Skeletal Muscle Ryanodine Receptor Mutations Associated with Malignant Hyperthermia Showed Enhanced Intensity and Sensitivity to Triggering Drugs when Expressed in Human Embryonic Kidney Cells

MALIGNANT hyperthermia (MH; Online Mendelian Inheritance in Man No. 145600) is a pharmacogenetic disorder triggered in MH susceptible (MHS) individuals by volatile anesthetics and depolarizing muscle relaxants. MH reactions are caused by abnormal calcium homeostasis in skeletal muscle via the skeletal muscle calcium channel ryanodine receptor 1 (RYR1) leading to a hypermetabolic state and muscle rigidity.¹  A single nucleotide change within its gene (RYR1; Online Mendelian Inheritance in Man No.180901) can lead to various skeletal muscle disorders including MH, the congenital myopathy central core disease (CCD) (Online Mendelian Inheritance in Man No. 117000),²  and multi-mini core disease (Online Mendelian Inheritance in Man No. 255320).³  Recent studies have shown that mutant forms of RYR1 have functional defects in calcium homeostasis and the defect is dependent on the position of the mutated amino acid. Mutations associated with MH including R163C have shown higher sensitivity to agonists compared with wild type (WT) in vitro,⁴  and R163C knock-in mice have also shown susceptibility in vivo to both triggering drugs and heat stress.^5,6  Mutations associated with CCD including I4898T have shown lower sensitivity and Ca²⁺ release to triggering agents, and muscle weakness has been demonstrated in mice and in human CCD individuals carrying the corresponding mutation.^7,8 

Identification of disease-related RYR1 mutations has potential importance for new diagnostic genetic tests to replace the current in vitro contracture test⁹  in Europe or caffeine-halothane contracture test¹⁰  in North America for MH diagnosis which requires invasive minor surgery. Although over 250 RYR1 mutations and polymorphisms have been reported from MHS and CCD individuals and families to date, functional studies have been carried out only for selected RYR1 mutations,¹¹  and most of the mutations are still functionally uncharacterized and are hence not recognized as “diagnostic.” It is important to understand the clinical effects of RYR1 mutations to implement simple DNA-based diagnostic tests. Functional characterization of MHS-associated mutations will also lead to a more complete understanding of the molecular mechanisms underlying skeletal muscle disorders.

Recently, we have reported functional studies of characterized RYR1 mutations using [³H]ryanodine binding assays with recombinant human RYR1 in vitro.¹²  Here, we report the analysis of four functionally uncharacterized RYR1 mutations for the first time by monitoring intracellular calcium release directly. We constructed four mutant RYR1 complementary DNA (cDNA) clones: R44C, R401C, R533C, and R533H. These mutations have been identified from MHS individuals or families but previously have not been functionally characterized. Importantly, the R401C mutation has been identified in New Zealand and Australian MHS families, and the probands have shown either fulminant clinical MH or an exercise-induced MH–like reaction.¹³  This mutation is also thought to be associated with multi-mini core disease,^14,15  which is a rare congenital myopathy and its incidence and pathology are still unknown. We also analyzed R163C and H4833Y mutations as positive controls. These are common MH-linked mutations and have been functionally characterized using calcium imaging and [³H]ryanodine binding assay with rabbit or human cDNA.^4,12,16 

Ethical approval to construct the human RYR1 cDNA clone was obtained from the Massey University Genetic Technology Committee, Palmerston North, New Zealand, acting as the Institutional Biological Safety Committee for the National Advisory Environmental Risk Management Authority.

The WT (National Center for Biotechnology Information: NM_000540.2) and mutant human RYR1 cDNAs were cloned in the plasmid cytomegalovirus vector (pcDNA3.1 (+), Life Technologies, Carlsbad, CA) using techniques described previously.^12,17  Site-directed mutagenesis was carried out using two steps: ≈1 kb polymerase chain reaction fragments were amplified to create primers for mutagenesis using a forward primer containing a specified mutation and a reverse primer without mutation. Purified polymerase chain reaction products were used as primers for mutagenesis polymerase chain reaction of RYR1 subclones which contained ≈3 kb fragments of RYR1. After sequencing both strands of purified subclones, successfully mutated plasmids were used to construct full-length RYR1 mutant clones. Primers used in amplification of mutagenesis primers are as follows: R163R; forward, 5′- GGAGAAAAGGTCTGCGTTGGGGAT-3′, R44C; forward, 5′-AGGGCTTCGGCAACTGCCTGTGCTT-3′, R401C; forward, 5′-CCGCCTGCATGATCCACAGCACCAA-3′, R533C; forward, 5′-GCACAGTTGCTACAATTGCCACGGAT-3′, R533H; forward, 5′-GCACAGTTGCTATGATTGCCACGGAT-3′. The same reverse primer was used for these reactions: reverse, 5′-TGCTTGTCCAGGAGGGAAGATG-3′. Primers for the H4833Y mutation were as follows: forward, 5′-GAACCCGCCCTGCGGTGTCTG-3′, and reverse, 5′-TTCCCATTGTAGGTGACAGAGGACA-3′. Inserts in the completed vectors were completely sequenced on both strands to confirm the absence of any unwanted amino acid substitutions due to polymerase chain reaction-induced errors.

WT and mutant RYR1 cDNAs were transiently expressed in human embryonic kidney (HEK)-293 cells expressing the large T-antigen of simian virus 40 (HEK-293T, American Type Culture Collection, Manassas, VA, CRL-11268) cells maintained in Dulbecco’s modified Eagle’s medium (Sigma–Aldrich, St. Louis, MO) containing 10% fetal bovine serum (Sigma–Aldrich) and 1% penicillin/streptomycin (Life Technologies) in a humidified atmosphere at 37°C under 5% CO₂ with media change every 2–3 days. Transient transfection was performed at 90% confluence using FuGENE HD (Hoffmann-La Roche Ltd., Basel, Switzerland) according to the manufacturer’s instructions to confirm RYR1 expression by Western blotting. Cells were harvested after 72 h, and total protein lysates were solubilized in cell lysis buffer containing 0.1 m Tris, 0.5% (v/v) Triton X-100, pH 7.8 (Sigma–Aldrich). Fifty microgram of solubilized protein was analyzed on a 7.5% sodium dodecyl sulfate polyacrylamide gel using standard protocols, and RYR1 proteins were detected using a monoclonal antibody 34C (Sigma–Aldrich) at a dilution of 1:5,000 and a monoclonal antibody against α-tubulin (Sigma–Aldrich, 1:5000) as loading control. Both antibodies were detected using horseradish peroxidase-conjugated antimouse secondary immunoglobulin G (Sigma–Aldrich).

HEK-293T cells were maintained on 0.01% (w/v) poly-d-lysine–coated glass cover slips. Transient transfection was performed using FuGENE 6 (Hoffmann-La Roche Ltd.) at 20–30% confluency. After 72-h incubation, cells were fixed with 3.7% (w/w) formaldehyde in phosphate-buffered saline (137 mm NaCl, 2.68 mm KCl, 8.1 mm Na₂HPO₄, and 1.47 mm KH₂PO₄, pH 7.4, Sigma–Aldrich) and then permeabilized with 0.1% Triton X-100 in phosphate-buffered saline. Proteins were detected using primary antibodies as follows: 34C (Sigma–Aldrich, mouse origin, 1:1000) for RYR1, and antiprotein disulfide isomerase (rabbit origin, Sigma–Aldrich, 1:1000) as an endoplasmic reticulum marker. Proteins were visualized under a fluorescence microscope BX51 (Olympus, Tokyo, Japan) using secondary antibodies: fluorescein isothiocyanate-labeled antimouse immunoglobulin G (Jackson ImmunoResearch, West Grove, PA, 1:200) for RYR1 and tetramethylrhodamine isothiocyanate-labeled antirabbit immunoglobulin G (Jackson ImmunoResearch, 1:200) for protein disulfide isomerase. 4′,6-diamidino-2-phenylindole was also used for nuclei staining.

HEK-293T cells were maintained on poly-d-lysine–coated glass slides. Transient transfection was performed as described earlier at 90% confluence using FuGENE HD. Cells were loaded after 72 h with 2 µm fura-2 AM (Life Technologies) and 0.02% pluronic F-127 (Sigma–Aldrich) in dimethyl sulfoxide (Sigma–Aldrich) at 37°C for 1 h in serum-free media. Cells were washed and analyzed in a HEPES-buffered physiological saline solution containing 25 mm HEPES, 125 mm NaCl, 5 mm KCl, 1.2 mm MgSO₄, 2 mm CaCl₂, and 6 mm glucose, pH 7.4, using an inverted fluorescence microscope IX81 (Olympus) at room temperature. Various concentrations of 4-chloro-m-cresol (4-CmC, Sigma–Aldrich, final concentrations: 50–1000 µm) were prepared in the solution and were added manually and mixed gently with the solution to obtain the final concentration to be analyzed, and the fluorescence ratio of 340/380 nm was recorded at intervals of 0.5 s. Calcium mobility was analyzed by calculation of F-F₀/F₀. Six to ten different transfection experiments were performed and 12–23 regions were analyzed per sample. Total releasable calcium was measured by addition of thapsigargin (Sigma–Aldrich).

Results were obtained as mean ± SD. Statistical analyses were performed using two-way factorial ANOVA followed by post hoc testing with Tukey–Kramer for multiple comparisons using GraphPad Prism 6 (GraphPad Software, La Jolla, CA). Sigmoidal curve fitting for EC₅₀ was determined using OriginPro 8.5 software (OriginLab Corporation, Northampton, MA) using the DoseResp function with four parameters.

Wild-type and mutant RYR1s were expressed in HEK-293T cells after 72-h transient transfection using cloned RYR1 cDNAs. Solubilized protein extracts (≈50 µg) were separated using sodium dodecyl sulfate polyacrylamide gel electrophoresis, and the RYR1 proteins were detected by Western blotting (fig. 1). No difference or abnormalities in expression were detected between WT and mutants.

RYR1 proteins are thought to be expressed on the membrane of the endoplasmic reticulum in HEK-293 cells.¹⁸  Immunofluorescence was performed to detect the colocalization of RYR1 proteins and the endoplasmic reticulum in HEK-293T cells. Merged figures indicate identical localization of RYR1 (green) and the endoplasmic reticulum marker protein disulfide isomerase (red) in HEK-293T cells (fig. 2). No difference was observed between WT and mutants.

Ca²⁺ imaging was performed using fura-2 to monitor calcium mobility in real time. In the resting state, no difference in fluorescence ratio (340/380) was observed between WT and mutants (fig. 3A, n = 12–23). Maximum response in F-F₀/F₀ was calculated and plotted to evaluate the intensity of response at each 4-CmC concentration (50, 100, 200, 300, 500, and 1,000 µm, fig. 3B). Between 90 and 100% of releasable Ca²⁺ (as measured by treatment with 800 nm thapsigargin) was released by 500 µm 4-CmC. Difference in maximum response was statistically significant in two-way factorial ANOVA (Genotype: P < 0.001, 4-CmC concentration: P < 0.001, Genotype ×4-CmC concentration: P < 0.001), and the post hoc Tukey–Kramer analysis showed that maximum response of six mutants (n = 12–23) against low concentrations of 4-CmC (50–500 µm) was significantly higher than that of WT (n = 18), except R163C and R533C at 50 µm of 4-CmC. These results indicate that these six RYR1 mutants associated with MH show a higher intensity of calcium release in response to 4-CmC compared with WT. Fluorescence ratios 340/380 were monitored and plotted at rest and after addition of 500 µm 4-CmC for each construct (fig. 4A) to analyze the response time against the agonist. It was not feasible, however, to measure accurately the time of 4-CmC addition due to the limitations of our experimental system. No significant differences were observed between WT and the six mutants. To calculate EC₅₀ of 4-CmC activation and compare sensitivity among constructs, fluorescence data were normalized using data at 1000 µm 4-CmC as a 100% response and dose–response curves were drawn using normalized data points (fig. 4B). Sigmoidal fitted curves for six mutant RYR1 proteins (n = 12–23) were shifted to the left compared with WT (n = 18), showing a lower EC₅₀ (21.2–170.4 µm) for 4-CmC activation than WT (453.2 µm). The difference between WT and mutants was statistically significant in two-way factorial ANOVA (Genotype: P < 0.001, 4-CmC concentration: P < 0.001, Genotype ×4-CmC concentration: P < 0.001), suggesting a higher sensitivity of mutants against 4-CmC than WT.

The aim of this study was to analyze the sensitivity to the RYR1-specific agonist 4-CmC of functionally uncharacterized mutations (R44C, R401C, R533C, and R533H) using calcium imaging with cloned human RYR1 cDNA expressed in HEK-293T cells. We previously sequenced RYR1 cDNA isolated from skeletal muscle of affected patients.¹⁹  This RYR1 cDNA sequence is identical to NM_000540.2, and hence this sequence was used as WT. We selected RYR1 mutations to be analyzed using the following criteria: the mutation had not been functionally characterized and each selected had been identified from MHS patients as measured by the “definitive standard” in vitro contracture test. All mutations exchange arginine, which is the most common nucleotide identified from MH-linked mutations. Each mutation clearly co-segregates with MH susceptibility in the families and none have been identified in a total of over 200 unaffected individuals. We have previously reported that MH-linked RYR1 mutants showed enhanced sensitivity to 4-CmC and CCD-linked mutants show decreased sensitivity against agonists compared with WT¹²  using a [³H]ryanodine binding assay which detects channel opening and indicates calcium release indirectly.²⁰  We performed calcium imaging in this study to detect calcium mobility in cells in vivo using F-F₀/F₀. HEK-293 cells are a widely used cell line that is thought to have low levels of nonfunctional endogenous RYR1 expression.²¹  Although epithelial HEK-293 cells are not a cell type specialized for RYR1-mediated calcium release, they have been used previously for transfection studies of RYR1 proteins.^18,22  Their main calcium store is the endoplasmic reticulum and these cells also express sarco/endoplasmic reticulum Ca²⁺-ATPase, but none of the components of the skeletal muscle calcium release channel. In conditions of transient expression of RYR1 proteins in HEK-293 cells, no detectable RYR1 proteins have been observed from a mock control without RYR1 cDNA,¹⁶  and no RYR1 was detected in mock-transfected HEK-293T cells in this study (fig. 1). HEK-293 cells have been used for calcium assays for RYR1 mutants in previous studies.^16,22  Although the experimental conditions using HEK-293T cells in this study do not mimic the in vivo situation in affected patients’ skeletal muscle, they can be used to detect differences in sensitivity against 4-CmC between WT and MH-linked mutants. Although we cannot state conclusively that these mutations cause MH in vivo, the results of this study nevertheless suggest that they cause an RYR1-specific increase in Ca²⁺ release and hence could result in susceptibility to MH. A previous study has shown that WT RYR1 may be expressed at higher levels than mutants in vivo.²³  To eliminate potential differences in expression levels of RYR1 proteins between WT and mutants, transient transfection was carried out using ≈90% confluent cells and FuGENE HD with 72-h incubation allowing very high expression levels of RYR1 for all constructs. Expression levels of RYR1 proteins were higher than that of the control protein α-tubulin as detected by Western blotting (fig. 1), and no difference was observed between WT and mutant RYR1. Expressed RYR1 proteins are thought to be localized on the internal membrane of the endoplasmic reticulum in HEK-293 cells,¹⁸  and immunofluorescence showed co-localization of RYR1 and the endoplasmic reticulum in HEK-293T cells (fig. 2). Although HEK-293T cells may not be able to mimic conditions of MH, which is a skeletal muscle disorder, these cells can be used for studies of calcium mobility and functional characterization of RYR1 mutations^16,24,25  as in this study.

Although no difference in 340/380 ratios in the resting state was observed between WT (n = 18) and mutants (n = 12–23, fig. 3A), all six mutants showed a higher intensity of response as measured by F-F₀/F₀ against low concentrations of 4-CmC (up to 500 µm) compared with WT (fig. 3B). Calcium mobility in 340/380 was monitored in real time at 0.5-s intervals and the response of WT and mutants against 500 µm 4-CmC was plotted (fig. 4A, n = 1). Apparent differences in response time between WT and mutants are evident, but it was not possible to obtain reliable data for kinetic analysis in this study because 4-CmC was added manually. Fluorescence changes would need to be monitored at less than 0.5-s intervals, and a flow cell would be required for accurate measurement of time zero for addition of agonist. Response data were normalized in percentage, and dose–response curves were used to calculate EC₅₀ values (fig. 4B). All six mutants (n = 12–23) were shifted to the left showing lower EC₅₀ values than WT (n = 18), indicating that these mutants react to 4-CmC at lower concentrations than WT and hence they are more sensitive to agonists than WT.

These observations clearly show functional differences in these mutant proteins compared with WT RYR1. This in vitro study, however, used a high density of cells (≈90% confluent) with high expression levels of RYR1 proteins. Although this protocol allows high levels of protein expression with clear differences between WT and mutants in functional analyses, it does not mimic in vivo conditions. This study focuses on functional differences between WT and MH-linked mutants. The techniques in this study provide basic methodology with which to analyze functional differences among constructs. These techniques are useful for studying MH-linked RYR1 mutations and other mutations related to CCD or multi-mini core disease and for selected mutations that may be able to discriminate between hypersensitive and hyposensitive channels. For example, the functionally characterized CCD mutation I4898T did not show any Ca²⁺ release to 4-CmC activation using either this calcium assay (data not shown) or in a [³H]ryanodine binding assay as previously reported.¹²  Most MH-linked mutations have been identified in three “hot spots” within RYR1, and these mutations may be located within functionally important domains for excitation–contraction coupling. Structural studies have revealed that N-terminal disease–related RYR1 mutations are clustered in specific key domains.²⁶  The amino acids R44 and R401 are located within a domain interacting with amino acids of other domains. Amino acid changes at these positions may cause disruption of interdomain interactions and consequent structural changes leading to a functional defect. The R533 is located on the surface of the N-terminal RYR1 protein and is close to the binding region of dantrolene and FKBP12.6.²⁷  An amino acid change at this position may change the surface structure and affect binding of agonists or modulators, leading to increased sensitivity.

Our results demonstrate functional effects of RYR1 mutations in vitro using calcium imaging with human RYR1 cDNA. MH-linked mutations show enhanced sensitivity and intensity to 4-CmC activation. This study characterizes functional effects of MH-linked mutations, and the results suggest that mutant protein functions differ depending on the amino acid change introduced within RYR1. Further investigation will be required to confirm the effects of these mutations in vivo and the cause of the disorder using different experimental approaches that more closely resemble the human situation.