Departments of 1Biochemical Pharmacology and 3Physiology, Innsbruck Medical University, A-6020 Innsbruck, Austria; 2Department of Pharmacology and Physiology, University of Rochester Medical Center, Rochester, NY 14642; and 4Department of Anesthesia Research, Brigham and Womens Hospital, Boston, Massachusetts 02115
Submitted 1 April 2004 ; accepted in final form 8 June 2004
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ABSTRACT |
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excitation-contraction coupling; calcium channel; muscle disease
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 1S-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 1S-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
1S. 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.
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MATERIALS AND METHODS |
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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 (0.130 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
R (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:
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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 1.83 M 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).
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The L channel conductance (G) for each test voltage (V) was obtained with the following equation:
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The voltage dependence of G, Qon, and intracellular Ca2+ release (F/F) was fitted according to a Boltzmann distribution:
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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
1S 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 Students t-test.
Immunofluorescence labeling. 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 x63 objective. Labeling patterns of expressing multinuclear myotubes were classified as either "clustered" or "endoplasmic reticulum (ER)/SR staining."
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RESULTS |
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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 1S-expressing dysgenic myotube, presumably because of the presence of a strong component of Ca2+-induced Ca2+ release (CICR). Nevertheless, caffeine-induced Ca2+ release in
1S-expressing myotubes typically required concentrations of
1 mM (Fig. 1A, left). The threshold for caffeine-induced Ca2+ release was approximately fivefold lower (0.10.3 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 1S 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
1S. However, similar to that observed for myotubes obtained from MHS pigs, the threshold potential for Ca2+ release in R1086H-expressing myotubes was 510 mV more hyperpolarized than that of wild-type
1S-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
1S-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).
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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 1S- 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
1S or R1086H. Our results show that the voltage dependence of charge movement as well as Qmax is comparable in
1S- and R1086H-expressing myotubes [Qmax was 4.6 ± 0.5 (n = 15) and 3.9 ± 0.5 (n = 16) nC/µF for
1S and R1086H, respectively; P = 0.26, unpaired Students t-test]. Gmax values calculated from these same experiments were 119 ± 15.8 and 81 ± 13.7 nS/nF for wild-type
1S- 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
1S- 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 1-subunit of the cardiac L channel (
1C). 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-
1C(R1217H)-expressing dysgenic myotubes were also significantly (P < 0.001) smaller than those attributable to wild-type
1C [Gmax was 383 ± 32 (n = 23) and 236 ± 25 (n = 24) nS/nF for wild-type
1C and R1217H, respectively]. Our observation that the R1217H mutation in
1C produced an effect on Gmax similar to that observed for the analogous mutation in
1S 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), 1C 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
1C 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
1S and
1C, the mutation selectively sensitizes the voltage-gated release mechanism triggered by
1S.
Analogous to their skeletal muscle counterparts, both wild-type 1C 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
1 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
1C and R1217H, respectively. Comparable values of DHPR-RyR1 coclustering were found in myotubes transfected with
1S (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.
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DISCUSSION |
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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 1S-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 1S-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
1S-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
1S 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 1S-expressing myotubes. In addition, compared with wild-type
1S, 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 1S 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
1S (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 1S and
1C 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
1-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 1S-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.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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