Functional and structural characterization of a novel malignant hyperthermia-susceptible variant of DHPR-β_1a subunit (CACNB1)

Abstract

Malignant hyperthermia (MH) susceptibility has been recently linked to a novel variant of β_1a subunit of the dihydropyridine receptor (DHPR), a channel essential for Ca²⁺ regulation in skeletal muscle. Here we evaluate the effect of the mutant variant V156A on the structure/function of DHPR β_1a subunit and assess its role on Ca²⁺ metabolism of cultured myotubes. Using differential scanning fluorimetry, we show that mutation V156A causes a significant reduction in thermal stability of the Src homology 3/guanylate kinase core domain of β_1a subunit. Expression of the variant subunit in β₁-null mouse myotubes resulted in increased sensitivity to caffeine stimulation. Whole cell patch-clamp analysis of β_1a-V156A-expressing myotubes revealed a −2 mV shift in voltage dependence of channel activation, but no changes in Ca²⁺ conductance, current kinetics, or sarcoplasmic reticulum Ca²⁺ load were observed. Measurement of resting free Ca²⁺ and Na⁺ concentrations shows that both cations were significantly elevated in β_1a-V156A-expressing myotubes and that these changes were linked to increased rates of plasmalemmal Ca²⁺ entry through Na⁺/Ca²⁺ exchanger and/or transient receptor potential canonical channels. Overall, our data show that mutant variant V156A results in instability of protein subdomains of β_1a subunit leading to a phenotype of Ca²⁺ dysregulation that partly resembles that of other MH-linked mutations of DHPR α_1S subunit. These data prove that homozygous expression of variant β_1a-V156A has the potential to be a pathological variant, although it may require other gene defects to cause a full MH phenotype.

INTRODUCTION

Malignant hyperthermia (MH) is a genetic disorder of skeletal muscle that confers abnormal sensitivity to certain anesthetic compounds, such as volatile hydrocarbon anesthetic gases (e.g., halothane) depolarizing neuromuscular blockers (e.g., succinylcholine) and rarely, in humans, to stressors such as vigorous exercise and heat (26, 34). In patients susceptible to MH, exposure to these agents triggers a series of events characterized by tachycardia, elevated core temperature, respiratory and metabolic acidosis, muscle rigidity, and in the absence of any intervention, potential death (34, 36). Although the pathophysiology of this syndrome is not yet fully understood, it is well accepted that the underlying causes of MH are alterations in the skeletal muscle calcium metabolism (32–34). Molecular genetic linkage studies have mapped the primary locus associated with the syndrome to chromosome 19q12-13.2, the gene encoding the skeletal ryanodine receptor (RyR1), a calcium release channel localized within skeletal muscle junctional sarcoplasmic reticulum (SR) key for muscle contraction (40).

A second locus linked to MH syndrome was mapped to chromosome 1q32.1, which includes the CACNA1S gene that encodes the α_1S subunit of the skeletal muscle dihydropyridine receptor (DHPR), the channel-forming unit of the voltage-dependent L-type Ca²⁺ channel. Currently, more than 40 sequence variants of the DHPR α_1S gene have been identified in humans with a family history of MH susceptibility (10, 22, 35). However, only three of these mutation-causing variants (R174W, R1086H, and T1354S) have been functionally characterized in vitro and confirmed to disrupt Ca²⁺ metabolism in cultured myogenic cells.

Currently, there are more than 300 variants of unknown significance linked to MH susceptibility in RyR1 and DHPR α_1S and only a handful of them have been shown to be causative of the disease. Moreover, in most of these cases, it is unknown whether these mutations appear alone or in combination with yet unidentified variants in other Ca²⁺ regulatory genes. In a recent genetic study, targeted to MH-susceptible individuals who have no variant in RyR1 or the DHPR α_1S subunit, several additional mutant variants were identified in other genes involved in Ca²⁺ regulation in skeletal muscle. These include the DHPR auxiliary subunit β_1a, the SR Ca²⁺ pump (SERCA1), and the Ca²⁺ binding proteins calsequestrin-1 (CASQ1), thus identifying potential additional Ca²⁺ signaling pathways involved in MH (8). The auxiliary β_1a subunit is an essential regulator of the DHPR α_1S subunit and a reported modulator of RyR-1 function (6, 17, 23, 24, 28, 39), two major proteins linked to MH pathology. However, whether MH susceptibility of the individual bearing the DHPR β_1a variant is a direct result of the amino acid mutation introduced by this variant (V156A) is currently unknown. Therefore, an understanding of the role of this novel variant on DHPR and RyR1 channel function and, more importantly, its direct effect on myoplasmic Ca²⁺ regulation could provide important new insights into the functional and genetic complexity of MH syndrome.

To study the role of DHPR β_1a variants in the pathophysiology of MH, here we expressed and functionally characterized in vitro the effect of mutation V156A on both β_1a protein stability and Ca²⁺ regulation of cultured myotubes. Our data indicate that structurally, β_1a-V156A leads to a reduction in the stability of the Src homology 3 (SH3)/guanylate kinase (GK) core domain of the β_1a protein and under basal conditions this novel mutation affects the cationic permeability of the plasma membrane in a process dependent on Na⁺/Ca²⁺ exchanger (NCX) and/or transient receptor potential canonical (TRPC) channels function, leading to chronic elevation of myogenic resting free Ca²⁺.

MATERIALS AND METHODS

DHPR β_1a expression and purification for in vitro studies.

Genes encoding hexa-histidine-tagged β_1a-constructs were synthesized and cloned into a custom vector pJ411^KanR with a high copy number origin of replication (DNA 2.0). Truncated β_1a constructs were constructed to include the following core residues: β_1a(68–192 ~252–462) and a comparable construct for V156A (with appropriate codon replacement). Bacteria (Escherichia coli BL21 strain) transformed with the plasmids were cultured at 37°C in 2× YT and 50 μg/ml kanamycin until OD₆₀₀ ~0.3. They were further cultured at 16°C for 1 h. Protein expression was induced with IPTG (0.4 mM), and the cells were cultured overnight. The bacterial pellet was resuspended in 50 mM phosphate buffer at pH 7.0, 500 mM NaCl and 30 mM imidazole (buffer A) containing an EDTA-free protease inhibitor cocktail (Roche), lysozyme (1 μg/ml), DNaseI (1 μg/ml), and RNaseH (1 μg/ml). Cells were lysed using a French Press (3 × 1,500 psi), and the cleared lysate was then applied to a HisTrap column. The resin was washed with buffer A followed by buffer A containing 1.5 M NaCl. The protein was eluted using a gradient of imidazole up to 500 mM. The hexahistidine tag was cleaved with HRV 3C protease (produced in-house) and simultaneously dialyzed overnight in buffer A containing 5 mM 2-mercaptoethanol at 4°C. The protein solution was applied to Ni-NTA resin to remove the hexahistidine tag-containing entities. The flow-through was further purified by size-exclusion chromatography.

Peptide synthesis.

A high-affinity binder of β_1a, DHPR α_1S AID peptide (³⁵⁷QQLEEDLRGYMSWITQGE³⁷⁴) was synthesized by the Biomolecular Resource Facility of the John Curtin School of Medical Research (Australian National University, Canberra, Australia) using an Applied Biosystems 430A peptide synthesizer and purified by reverse phase high-performance liquid chromatography (RP-HPLC) on a Jupiter 300 C4 column. The peptide was eluted using a linear gradient from buffer A (deionized water and 0.1% trifluoroacetic acid, TFA) and buffer B (acetonitrile and 0.1% TFA). Purified peptide fractions were identified by mass spectroscopy using an AB MDS Sciex 4800 MALDI-TOF-TOF Mass Analyzer.

Measurement of protein stability.

Protein stability was measured using differential scanning fluorimetry (DSF), which was undertaken at the Commonwealth Scientific and Industrial Research Organisation (CSIRO)-C3 Center (Melbourne, Australia). The protocol uses sypro-orange as reporter dye and involves screening protein samples against 28 buffer/salt formulations. Each data set is made up of quadruplicate measurements, and two separate experiments were performed.

cDNA constructs and virus packaging for myotube studies.

Cloning and expression of the full-length cDNA of mouse DHPR β_1a-subunit (GenBank NM_031173) into the bicistronic retroviral vector pCMMP-MCS-IRES-Puro were described previously (17). Point mutation V156A of human β_1a-subunit (GeneBank, NM_199247.2) was introduced into the homologous residue (V156) of the mouse gene using Gibson assembly (New England Biolabs) and double-stranded DNA fragments (gBlocks, Integrated DNA Technologies). The presence of mutation and proper reading frame of all selected clones were confirmed by sequencing before use. Virus packaging was performed with a set of three packaging vectors as described elsewhere (17).

Cell culture and calcium imaging.

Primary myoblasts from DHPR β₁-null mouse muscles (7, 23) were infected with virion particles carrying β_1a gene at a multiplicity of infection (MOI) = 0.5 and then selected with 1.6 µg/ml Puromycin for 2 wk to obtain stably transduced myoblasts. Cells expressing mouse Wt-β_1a or β_1a-V156A subunit were then grown and differentiated in 96-well plates as described previously (18). Calcium imaging was performed 5–6 days after differentiation in myotubes loaded with 5 μM Fura-2-AM or 2 μM Fluo-4 (Molecular Probes) in imaging buffer (125 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1.2 mM MgSO₄, 6 mM glucose, and 25 mM HEPES-Tris, pH 7.4). Cell membrane depolarization was performed by 5 s of perfusion with 5–7 volumes of imaging buffer containing increasing concentrations of KCl. To preserve osmolarity of the depolarization buffer, the increased K⁺ concentration was compensated with an equivalent reduction in total NaCl concentration. Sarcoplasmic reticulum Ca²⁺ content of Fura-2-AM-loaded cultured myotubes was estimated from the peak fluorescent amplitude of the Ca²⁺ transient induced by 40 mM caffeine stimulation in the presence of 1 μM thapsigargin (Tg) and in the absence of extracellular Ca²⁺ (11, 19). Extracellular Ca²⁺ entry at rest was estimated from the rate of dye quench by Mn²⁺ entry in myotubes loaded with 5 μM Fura-2-AM in the presence and in the absence of 10 µM KB-R7943 (glycerol, 2-(2-(4-(4-nitrobenzyloxy) phenyl)ethyl)-isothiourea methane sulfonate), as described previously (16, 20). Cells were then imaged at 360 nm (±5 nm) with an intensified 10-bit digital CCD camera (XR-Mega-10, Stanford Photonics, Stanford, CA) using a DG4 multiwavelength light source. Fura-2 images were acquired with a ×40 objective (Olympus, UApo/340), and fluorescent emission at 510 nm was then captured from regions of interest within each myotube at 33 frames/s using Piper-controlled acquisition software (Stanford Photonics) and are expressed as the ratio of signal collected at alternating 340 nm/380 nm excitation wavelengths. For Mn²⁺ quench studies, images were acquired at 510 nm as the average of 5 individual recordings captured at 6 frames/s. Fluo-4 fluorescent images were acquired using a ×60 objective, and emission at 510 nm was then collected from regions of interest from individual cells excited at 490 nm and are expressed as arbitrary fluorescent units.

Cell homogenate preparation and Immunoblotting.

Whole cell homogenates were prepared from myoblasts stably transduced with Wt-β_1a or β_1a-V156A subunit. Cells were differentiated in a six-well plate format for 5 days followed by harvesting and suspension in solubilization buffer (250 mM sucrose and 10 mM HEPES, pH 7.4, 1% Triton X-100) supplemented with 1 mM EDTA, 10 μg/ml leupeptin, 0.7 μg/ml pepstatin A, 5 μg/ml aprotinin, and 0.1 mM Pefabloc SC. Homogenates were centrifuged at 12,000 g for 20 min, and the supernatants were then separated in SDS-polyacrylamide gel electrophoresis (30) and transferred to PVDF membrane. Membranes were immunoblotted with anti-β₁ monoclonal antibody (N7/18, NeuroMab, Davis, CA) followed by incubation with horseradish peroxidase (HRP)-conjugated secondary antibody. To correct for variations in protein expression among preparations as well as differences in protein loading between lanes control membranes were incubated with polyclonal anti-β-actin antibody HRP-conjugated (MA5-15739-HRP, ThermoFisher). Blots were finally developed with SuperSignal ultra chemiluminescent substrate (Pierce). Net band intensities of unsaturated blot signals were corrected to β-actin expression and are plotted as fraction of the Wt signal (β_x/β_1a).

Measurement of ionic currents.

Macroscopic Ca²⁺ currents were measured using whole cell patch clamp as described previously (14, 17). Myotubes were voltage clamped with the use of an Axopatch 200B amplifier (Axon Instruments, Foster City, CA). Effective series resistance was compensated up to the point of amplifier oscillation with the Axopatch circuit, and the leak current was subtracted by using a –P/6 protocol before each sweep. The patch pipettes were coated using Sylgard and had a resistance of ~2 MΩ when filled with the pipette solution. The patch pipette internal solution consisted of 140 mM Cs-Aspartate, 5 mM MgCl₂, 10 mM Cs-EGTA, and 10 mM HEPES titrated with CsOH to pH 7.2. The composition of the external bath solution was 145 mM TEA-Cl, 10 mM CaCl₂, 10 mM HEPES titrated with TEA(OH) to pH 7.4.

The voltage dependence of the Ca²⁺ currents was fitted to the following equation.

$$I=G_{\max }(V-V_{\text{r}})/\{1+\exp [(V_{1/2}-V)/k]\}$$

where G_max is maximal conductance, V corresponds to the test potential, V_1/2 is the potential at which G = 1/2 G_max, k represents a slope parameter, and V_r is reversion potential.

Resting free Ca²⁺ and Na⁺ measurements.

Resting [Ca²⁺] and [Na⁺] measurements were performed with double-barreled Ca²⁺- or Na⁺-selective microelectrodes on myotubes differentiated in ECM-coated 35-mm culture dishes as described previously (1, 2). Briefly, potentials from the 3 M KCl (V_m), Ca²⁺ (V_Cae), and Na⁺ (V_Nae) barrels were acquired with high-impedance amplifiers (model FD-223; WPI, Sarasota, FL) and then V_m was subtracted electronically from the V_Cae and V_Nae to give a differential signal (V_Ca and V_Na) that represented the resting myoplasmic Ca²⁺ and Na⁺ concentrations, respectively. Records of V_m, V_Ca, and V_Na potentials were filtered at 200 Hz (WPI-LPF-30) and acquired using AxoGraph software (version 4.8; Axon Instruments) for future analysis.

Data collection and statistical analysis.

The Fura-2 fluorescent Ca²⁺ release signal is expressed as average peak transient amplitude (F − F_o). The Fluo-4 fluorescent signal is expressed as area under the curve, corresponding to the integral of the total transient signal (Sum of F − F_o), that better reflects the overall magnitude of Ca²⁺ release. The Fluo-4 signal was then normalized to F_max (20 mM caffeine). Unless otherwise indicated, statistical significant differences among data sets were calculated by one-way ANOVA analysis (Tukey’s test) using Prism-7 software (GraphPad, San Diego, CA). Data are expressed as means ± SE.

RESULTS

The c.467T>C variant of CACNB1 (DHPR β_1a-subunit) was recently identified in a MH-susceptible individual with altered in vitro contracture test (IVCT) for caffeine and halothane stimulation (22). This variant affects a highly conserved locus, thus suggesting a region of functional importance (8), and results in the mutation of a conserved Val residue (V156A mutation) of the Src homology 3 (SH3) domain of β_1a subunit (Fig. 1). SH3 domains are known mediators of protein-protein interactions (27, 41) and have been shown to be critical for the voltage-sensing properties of the DHPR complex (12). The effects of this mutant were examined in the context of protein stability, and the effects of β_1a-V156A mutation on Ca²⁺ homeostasis were characterized using cultured mouse β_1a-null myotubes.

Fig. 1.

Structure and amino acid sequence alignment of β_1a. Schematic organization of the β_1a subunit highlighting the location of conserved (SH3, HOOK, GK) and variable domains. The sequence alignment compares the SH3 domain from several vertebrate species showing the conservation of the amino acid sequence harboring residue Val-156 (black box) among different β isoforms. GeneBank accession numbers: human (AAB58780); mouse (NP_112450); rabbit (P19517); rat (EDM05897); zebrafish (AY952462); mouse β_2a (NP_075605); mouse β₃ (NP_036960); rat β₄ (NP_001099203).

V156A mutation reduces the stability of the β_1a-core subunit.

We have previously employed differential scanning fluorimetry (DSF) as a measure of protein stability (37). For this study, we wished to evaluate how substituting V156 with an alanine residue would impact upon β_1a protein stability. Because of the well-behaved properties of the SH3/GK core domains of β_1a, stability comparisons were performed for this construct. Both the wild-type (Wt) and mutant constructs gave rise to melting profiles that typified a stable globular protein (Fig. 2, A and B). Comparison of the Wt and mutant melting temperature (T_m) over a range of diverse pH, salt, and buffer conditions (Fig. 2, C and D) revealed a statistically significant ~2.0°C reduction (P = 0.019, ANOVA single factor) in protein stability upon the introduction of the mutation. This reduction in protein stability was observed under all conditions examined, indicating that external variables such as pH or salt concentration had little bearing on the difference in T_m. Addition of the high-affinity AID peptide (DHPR α_1S interacting domain) resulted in an overall stabilization for both the Wt and V156A mutant ranging from 2.5 to 8°C; however, the overall T_m difference between the constructs in the presence of AID was in the order of ~0.5–1.0°C (P = 0.066).

Fig. 2.

Thermal denaturation measurements of purified Wt- and β_1a-V156A cores. A: melting curves for Wt-β_1a in Tris·HCl, pH 8.0, at 50 mM (blue) and 200 mM (orange) NaCl, respectively. B: melting curves for β_1a-V156A in Tris·HCl, pH 8.0, at 50 mM (blue) and 200 mM (orange) NaCl, respectively. C: summary of transition melting temperatures for Wt and V156A constructs in various buffers in 200 mM NaCl. D: same as in C but in the presence of two molar excess AID peptide.

MH-susceptible variant β_1a-V156A enhances sensitivity to caffeine activation in cultured myotubes.

Increased sensitivity to caffeine activation has been one of the primary readouts for MH susceptibility linked to both RyR1 (13, 21, 22, 31) and skeletal DHPR (10, 22). Therefore, to assess the effect of the β_1a-V156A mutation on caffeine sensitivity, here we stably transduced DHPR β₁-null myoblasts with either DHPR Wt-β_1a or β_1a-V156A subunit using retroviral infection followed by puromycin selection and further differentiation into myotubes. Western blot analysis with anti-β_1a antibody of whole cell homogenates from cultured myotubes (Fig. 3A) indicated that both cell genotypes expressed β_1a protein at nearly identical levels (P > 0.05). Figure 3B shows average caffeine dose responses of Fura-2-loaded myotubes exposed to increasing concentrations of caffeine for each cell phenotype. These data show that, compared with myotubes expressing Wt-β_1a subunit, the β_1a-V156A-expressing myotubes displayed a slight but statistically significant (P < 0.05) leftward shift in caffeine sensitivity, with average EC₅₀ of 5.9 ± 0.15 mM (n = 82) and 5.1 ± 0.19 mM (n = 77) for Wt-β_1a and β_1a-V156A myotubes, respectively. To further verify this finding, we also measured caffeine-induced calcium release in myotubes loaded with Fluo-4, a Ca²⁺ dye that presents a quantum yield superior to that of Fura-2 and, therefore, produces larger increase in fluorescence intensity in response to Ca²⁺ binding. Analysis of normalized Ca²⁺ release signal (area under the curve) further confirmed that expression of β_1a-V156A conferred myotubes with enhanced sensitivity to caffeine activation (Fig. 3C), with measured average EC₅₀ of 6.5 ± 0.18 mM (n = 76) and 5.2 ± 0.20 mM (n = 68) for the Wt- and β_1a-V156A myotubes (P < 0.05), respectively.

Fig. 3.

MH-susceptible variant β_1a-V156A modulates caffeine sensitivity of SR Ca²⁺ release in cultured myotubes. A: representative immunoblots of whole cell homogenates from Wt-β_1a (Wt) and β_1a-V156A (V156A) expressing myotubes. Gels were loaded with 20 μg total protein/lane and immunoblotted for expression of β_1a. Band intensity was corrected to loading differences (β-actin signal) and is expressed as fraction of the Wt-β_1a signal (means ± SD). Numbers in the bars indicate the total number of lanes analyzed from 4 immunoblots of 2 independent myotubes preparations (P > 0.05). β₁-Null myotubes stably transfected with Wt-β_1a (black) or β_1a-V156A (gray) were loaded with Fura-2 or Fluo-4 and then challenged with increased concentrations of caffeine. B and C: average caffeine-dose response relationship for myotubes loaded with either Fura-2 (B) or Fluo-4 (C) fitted to a sigmoidal dose-response curve. Data are presented as means ± SE; n = 3 (B) and n = 4 (C) independent experiments.

Previous studies on myotubes bearing MH mutant R174W of the DHPR α_1S subunit revealed that in addition to increased caffeine sensitivity, this mutation caused significant alteration in total SR calcium load of cultured cells (14). To quantify the effect of β_1a-V156A mutant variant on SR calcium content, we estimated total Ca²⁺ release by depleting SR stores of Fura-2-loaded myotubes with 40 mM caffeine in the presence of 1 μM thapsigargin (Tg) plus Cd²⁺ and La³⁺. These experimental conditions selectively target SR Ca²⁺ store, preventing signal distortion by Ca²⁺ reloading into the SR stores or Ca²⁺ entry from the extracellular medium. Figure 4A shows that under these conditions, Wt-β_1a and β_1a-V156A-expressing myotubes had biphasic responses to caffeine, with a fast Ca²⁺ release component followed by a slower and sustained release event. Quantitative analysis of the calcium signal indicates that both cell phenotypes have peak Ca²⁺ amplitude and total Ca²⁺ released (area under the curve of a 2-min interval following caffeine addition) that were indistinguishable from one another, suggesting no difference in overall SR calcium content (Fig. 4B).

Fig. 4.

β_1a-V156A mutation does not affect SR Ca²⁺ load of cultured myotubes. A: representative traces of caffeine-induced Ca²⁺ release transients of Fura-2-loaded myotubes expressing Wt-β_1a (black) or β_1a-V156A mutation (gray). The buffer was supplemented with 1 μM thapsigargin (Tg) and 0.5 mM Cd²⁺/0.1 mM La³⁺ to prevent Ca²⁺ reload to the SR stores and extracellular Ca²⁺ entry, respectively. B: comparison of average peak Ca²⁺ transient amplitude (ΔF) and overall Ca²⁺ released (area under the curve) of β₁-null myotubes expressing either Wt-β_1a (black) or β_1a-V156A (gray) mutant subunit. Data are presented as means ± SE. Numbers in the bars indicate the total number of cells analyzed per condition; n = 3 independent experiments. a.u., Arbitrary fluorescent units.

Effect of β_1a-V156A mutation on L-type Ca²⁺ channel function.

Previous studies of MH-linked mutations of DHPR α_1S subunit revealed significant effects of these mutations on the L-type Ca²⁺ channel function (3, 14, 38, 42). Here we evaluate the role of β_1a-V156A on DHPR function by analyzing the effect of the β_1a-V156A mutation on excitation-contraction (E-C) coupling, L-type Ca²⁺ current density, and current kinetics. Figure 5, A and B, shows representative KCl dose-response curves of Fura-2-AM-loaded for myotubes expressing either Wt- or β_1a-V156A mutation. Analysis of these data indicated that both cell phenotypes have similar sensitivity to K⁺ depolarization, with average EC₅₀ of 18.7 ± 0.5 mM (n = 90) and 20.2 ± 0.7 mM (n = 76) for Wt- and β_1a-V156A-expressing cells, respectively (P > 0.05). Figure 4A suggests that at high K⁺ concentrations, β_1a-V156A myotubes displayed seemingly smaller average peak Ca²⁺ transient amplitude than wild type cells. However, further analysis of Ca²⁺ transients induced by 60 mM KCl, in the presence of either extracellular Ca²⁺ or calcium channel blockers Cd²⁺ and La³⁺, found no statistical differences in peak Ca²⁺ amplitude of K⁺-induced Ca²⁺ transients between Wt-β_1a and β_1a-V156A myotubes (Fig. 5, B and C). Overall, these data suggest that β_1a-V156A mutation had no noticeable effects on the sensitivity of SR Ca²⁺ release to KCl-induced membrane depolarization, thus E-C coupling.

Fig. 5.

Depolarization-induced Ca²⁺ release is not affected by expression of β_1a-V156A subunit. A: representative fluorescent traces of K⁺-dose responses from Wt- and V156-expressing myotubes. B: average peak fluorescent amplitude of K⁺-induced Ca²⁺ release transients observed in Fura-2 loaded myotubes expressing Wt-β_1a (black) and β_1a-V156A subunit (gray). Data are expressed as means ± SE and were fitted to a sigmoidal dose-response curve. C: comparison of average peak Ca²⁺ transient amplitude induced by 60 mM K⁺ in the presence (2 mM Ca²⁺) and in the absence of extracellular Ca²⁺ (Cd²⁺/La³⁺) shows no differences in E-C coupling signaling between Wt-β_1a (black) and β_1a-V156A (gray) expressing myotubes. Data are presented as means ± SE. Numbers in the bars indicate the total number of cells analyzed per condition; n = 3 independent experiments. r.u. = 340/380 rate units.

The effects of β_1a-V156A mutation on DHPR channel function were further analyzed using whole cell patch-clamp technique. Comparison of Ca²⁺ current recording shows that Wt-β_1a and β_1a-V156A-expressing myotubes display classic slow-activating Ca²⁺ currents (Fig. 6A). Further analysis revealed that β_1a-V156A-expressing myotubes displayed Ca²⁺ currents that were indistinguishable from Wt-β_1a myotubes in peak amplitude (Fig. 6, A and B), magnitude of L-type conductance (G_max), or current kinetics (Fig. 6, C and D). However, average voltage dependence of the channel activation (V_1/2) in β_1a-V156A-expressing myotubes was shifted by ~2 mV to more negative potential (Fig. 6B and Table 1, P < 0.05). The similarity in Ca²⁺ current densities between Wt- and the V156A-cell line (Fig. 6, A and B) also suggests that equivalent levels of DHPR complex were expressed at the plasma membrane of both genotypes and, therefore, β_1a-V156A variant did not selectively affect expression or targeting of the DHPR complex.

Fig. 6.

Expression of the β_1a-V156A subunit has limited effect on L-type channel function of cultured myotubes. A: representative Ca²⁺ currents traces recorded from β₁-null myotubes expressing either Wt-β_1a (black) and β_1a-V156A (gray). Scale bars represent 20 pA/pF and 50 ms. B: comparison of current-voltage (I–V) relationship. Myotubes were depolarized for 200 ms from a holding potential of −50 mV to test potentials between −40 mV and +90 mV applied in 10 mV increments. C: current kinetics of Wt-β_1a and β_1a-V156A expressing myotubes. Currents were fitted to a bi-exponential curve, and the corresponding τ slow and τ fast values at +30 mV were plotted as means ± SE. Numbers in the bars indicate the number of cells analyzed per condition; n = 3 independent experiments. D: average contribution of τ slow and τ fast to the total current. Values are expressed as fraction of the total amplitude. The I–V relationships were fitted using Eq. 1 (see materials and methods). Fitting parameters for each plot between −60 mV and +90 mV are presented in Table 1. Data are expressed as means ± SE; n = 3 independent experiments.

Table 1.

Comparison of Ca²⁺ conductance parameters of myotubes expressing Wt- and β_1a–V156A subunit

	G–V Curves
	Gmax, pS/pF	V1/2, mV	k, mV
Wt-β1a	501 ± 28 (20)	17.4 ± 0.8 (20)	6.3 ± 0.1 (20)
β1a-V156A	505 ± 33 (27)	15.2 ± 0.5* (27)	6.4 ± 0.2 (27)

Data are means ± SE of Boltzmann parameters fitted between −50 mV and +90 mV. References shown in parentheses.

^*P < 0.05, compared with Wt-β_1a (unpaired t-test).

β_1a-V156A-expressing myotubes display elevated [Ca²⁺]_i and [Na⁺]_i at rest.

MH-linked mutations of RyR1 and the DHPR α_1S subunit have often been associated with elevated myoplasmic resting free [Ca²⁺]. To evaluate whether β_1a-V156A mutation bears this distinctive phenotype, we directly measured [Ca²⁺]_i of Wt-β_1a and β_1a-V156A-expressing myotubes using calcium-selective microelectrodes. Average data show that [Ca²⁺]_i in Wt-β_1a-expressing myotubes was 140 ± 2 nM (n = 60), a concentration slightly higher than that recorded in primary myotubes [123 ± 0.4 nM (n = 91), Fig. 7A]. In contrast, average resting Ca²⁺ concentration for myotubes expressing β_1a-V156A was 296 ± 4 nM (n = 101). No difference in resting membrane potential was observed between Wt-β_1a and β_1a-V156A-expressing myotubes (−63 ± 0.3 mV, n = 45 vs. −62 ± 0.4 mV, n = 60). It is worth noting that, because of the buffering effect of Fura-2 on intracellular Ca²⁺ (Refs. 5 and 43 and data not shown), these differences in [Ca²⁺]_i were not readily apparent from the basal fluorescence signal in Fura-2-loaded myotubes (Fig. 5A).

Fig. 7.

β_1a-V156A expressing myotubes have chronically elevated [Ca²⁺]_i and [Na⁺]_i at rest. A and B: comparison of average myoplasmic resting Ca²⁺ concentration measurements on wild-type primary myotubes (white) and β₁-null myotubes stably transfected with either Wt-β_1a (black) or β_1a-V156A subunit (gray). Calcium concentrations were measured in the presence (+) and in the absence (−) of 1 μM Gd³⁺ (A) or before (−) and after (+) 5 min incubation with 10 µM KB-R7943 (B) to inhibit cation influx. C: average intracellular Na⁺ concentration measurement of wild-type primary myotubes (white) and β₁-null myotubes stably transfected with either Wt-β_1a (black) or β_1a-V156A subunit (gray) in the presence (+) and in the absence (−) of 1 μM Gd³⁺. Extracellular Gd³⁺ significantly reduced [Na⁺]_i of Wt-β_1a and β_1a-V156A expressing myotubes but not wild-type primary myotubes. Data are presented as means ± SD. Numbers in the columns indicate the number of cells analyzed per condition; n = 3 independent experiments. *P < 0.05, ****P < 0.0001 (one-way ANOVA, Sidak’s multiple comparison test).

Inhibition of plasmalemmal Ca²⁺ entry with 1 μM GdCl₃ resulted in a drastic reduction of [Ca²⁺]_i to 98 ± 1 nM (n = 60) in Wt-β_1a-expressing myotubes and to 147 ± 2 nM (n = 64) in β_1a-V156A-expressing myotubes, with no effect on resting membrane potential in either genotype. These data reveal reductions in resting Ca²⁺ levels of ~30% and 50%, respectively, suggesting a differential inhibitory effect of Gd³⁺ on each cell group. Specific inhibition of Ca²⁺ entry through TRPC and NCX pathways by incubation with 10 μM KB-R7943 resulted in the reduction of resting Ca²⁺ concentration to levels similar to those observed with GdCl₃ (Fig. 7B).

Consistent with the above results, direct recording of intracellular [Na⁺] using Na⁺ selective microelectrodes shows that β_1a-V156A myotubes also display ~75% higher [Na⁺]_i than myotubes expressing wild-type β_1a [14.7 ± 0.1 mM (n = 55) and 8.5 ± 0.04 mM (n = 35), respectively] (Fig. 7C), with no differences in resting membrane potential [−63 ± 0.5 mV (n = 25) vs. −62 ± 1 mV (n = 25), respectively]. Similar to the effect of Gd³⁺ in resting Ca²⁺, the presence of 1 μM Gd³⁺ significantly reduced [Na⁺]_i in both cell groups although with a more drastic effect on β_1a-V156A-expressing myotubes (to 7.4 ± 0.03 mM, n = 35 in Wt-β_1a and to 9.3 ± 0.2 mM, n = 55 in β_1a-V156A myotubes) (Fig. 7C).

To explore differences in plasmalemmal Ca²⁺ entry under resting conditions between genotypes, we also measured the rates of Fura-2 fluorescence quenching induced by Mn²⁺. Figure 8 shows that β_1a-V156A-expressing myotubes displayed significantly higher rates of Mn²⁺ quench than the counterpart expressing Wt-β_1a subunit [−0.182 ± 0.008 fluorescent units (f.u.)/s (n = 164) and −0.113 ± 0.005 f.u./s (n = 131) for β_1a-V156A and Wt-myotubes, respectively]. As expected, the addition of 1 μM Gd³⁺ significantly reduced the average slope of Mn²⁺ quench in both cell genotypes (Fig. 8B). Likewise, rates of Mn²⁺ quench in the presence of 10 μM KB-R7943 were significantly lower than in untreated myotubes (Fig. 8C), with MH-susceptible cells showing higher sensitivity to KB-R7943 inhibition than wild-type cells (40% vs. 21% reduction for MH and Wt myotubes, respectively).

Fig. 8.

Effect of β_1a-V156A expression on resting Ca²⁺ entry in cultured myotubes. A: representative traces of Mn²⁺ quench of Fura-2 fluorescence of Wt-β_1a and β_1a-V156A expressing myotubes under resting conditions showing significant differences in the rate of Mn²⁺ entry between cell types. B: average rate of Mn²⁺ quench, calculated from the slope of the fluorescent signal (dashed lines), is shown for each cell phenotype in the presence and in the absence of extracellular Gd³⁺ (1 μM). C: average rate of Mn²⁺ quench in the presence and in the absence of 10 µM KB-R7943. Following loading with Fura-2 (30 min.), cells were incubated for 5 min with KB-R7943 and imaged in the presence of the inhibitor. Data are presented as means ± SE. Numbers in the bars indicate the number of cells analyzed per condition; n = 3 independent experiments. *P < 0.05, ****P < 0.0001 (unpaired t-test).

DISCUSSION

New evidence has linked malignant hyperthermia susceptibility in skeletal muscle with spontaneous mutations in several Ca²⁺ regulatory proteins other than the RyR1 calcium release channel or the DHPR α_1S subunit (8). In this study, we show evidence that expression in mouse cultured myotubes of a newly identified MH-susceptible variant of DHPR β_1a subunit (mutation V156A) results in alteration of the overall cell Ca²⁺ homeostasis that partly resembles the Ca²⁺ phenotype reported for MH-linked mutations of DHPR α_1S subunit.

Similar to MH-linked mutations of DHPR α_1S, expression of variant β_1a-V156A in β₁-null myotubes resulted in enhanced sensitivity of calcium release to caffeine stimulation. While this caffeine sensitivity appeared less prominent than that induced by many of the MH-linked mutations of RyR1, the effect of β_1a-V156A on caffeine-induced Ca²⁺ release seems consistent with the agonist sensitivities reported for some MH-linked mutations of DHPR α_1S subunit (14, 22, 38, 42). These results are also fully consistent with IVCT data that suggest that DHPR MH variants tend to have a smaller increase in the response to both caffeine and halothane than RyR1 MH variants (22).

The effects of variant β_1a-V156A on depolarization-induced Ca²⁺ release, on the other hand, was in stark contrast to the effects of MH mutations in α_1S subunit. To date, only three of the currently known MH-linked mutations of α_1S subunit have been studied in vitro (R174W, R1086H, and T1354S) and each of these mutations affected the properties of the Ca²⁺ current. The changes include total (3) or partial (42) inhibition of L-type Ca²⁺ currents, a shift in voltage dependence (42), and changes in Ca²⁺ current kinetics (38). None of the mutations, however, appeared to affect the gain of E-C coupling in the MH myotubes (orthograde signal). In contrast, our findings revealed that L-type Ca²⁺ currents of myotubes expressing β_1a-V156A were essentially unaffected, showing no significant differences in either Ca²⁺ current density or activation kinetics when compared with cells expressing Wt-β_1a. However, a slight but statistically significant leftward shift in voltage dependence of channel activation was observed in β_1a-V156A-expressing myotubes, a finding that appears consistent with studies in zebrafish muscle that show a role for β_1a-SH3 domain in the voltage-sensing properties of the skeletal DHPR (12).

Previous studies in myotubes expressing the MH mutant DHPR-α_1S-R174W described a dramatic reduction in SR Ca²⁺ content, suggesting a direct effect of this mutation on passive SR Ca²⁺ leak mediated by RyR1 and, thus, a potential model of MH susceptibility involving altered RyR1-DHPR coupling at rest (14). In our study, β_1a-V156A-expressing myotubes showed no differences in total SR Ca²⁺ load with wild-type myotubes, indicating a lack of effect of β_1a-V156A on RyR1-DHPR coupling. However, we cannot rule out a milder effect of β_1a-V156A on RyR1 function, and thus an increase in passive SR Ca²⁺ leak, that was beyond the resolution of our fluorescent assay.

We found that β_1a-V156A-expressing myotubes have chronically elevated levels of myoplasmic [Ca²⁺]_i and [Na⁺]_i under quiescent conditions in the absence of voltage or ligand-triggered Ca²⁺ release. Such elevated levels of myoplasmic [Ca²⁺]_i and [Na⁺]_i are commonly reported with MH-linked mutations in RyR1 (5, 43) and DHPR α_1S (14) and are usually associated with either RyR1 or α_1S channel dysfunction. Intriguingly, elevated resting free Ca²⁺ in β_1a-V156A-expressing myotubes seems to arise not only in the absence of a discernably altered DHPR channel phenotype but also under resting conditions where DHPR channels would not be expected to contribute to Ca²⁺ influx. We found that resting [Ca²⁺]_i and [Na⁺]_i in these cells were highly sensitive to extracellular Gd³⁺, thus strongly suggesting that alteration of cationic fluxes across the plasma membrane is a major contributor to elevated [Ca²⁺] and [Na⁺] in myotubes expressing β_1a-V156A subunit. Treatment of myotubes with KB-R7943, a widely used inhibitor of the reverse Na⁺/Ca²⁺ exchanger (NCX) that also blocks transient receptor potential canonical (TRPC) channels (29), was as effective as Gd³⁺ in lowering [Ca²⁺]_i, thus indicating that both NCX and TRPC channels are the main cationic pathways affected in these β_1a-V156A-expressing cells.

Analysis of the rates of Mn²⁺ quench further supports the hypothesis that NCX and TRPC channels are the main cationic pathways affected, because β_1a-V156A-expressing myotubes displayed significantly higher rates of extracellular Ca²⁺ entry than wild-type cells under resting conditions. Importantly, extracellular Gd³⁺ and KB-R7943 strongly inhibited Mn²⁺ quench, which is fully consistent with the observed inhibitory effect of both reagents on myoplasmic resting [Ca²⁺]. This result, therefore, confirms the hypothesis that β_1a-V156A-expressing myotubes have enhanced plasmalemmal Ca²⁺ influx.

Likewise, the inhibitory effect of extracellular Gd³⁺ on the intracellular [Na⁺] strongly suggests that this Na⁺ overload is likely mediated by an increase in net Na⁺ influx rather than a decrease in Na⁺ efflux. More importantly, as showed in Fig. 3A, these differences in Ca²⁺ and Na⁺ regulation could not be explained by a differential expression of the β_1a protein between cell genotype but rather are the result of intrinsic differences between the two proteins.

A similar alteration in [Ca²⁺]_i and [Na⁺]_i was recently reported in muscle fibers from an MH mouse model carrying RyR1-R163C mutation (2, 15). These studies linked the increased [Ca²⁺]_i and [Na⁺]_i to chronic overexpression of nonselective cationic channels (TRPC3 and TRPC6) and enhanced NCX activity (2, 15). However, it is worth noting that RyR1-R163C mutation results in Ca²⁺ dysregulation that is far more severe than that observed in β_1a-V156A mutant. In particular, RyR1-R163C shows alterations in SR Ca²⁺ content (43), L-type channel function, and gain of E-C coupling (4, 44) that were not observed in our study. Interestingly, these different characteristics seem to suggest that changes in Ca²⁺ fluxes associated to NCX and/or TRPC channel dysfunction in MH-susceptible muscles are independent of changes in SR Ca²⁺ dynamics.

The molecular mechanism by which β_1a-V156A mutation affects membrane permeability is still unclear. However, because of the location of residue V156 in the SH3 region, a domain known to be involved in protein-protein interactions (27, 41), it is plausible that the V156A mutation could prompt noncanonical interactions of the SH3 domain with ion channels other than the DHPR-α_1S subunit (41). Indeed, there is evidence for a functional association between DHPR-β isoforms and K⁺ and Cl⁻ channels as well as Zn²⁺ transporters, with consequential effects on membrane permeability (for review see Ref. 9). Likewise, a recent report has revealed novel interactions between the β_1a subunit and proteins associated with Ca²⁺ membrane transport and protein synthesis (25), suggesting that the β_1a subunit in particular is involved in a wider range of functional interactions than previously thought. Therefore, it will be important to further study the impact of β_1a-V156A on function and expression levels of TRPCs, NCXs as well as other channels involved in sarcolemmal cation transport.

At first glance, the V156A represents only a conservative amino acid substitution, but a 2°C reduction in the T_m of the purified β_1a protein suggests that this amino acid substitution does affect its core stability. A possible explanation lies in the location of this residue within the core structure of β_1a. The crystal structure of β_1a (Fig. 9) shows that V156 is located before the β4 strand in the n-Src loop of the SH3 domain, which is nestled in a hydrophobic pocket flanked by other hydrophobic residues, L93, A96, and I146. Substitution with an alanine residue reduces the hydrophobicity of this residue and could impact the assembly and hence the stability of the hydrophobic pocket. Interestingly, the corresponding residue for all three other β-subunit isoforms is isoleucine, an even more hydrophobic residue than valine. This prompts us to speculate that the role of an intermediate hydrophobic residue such as valine is to promote a degree of conformational plasticity in this region that, when altered, impacts upon protein function and normal Ca²⁺ regulation. The effect of AID peptide on thermal stability resulted in a reduction of the T_m difference between Wt and V156A of 0.5–1.0°C. The normalization of V156A is consistent with our previous studies where the addition of AID reduced the T_m difference between various β isoforms and its chimeras (35). This suggests that binding of the β_1a to the DHPR α_1S subunit through the AID region likely restores normal stability to the mutant protein. Moreover, the limited effect of V156A mutation on the overall DHPR channel function observed in this study supports the idea that Ca²⁺ dysregulation mediated by the β_1a-V156A variant is linked to an association of the mutant subunit with alternative Ca²⁺ pathways.

Fig. 9.

Locality of V156 on the β_1a crystal structure (PDB-4ZW2). The SH3 domain is shown in green, the GK domain in blue, the AID peptide in red, and the hook regions in light blue. The V156 residue is located within the SH3 domain of β_1a and forms part of a hydrophobic pocket along with residues L93, A96, and I146.

In summary, our data suggest that homozygous expression of β_1a-V156A would be sufficient to cause an MH-like phenotype in cultured myotubes and, therefore, β_1a-V156A by itself has the potential to be a pathogenic variant. It is important to note, however, that no homozygotes have been identified in any of the subjects currently reported to bear variant β_1a-V156A (ExAC browser). This suggests that the effects of β_1a-V156A might combine with other gene defects to produce an MH phenotype rather than β_1a-V156A being a single causative pathogenic variant. These results suggest that mutant variants in calcium regulatory proteins other than RyR1 and DHPR α_1S subunit can contribute significantly to both the development and the severity of the MH phenotype. Finally, our data further support the hypothesis that alteration of nonselective sarcolemmal cation permeability is a constitutive factor of MH susceptibility in skeletal muscle (2, 15).

GRANTS

Research reported in this publication was supported by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under Award Numbers R03 AR-066359 and 1R01 AR-067738 (to C. F. Perez). The work was also supported by Australian National Health and Medical Research Council Grants APP1020589 and APP1002589 (to A. F. Dulhunty and M. G. Casarotto). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or the Australian National Health and Medical Research Council.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

C.F.P. and M.G.C. conceived and designed research; C.F.P., J.M.E., J.R.l., D.B., S.A., and M.G.C. performed experiments; C.F.P., J.M.E., J.R.l., D.B., A.F.D., S.A., and M.G.C. analyzed data; C.F.P., J.M.E., J.R.l., A.F.D., and M.G.C. interpreted results of experiments; C.F.P., J.M.E., D.B., S.A., and M.G.C. prepared figures; C.F.P. and J.M.E. drafted manuscript; C.F.P., A.F.D., and M.G.C. edited and revised manuscript; C.F.P. approved final version of manuscript.