channel activation and that the R1086H MH mutation in α1S disrupts this critical negative allosteric regulatory mechanism.
We found that both resting indo 1 fluorescence and the sensitivity of caffeine-induced Ca2+ release are increased in R1086H-expressing myotubes. These results suggest that increased “sensitivity” of RyR1 to activation by caffeine may be a consequence of an elevated resting Ca2+ level in R1086H-expressing myotubes, rather than a reflection of an intrinsic change in SR Ca2+ release channel sensitivity, as suggested by others (33). In support of this idea, it was found that reducing resting Ca2+ levels in MHS myoballs with BAPTA-AM is sufficient to eliminate increased sensitivity to activation by caffeine and 4-chloro-m-cresol (34).
However, we also found that noninjected dysgenic myotubes also exhibit increased sensitivity of caffeine-induced Ca2+ release despite having resting Ca2+ levels identical to those of α1S-expressing myotubes. In addition, compared with wild-type α1S, voltage-gated Ca2+ release occurs at more negative potentials in R1086H-expressing myotubes under conditions in which intracellular Ca2+ levels were set to identical levels (via dialysis with a 60 nM free Ca2+ internal solution; Ref. 4). Thus increased RyR1 sensitivity to activation in R1086H-expressing myotubes most likely involves a combined effect of both an increase in resting Ca2+ and a disinhibition of a DHPR-mediated negative allosteric regulation of the release mechanism (16, 31).
Two different point mutations in Arg1086 of the intracellular III-IV loop of the skeletal muscle DHPR (R1086H and R1086C) are linked to MH (27, 39). Although the II-III loop is clearly an essential structural determinant of both orthograde (41, 46) and retrograde (24) DHPR-RyR1 coupling, a related functional role of the intracellular III-IV loop has not been suggested previously. Our results showing alterations in orthograde coupling (hyperpolarizing shift in VF1/2) created by an MH-causing mutation in α1S are the first to assign a functional role of the III-IV linker in DHPR-RyR1 coupling. Interestingly, the skeletal muscle DHPR II-III and III-IV loops have both been shown in biochemical experiments to bind to contiguous and/or overlapping regions of RyR1 (32). If such interactions were to influence orthograde coupling with RyR1, then our results could either be explained by a direct effect of the R1086H mutation on this interaction or by a more indirect effect such as the mutated III-IV linker altering the structural conformation and activity of the II-III loop. Thus it will be important for future studies to determine whether the R1086C MH mutation in α1S (as well as other MH mutations in the DHPR identified in future genetic studies) imparts a similar effect on RyR1 sensitivity to activation by the voltage sensor.
How could an important role of the III-IV loop in orthograde coupling have evaded detection in previous studies? One possibility is that the determination of DHPR regions critical for DHPR-RyR1 coupling in intact muscle cells has primarily been inferred from experiments expressing α1S and α1C chimeras in dysgenic myotubes (24, 41, 46). Because the amino acid sequence of the III-IV loop is highly conserved (85% identity) between skeletal and cardiac muscle DHPR α1-subunits, chimeric skeletal-cardiac III-IV loop chimeras may fail to reveal important functional domains that are conserved in both proteins. In this way, functional analysis of DHPR disease mutations reconstituted in dysgenic myotubes can potentially unveil new insights into the structure/function of skeletal muscle EC coupling that may have eluded detection with the chimeric approach.
Our experiments are the first to determine effects of a MH disease mutation in the skeletal muscle DHPR (R1086H) on the orthograde and retrograde signals of skeletal muscle EC coupling. Similar to that observed for MH mutations in RyR1, the R1086H MH mutation in the α1S-subunit of the skeletal muscle DHPR increases the sensitivity of the Ca2+ release mechanism to activation by both pharmacological (caffeine) and endogenous (voltage sensor) activators. As similar effects have also been documented for MH mutations in RyR1 (3, 9, 47, 48), a generalized increase in sensitivity of the Ca2+ release mechanism to activation by a wide range of triggering agents may indeed represent a unifying principle that underlies increased susceptibility of MH muscle to activation by halothane during anesthesia.
This work was supported in part by the National Institute of Arthritis and Musculoskeletal and Skin Diseases (Grants AR-44657 to R. T. Dirksen and AR-46513 to P. D. Allen), a Neuromuscular Disease Research grant from the Muscular Dystrophy Association (to R. T. Dirksen), the Austrian National Bank (to M. Grabner), the Fonds zur Förderung der Wissenschaftlichen Forschung, Austria (Grants P13831-GEN and P16098-B11 to M. Grabner and P16532-B05 to B. E. Flucher), and European Union Grant HPRN-CT-2002-00331 to B. E. Flucher.
We thank Linda Groom and Dr. J. Hoflacher for excellent technical assistance, Dr. N. Kasielke for valuable contributions to the cardiac DHPR recordings, and Dr. H. Glossmann for continuous support. This work is part of the Ph.D. thesis of R. G. Weiss.
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- Copyright © 2004 the American Physiological Society