Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M403612200 on August 8, 2004

J. Biol. Chem., Vol. 279, Issue 42, 43838-43846, October 15, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/42/43838    most recent
M403612200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ducreux, S.
Right arrow Articles by Treves, S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ducreux, S.
Right arrow Articles by Treves, S.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Effect of Ryanodine Receptor Mutations on Interleukin-6 Release and Intracellular Calcium Homeostasis in Human Myotubes from Malignant Hyperthermia-susceptible Individuals and Patients Affected by Central Core Disease*

Sylvie Ducreux{ddagger}, Francesco Zorzato§, Clemens Müller¶, Caroline Sewry||**, Francesco Muntoni||, Ros Quinlivan**, Gabriella Restagno{ddagger}{ddagger}, Thierry Girard{ddagger}, and Susan Treves{ddagger}§§§

From the {ddagger}Departments of Anaesthesia and Research, Kantonsspital Basel, 4031 Basel, Switzerland, the §Dipartimento di Medicina Sperimentale e Diagnostica, Universitá di Ferrara, 44100 Ferrara, Italy, the Institut für Humangenetik, Biozentrum der Universität Würzburg, 97074 Würzburg, Germany, ||The Dubowitz Neuromuscular Centre, London W120NN, United Kingdom, the **Neuromuscular Centre, Robert Jones & Agnes Hunt Orthopaedic Hospital NHS Trust, Oswestry SY10 7AG, United Kingdom, and the {ddagger}{ddagger}Dipartimento di Patologia Clinica, S.C. Genetica Molecolare, Azienda Ospedaliera S.Anna, 10126 Torino, Italy

Received for publication, April 1, 2004 , and in revised form, August 7, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In this study we report for the first time the functional properties of human myotubes isolated from patients harboring the native RYR1 I4898T and R4893W mutations linked to central core disease. We examined two aspects of myotube physiology, namely excitation-contraction and excitation-secretion coupling. Our results show that upon activation of the ryanodine receptor (RYR), myotubes release interleukin-6 (IL-6); this was dependent on de novo protein synthesis and could be blocked by dantrolene and cyclosporine. Myotubes from the two patients affected by central core disease showed a 4-fold increase in the release of the inflammatory cytokine IL-6, compared with cells derived from control or malignant hyperthermia susceptible individuals. All tested myotubes released calcium from intracellular stores upon stimulation via surface membrane depolarization or direct RYR activation by 4-chloro-m-cresol. The functional impact on calcium release of RYR1 mutations linked to central core disease or malignant hyperthermia is different: human myotubes carrying the malignant hyperthermia-linked RYR1 mutation V2168M had a shift in their sensitivity to the RYR agonist 4-chloro-m-cresol to lower concentrations, whereas human myotubes harboring C-terminal mutations linked to central core disease exhibited reduced [Ca2+]i increase in response to 4-chloro-m-cresol, caffeine, and KCl. Taken together, these results suggest that abnormal release of calcium via mutated RYR enhances the production of the inflammatory cytokine IL-6, which may in turn affect signaling pathways responsible for the trophic status of muscle fibers.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Central core disease (CCD)1 is a congenital myopathy of autosomal dominant inheritance. Affected individuals often present with infantile hypotonia (floppy infant syndrome), weakness, hip dislocation, and a delay in achieving motor milestones. Later in life, the predominant symptom is a generalized muscle weakness affecting the proximal muscle groups more than the distal ones. The clinical severity is highly variable, but disease course is usually slow or nonprogressive (1–4). On the basis of clinical findings alone the diagnosis is difficult, and a histological examination of muscle tissue is essential. Typically, type I fibers are predominant and contain well demarcated and centrally located cores, which lack mitochondria and do not stain with histochemical dyes for oxidative enzymes. In longitudinal sections the core area is extensive and covers a considerable length of the fiber (4–6).

From a molecular point of view CCD is closely associated with malignant hyperthermia (MH), because both disorders are caused by mutations in the ryanodine receptor (RYR1), the calcium release channel of the sarcoplasmic reticulum. MH is a pharmacogenetic disease triggered by volatile anesthetics and the depolarizing muscle relaxant suxamethonium in predisposed (MH-susceptible (MHS)) individuals (7, 8). The clinical signs of an impending MH reaction are due to a hypermetabolic state and include muscle rigidity, metabolic acidosis, rhabdomyolysis, tachycardia, and/or an increase in body temperature. Histological examination of muscle fibers from MHS individuals does not reveal alterations of the status of the muscle.

To date more than 40 mutations have been identified in the ryanodine receptor gene (RYR1, human chromosome 19q13.1) and are associated with MH and/or CCD (for review see Refs. 9–11). The RYR1 gene is comprised of 106 exons that code for one of the largest known proteins (5038 amino acids) (12, 13). The first 4000 amino acids are predicted to form the hydrophilic cytoplasmic domain, whereas the last 1000 amino acids form the hydrophobic membrane-spanning pore (14–16). The majority of RYR1 mutations giving rise to the MHS phenotype appear to be clustered in the hydrophilic domain, whereas most mutations linked to CCD have been found in hydrophobic pore-forming domain 3 (17–20).

To understand the molecular mechanism underlying MH and CCD, it is essential to evaluate the impact of mutations on the functional properties of the RYR calcium channels and on excitation-contraction coupling. As far as MH-linked RYR1 mutations are concerned, in most cases they cause an increased sensitivity of the RYR to pharmacological agonists such as caffeine, 4-chloro-m-cresol, and halothane (21–25). On the other hand, mutations causing CCD have been shown to lead to leaky calcium channels that can cause depletion of intracellular calcium pools (17, 20), or to uncoupled RYR calcium channels in which depolarization of the dihydropyridine receptor does not lead to opening of the RYR calcium channel (26, 27). Thus, the impact of most disease-causing RYR1 mutations will necessarily affect calcium homeostasis of the muscle cell. To date, however, only few downstream effects of the altered intracellular calcium homeostasis have been investigated.

Calcium-dependent pathways regulate many aspects of muscle physiology, including growth and regeneration (28, 29). Increases in [Ca2+]i have been shown to activate transcription of the pro-inflammatory regulators nuclear factor {kappa}B (NF-{kappa}B), c-Jun N-terminal kinase (JNK), and nuclear factor of activated T cells (NF-AT) via activation of the calcium sensitive-phosphatase calcineurin (30). Skeletal muscle cells are capable of producing and responding to several cytokines such as IL-4, IL-6, IL-15, and tumor necrosis factor (31–34). Although the fine mechanism controlling cytokine release is not fully understood, muscle exercise has been shown to increase the plasma concentration of IL-6, which in turn has been proposed to exert a role on the maintenance of glucose homeostasis, its full physiological role, however, has yet to be established (for review see Ref. 32). When released in large quantities, IL-6 and IL-1 cause fever; i.e. they constitute an endogenous pyrogen and can induce the synthesis of acute phase plasma proteins and initiate metabolic wasting (35). Because the mechanism underlying CCD appears to be controversial, in the present report we examined how two endogenously expressed CCD-linked RYR1 mutations affect [Ca2+]i homeostasis and cytokine release from cultured human myotubes and compared their properties to those of myotubes from control individuals or MHS individuals carrying a RYR1 mutation in a different domain of the calcium release channel.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Dulbecco's modified Eagle's medium containing 4.5 mg/ml glucose, penicillin G, streptomycin, were all purchased from Invitrogen. Insulin was purchased from Eli Lilly and Co. (Indianapolis, IN). Cell culture plastic ware was from BD Biosciences. Glutamine, HEPES, bovine serum albumin, epidermal growth factor, cycloheximide, anti-{alpha}-actinin, anti-goat-FITC, anti-mouse-cy3, creatine, and dantrolene were from Sigma. Fura-2/AM and cyclosporine A were from Calbiochem. The calcium calibration kit was from Molecular Probes, Inc. A goat anti-{alpha}1.1 subunit of the skeletal muscle dihydropyridine receptor polyclonal antibodies was from Santa Cruz Biotechnology Inc. PeliKine-compact human IL-1{beta} and IL-6 ELISA kits were from CLB, Amsterdam, The Netherlands. Mouse monoclonal antibody anti-IL-6R (CD130) was from BIOSOURCE International (Camarillo, CA). All other chemicals were reagent or highest available grade.

Patients, Identification of RYR1 Mutations, and Human Skeletal Muscle Cell Culture—Primary human muscle cell cultures were established after obtaining informed consent from muscle biopsies of patients undergoing the in vitro contracture test for the diagnosis of MH susceptibility, in 10 control patients from 3 different families, 9 patients with a positive family history for MH in which the V2168M mutation in the RYR1 gene was identified (from 5 different families), and from 2 patients affected by CCD. In these patients, exons 98–103 of the RYR1 gene, the major mutation hot spot, were screened for mutations by single strand conformation analysis and direct sequencing as described previously (20).

Human skeletal muscle cell cultures were established as previously described (22). To differentiate cells into myotubes, the medium was switched to Dulbecco's modified Eagle's medium plus 4.5 mg/ml glucose, 0.5% bovine serum albumin, 10 ng/ml epidermal growth factor, 0.15 mg/ml creatine, 5 ng/ml insulin, 200 mM glutamine, 600 ng/ml penicillin G and streptomycin, and 7 mM HEPES, pH 7.4 (36).

Intracellular [Ca2+] Measurements—Cells were transferred onto glass coverslips (5 cm in diameter) and allowed to proliferate in growth medium until visible groups of >10 cells were formed. The medium was then switched to differentiation medium, and [Ca2+] measurements were performed 7–10 days later. The free cytosolic [Ca2+] was determined using the fluorescent Ca2+ indicator Fura-2 as described (22). Coverslips were mounted onto a 37 °C thermostatted chamber, which was continuously perfused with Krebs Ringer medium; individual cells were stimulated with a 12-way 100-mm diameter quartz micromanifold computer-controlled microperfuser (ALA Scientific) as previously described (37). On-line (340 nm, 380 nm, and ratio) measurements were recorded using a fluorescent Axiovert S100 TV inverted microscope (Carl Zeiss GmbH, Jena, Germany) equipped with a x20 water-immersion FLUAR objective (0.17 numerical aperture) with filters (BP 340/380, FT 425, and BP 500/530) and attached to a Hamamatsu multiformat charge-coupled device camera. The cells were analyzed using an Openlab imaging system, and the average pixel value for each cell was measured at excitation wavelengths of 340 and 380 nm. Ca2+ calibration was performed following the instructions provided with the calcium calibration kit.

Release of Cytokines—Human skeletal muscle cells were placed in the wells of a 24-microtiter plate and allowed to grow until visible aggregates of >10 cells were seen. The medium was then switched to differentiation medium for 7–10 days. On the day of the experiment the medium was changed, and cells were stimulated at 37 °C under the specified conditions. The amount of IL-1{beta} or IL-6 released into the supernatant was determined by using the CLB PeliKine Compact indirect ELISA kit following the manufacturer's instructions. All tests were performed in duplicate.

Immunofluorescence—Undifferentiated myoblasts, growing in growth medium or myotubes 7–10 days after being grown in differentiation medium, were rinsed, fixed in ice-cold methanol:acetone (1:1) for 20 min, and processed as previously described (22). Fluorescence was detected using a fluorescence Axiovert S100 TV inverted microscope (Carl Zeiss GmbH, Jena, Germany) equipped with a x20 FLUAR objective and Zeiss filter sets (BP 475/40, FT 500, and BP 530/50; and BP 546, FT 560, and BP 575–640) for detection of FITC and Cy3 fluorescence, respectively.

Statistical Analysis—Statistical analysis was performed using the Student's t test for paired samples when there were two groups and ANOVA for more than two groups. Origin computer program (Microcal Software, Inc., Northampton, MA) was used for statistical analysis and curve generation.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
RYR1 Mutation Analysis—RYR1 mutations of the nine MH patients have been identified and described earlier (38). In the two patients diagnosed clinically and histologically as having CCD we screened exons 98–103 of the RYR1 gene by SCCA. We concentrated on this region, because it is part of the highly conserved hydrophobic C-terminal domain of the RYR1 protein and has been found to be a hot spot for CCD-linked mutations (19, 20). Aberrant bands were identified, and direct DNA sequencing confirmed the presence of mutations I4898T in patient CCD1 and of R4893W in patient CCD2.

Differentiated Cultured Human Skeletal Muscle Cells Express Proteins Characteristic of Differentiated Skeletal Muscle—To demonstrate that under our experimental culture conditions human primary muscle cells differentiate sufficiently, we performed immunofluorescence studies. Fig. 1 shows the appearance of human myoblasts cultured under growth conditions (Fig. 1, A–E) or myotubes after 7–10 days of culture in differentiation medium (Fig. 1, F–J). The expression of cytoskeletal proteins specific for skeletal muscle was demonstrated using an anti-{alpha}-sarcomeric actinin mouse monoclonal antibody. This protein is expressed only in striated muscle cells and is used to identify cells or tissues of striated muscle lineage (39). Fig. 1 (B and G) shows the fluorescence pattern of cultured human muscle cells: undifferentiated myoblasts are immunonegative, while differentiated myotubes exhibit a fluorescent pattern that is compatible with that of a cytoskeletal protein. Fig. 1 (C–I) shows the immunofluorescent patterns obtained using anti-RYR1 (Fig. 1, C and H) and anti-{alpha}1.1 dihydropyridine antibodies (Fig. 1, D and I), confirming that myotubes express the two main protein components of skeletal muscle excitation-contraction coupling (40–42). We also verified whether cultured muscle cells express the IL-6 receptor, because IL-6 is released in large quantities during intense muscle activity (32). Of interest, immature myoblasts did not express the IL-6 receptor (Fig. 1E), whereas differentiated cells were immunopositive (Fig. 1J).



View larger version (45K):
[in this window]
[in a new window]
  		FIG. 1.
Photomicrograph of cultured human myoblasts and myotubes. A–E, myoblasts cultured in growth medium. F–J myotubes cultured for 7–10 days in differentiation medium. A and F, phase-contrast micrograph. Note the presence of >5 nuclei in the uppermost cell of panel F. B–J, indirect immunofluorescent staining for sarcomeric {alpha}-actinin (B and G), skeletal muscle RYR (C and H), {alpha}1.1 subunit of the dihydropyridine receptor (D and I), and IL-6 receptor (E and J). x20 Zeiss FLUAR objective; magnification, x500).

 
Activation of the RYR Leads to IL-6 Release from Cultured Human Myotubes—We next carried out experiments aimed at determining whether activation of the RYR leads to IL-6 release from cultured human myotubes. On the day of the experiment, differentiation medium was replaced with fresh differentiation medium containing no agonist or 4-chloro-m-cresol or caffeine. Cells from three control individuals were incubated at 37 °C for different time periods, and the amount of IL-6 released into the supernatant was determined by ELISA. Fig. 2 shows the time course of IL-6 release after treatment with 600 µM 4-chloro-m-cresol (Fig. 2A) and 10 mM caffeine (Fig. 2B). Maximal release was obtained 4–6 h after addition of either agonist, indicating that activation of the RYR most likely does not result in the release of pre-formed cytokine but stimulates activation of transcription and protein synthesis. The total amount of IL-6 released into the supernatant after 6 h of incubation of muscle cells from control individuals with 600 µM 4-chloro-m-cresol and 10 mM caffeine was 5.0 ± 0.7 and 7.5 ± 1.2 ng/106 cells, respectively.



View larger version (12K):
[in this window]
[in a new window]
  		FIG. 2.
Time course of IL-6 release by human myotubes from normal individuals, induced by pharmacological activation of the ryanodine receptor. A and B, cells were incubated in differentiation medium plus 600 µM 4-chloro-m-cresol (A) or 10 mM caffeine (B) at 37 °C for the indicated times. The amount of IL-6 released into the supernatant was determined by ELISA. The results are expressed as percent increase in IL-6 released; the maximal amount of IL-6 released by the RYR1 agonists was considered 100%. The results are the mean ± S.E. of three experiments carried out in duplicate on cultures established from three different control individuals. The mean maximal IL-6 released by 600 µM 4-chloro-m-cresol and 10 mM caffeine was 5.0 ± 0.7 and 7.5 ± 1.2 ng/106 cells, respectively. C, IL-6 release induced by a 5-h incubation with 600 µM 4-chloro-m-cresol is significantly reduced in the presence of dantrolene (dantro, 20 µM), cyclosporine A (CsA, 100 nM), and cycloheximide (CHX, 3 µg/ml). Bckg represents the background IL-6 release of myotubes incubated for 5 h in the absence of added agonist. Results represent the mean (± S.E.) of at least four separate experiments carried out in duplicate (Student's t test; *, p < 0.00001).

 
To verify that this IL-6 release was (i) dependent on protein synthesis and (ii) dependent on activation of the RYR1, we performed experiments on cells stimulated with 600 µM 4-chloro-m-cresol, which had been pre-treated with cycloheximide to inhibit protein synthesis (3 µg/ml, 30 min) (43), dantrolene (20 µM, 30 min) (44) to prevent activation of RYR, or cyclosporine (100 nM, 30 min) (45) to block calcium-dependent calcineurin activation of the transcription factor NF-AT. Pre-treatment with any of the three compounds resulted in a significant inhibition (>50%, p < 0.00001, Student's t test) of 4-chloro-m-cresol-activated IL-6 release (Fig. 2C). Background IL-6 release after 5 h of incubation of the myotubes at 37 °C in the absence of any RYR agonist was 31.3% ± 3.2 of the maximal release (mean ± S.E.; n = 6). Contrary to what was observed in lymphocytes (46), treatment of human myotubes with 4-chloro-m-cresol or caffeine did not result in the release of significant amounts of IL-1 (<50 pg/106 cells).

We next determined the dose-response curves of 4-chloro-m-cresol-induced and caffeine-induced IL-6 release from human myotubes obtained from control individuals. Fig. 3 shows that treatment of cells with either agonist resulted in the significant release of IL-6 (p < 0.0018; ANOVA). The release could be blocked by pre-treatment with 20 µM dantrolene. In cells from control individuals, the maximal release was obtained at a concentration of 300 µM 4-chloro-m-cresol and 10 mM caffeine: the mean (± S.E.) IL-6 released was 6.0 ± 1.1 ng/106 cells and 7.1 ± 1.1 ng/106 cells, respectively (Fig. 3, A and E). These results represent the mean of eight different experiments carried out on muscle cells from at least four different individuals. The amounts of IL-6 released by 10 mM caffeine or 300 µM 4-chloro-m-cresol were not significantly different (p = 0.497). Of interest, treatment with either agonist resulted in a larger amount (1.4-fold) of cytokine release compared with that obtained by treating the cells with 1 µM ionomycin (mean ± S.E.; IL-6 released was 5.3 ± 0.8 ng/106 cells; not shown).



View larger version (28K):
[in this window]
[in a new window]
  		FIG. 3.
Dose-dependent 4-chloro-m-cresol and caffeine-induced IL-6 release from human myotubes from control individuals and patients bearing MH and CCD-linked RYR1 mutations. Cells were incubated for 5 h in differentiation medium plus the indicated concentrations of 4-chloro-m-cresol (A–D) or caffeine (E–H); the amount of IL-6 released in the supernatant was determined by ELISA. Data represent the mean (± S.E.) of experiments carried out at least three times in duplicate. Empty boxes, cells from controls (at least four different individuals; A and E); filled boxes, cells from MHS individuals bearing the V2168M RYR1 mutation (eight different individuals; B and F); crossed boxes, cells from the CCD individual bearing the I4898T RYR1 mutation (C and G); lined boxes, cells from the CCD individual bearing the R4893W RYR1 mutation (D and H). dantro, cells incubated with 20 µM dantrolene plus 600 µM 4-chloro-m-cresol.

 
When the same experiment was performed on cells from MHS individuals carrying the RYR1 V2168M mutation, addition of 4-chloro-m-cresol and caffeine resulted in the significant release of IL-6 (p < 0.0001; ANOVA). Maximal release was obtained at a concentration of 75 µM 4-chloro-m-cresol and 10 mM caffeine: the mean (± S.E.) IL-6 released was 7.2 ± 1.3 ng/106 cells and 8.4 ± 1.3 ng/106 cells, respectively (Fig. 3, B and F). Pre-treatment of the cells with 20 µM dantrolene inhibited IL-6 release, whereas treatment with 1 µM ionomycin resulted in the release of 5.5 ± 0.8 ng/106 cells. Background release of IL-6 in the absence of added RYR agonist was similar in myotubes from control and MHS individuals (mean ± S.E.; release was 2.1 ± 0.5 and 2.99 ± 0.2 ng/106 cells, respectively).

Fig. 3 (C, D, G, and H) summarize the results obtained when experiments were performed on myotubes from the CCD patients harboring the I4898T and R4893W RYR1 mutations. The presence of either mutation in the C-terminal region of RYR1 caused a significant increase in the background levels of IL-6 released in the absence of an exogenous pharmacological activator of the RYR: the mean (± S.E.) amount of IL-6 released was 8.7 ± 0.4 and 14.7 ± 2.4 ng/106 myotubes carrying the RYR1 I4898T and R4893W mutations, respectively (p < 0.0001, ANOVA; compared with the background release of myotubes from control individuals). This background release of IL-6 blunted the stimulatory effect of 4-chloro-m-cresol to myotubes from the CCD patient harboring the I4898T mutation; the release induced by 600 µM 4-chloro-m-cresol was inhibited by pre-treatment with 20 µM dantrolene (Student's t test p < 0.0012; Fig. 3C). On the other hand, addition of caffeine caused a significant increase in IL-6 release, compared with background release (p < 0.004; ANOVA, Fig. 3G). 4-chloro-m-cresol (figure 3 panel D) and caffeine (figure 3 panel H) induced IL-6 release in myotubes from the CCD patient harboring the R4893W mutation were not significantly different; pre-treatment of these cells with 20 µM dantrolene, caused a significant reduction in IL-6 release by 600 µM 4-chloro-m-cresol (Student's t test p < 0.007).

[Ca2+]i Homeostasis in Cultured Human Myotubes Carrying RYR1 Mutations—We assessed the intracellular calcium homeostasis of myotubes from individuals carrying different mutations in RYR1 by studying: (i) resting myoplasmic free calcium concentration; (ii) KCl-induced (depolarization) calcium release and (iii) 4-chloro-m-cresol induced calcium release. All mutations caused a small but significant increase in the [Ca2+]i (Student's t test, p < 0.0001); the difference between the mean resting [Ca2+]i of muscle cells from control and RYR1-mutation bearing individuals was ~13 nM. However no difference was found when comparing [Ca2+]i from muscle cells bearing the different RYR1 mutations (p = 0.937; ANOVA, Fig. 4).



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 4.
Resting calcium concentration of human myotubes from control individuals and individuals bearing RYR1 mutations. Average resting myoplasmic [Ca2+] of cells from control, MHS individuals, and CCD individuals bearing the indicated RYR1 mutation. Values are the mean (± S.E.) of the indicated number of analyzed cells. The [Ca2+]i was measured using the fluorescent Ca2+ indicator Fura-2, in Krebs Ringer containing 1 mM CaCl2. (Student's t test, **, p < 0.0001).

 
In the next series of experiments we determined whether cells carrying different RYR1 mutations respond differently to treatment with RYR1 agonists. Fig. 5 shows a representative single cell [Ca2+]i measurement experiment: addition of the agonists resulted in an increase in the myoplasmic [Ca2+]i, which varied in extent depending on the mutation present and agonist concentration used. Fig. 6 shows the changes in fluorescence induced by the addition of different concentrations of KCl to myotubes from controls, MHS and CCD individuals. In control cells the EC50 for KCl-induced calcium release was 30 mM KCl, and maximal release was observed at 60 mM KCl, whereas in cells from MHS individuals harboring the V2168M mutation the EC50 for KCl-induced calcium release was 10 mM. In cells from the CCD patient harboring the RYR1 I4898T mutation the EC50 for KCl-induced calcium release was 40 mM, whereas cells harboring the R4893W RYR1 mutation were more sensitive to KCl-induced calcium release (EC50 for KCl was 10 mM). Cells from both CCD patients, however, released significantly less calcium compared with cells from control or MHS individuals.



View larger version (32K):
[in this window]
[in a new window]
  		FIG. 5.
Calcium release stimulated by 4-chloro-m-cresol and KCl in human myotubes from control individuals and individuals bearing RYR1-mutations. B–E, single cell intracellular Ca2+ measurements of Fura-2-loaded human myotubes. A, phase contrast; B, resting [Ca2+]i before the application of 100 mM KCl and 10 (C), 30 (D), and 40 (E) s after the application of 100 mM KCl. Myotubes were individually stimulated by addition of the agonist in Krebs-Ringer buffer containing 0.5 mM EGTA, 100 µM La3+, thus the increase in [Ca2+]i represents only release of calcium from intracellular stores. Experiments were as described under "Experimental Procedures." The bottom traces show representative curves obtained after stimulation of single cells (arrow) with 4-chloro-m-cresol (25 µM, top left; 600 µM, top right) and KCl (10 mM, bottom left; 100 mM, bottom right). Solid lines, control; dashed and dotted lines, MHS; and dotted lines, CCD.

 



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 6.
Calcium release stimulated by KCl in human myotubes. Calcium imaging was performed as described for Fig. 5 and under "Experimental Procedures": the curves show the KCl dose-dependent change in calcium, expressed as a change in fluorescence ratio (peak ratio 340/380 nm; resting ratio 340/380 nm). Each point represents the mean (±S.E.) of the change in fluorescence of 5–10 measurements. Dose-response curves were generated using the Origin software.

 
We next assessed cells from the different patients for their response to the RYR-specific activator 4-chloro-m-cresol (Fig. 7); muscle cells from control individuals had an EC50 of 120 µM for 4-chloro-m-cresol-induced calcium release, whereas all cells harboring RYR1 mutations were more sensitive to the agonist and had a lower EC50 for 4-chloro-m-cresol-induced calcium release (the EC50 values were 10 and 50 µM for cells from MHS individuals harboring the V2168M mutation and cells from CCD individuals, respectively). Of interest the cells from the CCD patient harboring the R4893W mutation released very little calcium when challenged even with high (up to 600 µM) concentrations of 4-chloro-m-cresol. As observed with KCl, cells from patients affected by CCD, irrespective of the mutation released less calcium than cells from MHS or control individuals.



View larger version (21K):
[in this window]
[in a new window]
  		FIG. 7.
Calcium release stimulated by 4-chloro-m-cresol in human myotubes. Curves show the 4-chloro-m-cresol dose-dependent change in calcium, expressed as change in fluorescence ratio (peak ratio 340/380 nm; resting ratio 340/380 nm). Each point represents the mean (±S.E.) of the change in fluorescence of 5–10 measurements. Dose-response curves were generated using the Origin software. Conditions as described for Fig. 6.

 
Finally we compared the calcium released from the intracellular stores of myotubes by 150 mM KCl, 600 µM 4-chloro-m-cresol, or 10 mM caffeine (Fig. 8). Addition of the agonists caused similar increases in the myoplasmic [Ca2+]i of the myotubes obtained from control and MHS individuals; on the other hand, irrespective of which mutation was present in the C-terminal domain of the RYR1, myotubes from CCD individuals were always less responsive, because stimulation with KCl, 4-chloro-m-cresol, or caffeine induced significantly smaller changes in the [Ca2+]i.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 8.
Increase in [Ca2+]i induced by KCl, 4 chloro-m-cresol, and caffeine in human myotubes from control, MHS, and CCD individuals. Change in fluorescence ratio (peak ratio 340/380 nm; resting ratio 340/380 nm) induced by 150 mM KCl, 600 µM 4-chloro-m-cresol, and 10 mM caffeine in Krebs-Ringer containing 0.5 mM EGTA and 100 µM La3+. Results are the mean (±S.E.) of n (6–34) independent measurements.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
This is the first report in which some of the physiological properties of myotubes harboring endogenous mutations in the C-terminal domain of RYR1 obtained from individuals affected by CCD were investigated. We (i) compared the calcium release properties of myotubes from patients bearing the different RYR1 mutations and (ii) investigated whether cultured human myotubes are capable of releasing IL-6 after activation of the ryanodine receptor and whether there are differences between cells from normal individuals and individuals bearing MH and CCD-linked RYR1 mutations.

During the past years a number of studies have addressed the role of growth factors and inflammatory cytokines in muscle physiology (31, 47). IL-6 is a member of pro-inflammatory cytokines, which also include IL-1{beta} and tumor necrosis factor{alpha}; these cytokines are also known as endogenous pyrogens, because they are capable of increasing body temperature and stimulating the release of acute phase proteins from the liver (35). Although IL-6 is released by cells of the immune system, other cell types, including skeletal muscle cells can produce and release this cytokine (32, 48, 49). Of interest, the plasma levels of IL-6 have been shown to increase by almost 100-fold after a marathon race, and a role for changes in the myoplasmic calcium concentration in IL-6 release from human muscle cells has been put forward (32, 48). Available data indicate that in skeletal muscle this cytokine may act as a hormone in the maintenance of glucose homeostasis during muscle contraction (32). However, IL-6 must also have an autocrine effect, because muscle cells express the IL-6 receptor and up-regulation of its transcript has been demonstrated following muscle injury (50).

The results of the present investigation are relevant to demonstrate an important role of calcium signaling in excitation-transcription coupling. Our results show that human myotubes are capable of releasing IL-6 several hours after RYR1 activation and that the presence of a RYR1 mutation affects the amount of IL-6 released. RYR1 activation specifically caused IL-6 release from cultured human myotubes since: (i) IL-6 release was elicited by both 4-chloro-m-cresol and caffeine, two agonists that activate calcium release via RYR1 (24, 46); (ii) IL-6 release could be blocked by pre-treatment with dantrolene, a substance that specifically closes the calcium channel, thereby blocking calcium release from the sarcoplasmic reticulum (44). Of interest, in cells from controls and MHS individuals, the concentration of 4-chloro-m-cresol causing maximal IL-6 release paralleled its maximal calcium releasing effect. On the other hand, IL-6 release from myotubes from CCD patients appears to be more complex. The most intriguing observation is that myotubes harboring RYR1 mutations in the C-terminal domain of the RYR exhibited a 4-fold increase in the background level of IL-6 released compared with cells from controls or MHS individuals. Such a background IL-6 release likely attenuates the IL-6-releasing effect of 4-chloro-m-cresol, because it would be difficult to overstimulate the IL-6-releasing pathway. The IL-6 release observed in the presence of 4-chlorocresol is sensitive to the RYR blocker dantrolene indicating that it involves RYR activation. Thus, it is plausible that the background IL-6 release may result, at least in part, from "unprompted calcium release events" via RYR. The hypothesis of unprompted release via RYR is consistent with (i) data obtained in CCD lymphocytes and with (ii) the observation that calcium release from intracellular stores of myotubes from CCD patients is decreased in the presence of EGTA in the extracellular medium, a condition that does not allow the full recharge of the calcium store (Fig. 8). The latter condition is in line with previous reports (15, 51) showing that recombinant RYR1 channels in which Ile-4898 was replaced by either Ala, Val, or Leu, or carrying a 5-amino acid deletion in the C-terminal region, failed to close completely for long time periods even in the presence of mM EGTA. We measured calcium release in the presence of EGTA to evaluate the release from the stores to exclude calcium signals generated by influx of calcium from the extracellular medium. During the determination of IL-6 release, the presence of extracellular calcium would facilitate recharging of calcium stores, enhancing the driving force of calcium release through mutated RYR via unprompted events or via RYR agonist activation. For the above reasons, we believe that the presence of extracellular calcium during the IL-6 release assay accounts for the apparent miss-correlation between the effect of 4-chloro-cresol on calcium release (Fig. 8) and on IL-6 release (Fig. 3). Altogether these results are consistent with the idea that the one or more calcium signals generated via activation of RYR1 is relevant to the signal transduction pathway leading to IL-6 release in human myotubes, but we can not exclude the possibility that other signals might also be involved. In this context it should be kept in mind that calcium-independent mechanisms have also been implicated in IL-6 release (32, 52, 53).

In many cases cytokine production is regulated by the intracellular calcium concentration, via the phosphatase calcineurin and the nuclear factors of activated T cells (NF-AT)-signal transduction pathway (30). The results of the present investigation show that activation of the RYR leads to a calcineurin-dependent IL-6 release, because pre-treatment with cyclosporine blocked release of this cytokine from human myotubes. It will be interesting to study whether release of other muscle-derived cytokines such as IL-4, IL-15, and IL-13 whose transcription is regulated by the NF-AT pathway can also be elicited by RYR activation. The release of cytokines via activation of RYR1 also has important implications in MH. This condition is characterized by hypermetabolism and a rapid increase in body temperature following the administration of trigger agents (7–9). Point mutations in RYR1 not only affect the amount of calcium released from the sarcoplasmic reticulum but also, in view of the present findings, the amount of IL-6 that can potentially be released. Because muscle mass constitutes more than 40% of total body weight, part of the downstream effects of mutations in RYR1 may include an increase in the circulating levels of IL-6, which may result in fever, muscle wasting, and fiber type switching (via Myf5 gene expression) (54). From a histological point of view MH does not induce major changes in muscle fiber appearance, whereas CCD is associated with fiber atrophy/hypotrophy (1–6). Thus alterations of calcium homeostasis caused by RYR-linked mutations in CCD patients, could potentially influence the signaling pathways responsible for the trophic status of muscle fibers and account for the different phenotypes (MH versus CCD), which are associated with mutations of the same molecule. Cytokine release experiments on a larger number of cells from patients affected by CCD may help clarify the clinical relevance of this novel observation.

During the past decade a number of studies have appeared concerning the functional impact of mutations in the RYR1 gene associated with CCD and/or the MHS phenotype (9–11). Most MH-linked mutations are clustered within two domains of the large hydrophilic region of the RYR protomer and cause a shift in the dose-response curve to agonists such as caffeine, 4-chloro-m-cresol, and halothane (9–11, 22–24). As far as the MH-linked RYR1 V2168M mutation is concerned, the results of the present investigation are consistent with our previous studies using immortalized lymphoblastoid cells of the same genotype (46) and those of Yang et al. (25) using dyspedic mouse myotubes transduced with wild type or mutated RYR1 cDNA. The presence of the mutation resulted in a shift in the EC50 of calcium release stimulated by 4-chloro-m-cresol from ~130 to 28 µM, in wild type and mutation carrying myotubes, respectively. Similarly, the dose-response curve for KCl-induced calcium release was shifted in myotubes from MHS individuals carrying the V2168M RYR1 mutation. These results confirm those of Yang et al. (25) and Gallant et al. (55) except that under our experimental conditions the EC50 for KCl-induced calcium release ~2-fold lower; the differences most likely reflect differences in the experimental approaches and the use of different calcium indicators.

CCD-linked RYR1 mutations in the C-terminal transmembrane domain have been shown to cause reduced calcium release following RYR activation (17, 20). This observation has been explained by the "leaky channel" hypothesis, which states that the presence of C-terminal CCD-linked RYR1 mutations result in a reduction of the intracellular calcium stores, which in turn cause muscle weakness, because the contractile force in normal excitation-contraction coupling is proportional to activating Ca2+. For some mutations, however, a second hypothesis has been put forward: the presence of C-terminal RYR1 mutations reduces voltage-dependent calcium release even in the absence of sarcoplasmic reticulum calcium store depletion, i.e. there is excitation-contraction uncoupling (26, 27). The results of the present investigation support the overall view that mutations in the C-terminal domain of RYR1 reduce the amount of calcium that is released via the RYR calcium channel, whereby cells expressing mutant C-terminal channels have an altered excitation-contraction coupling. As to the differences in agonist-induced calcium release between our results and those of Avila et al. (27), they probably reflect the different experimental models used; i.e. dyspedic mouse myotubes reconstituted with recombinant homozygous mutated rabbit RYR1 on a homogeneous genetic background, versus expression of the endogenous heterozygous mutation in cultured human myotubes on the patient's individual genetic background.

As far as the resting myoplasmic calcium concentration is concerned, cells carrying any RYR1 mutation had a small but significantly higher resting calcium concentration, supporting previous observations that the presence of RYR1-linked mutations does not grossly alter the resting calcium concentration of cells expressing the native endogenous mutation (22, 27). This is expected, because resting calcium is the net balance between efflux from the sarcoplasmic reticulum via the RYR, re-uptake into the sarcoplasmic reticulum via the sarcoplasmic reticulum Ca2+-ATPase, and the activity of other calcium extrusion pathways. A high myoplasmic calcium concentration would be detrimental to the cells, possibly activating calcium-dependent proteases and eventually leading to cell death.

In conclusion our data show that skeletal muscle cells respond to RYR activation by releasing IL-6, and that the presence of RYR1 mutations not only affects calcium homeostasis but, indirectly, also the amount of cytokine released. Our data also show that myotubes from patients affected by CCD release significantly more IL-6 compared with cells from control and MHS individuals and point toward a complex and heterogeneous pathogenetic pathway leading to the myopathy in CCD patients with different mutations.


   FOOTNOTES
 
* This work was supported by Swiss National Science Foundation (Grants 3200-063959.00 and 3200-067820.02, by Association Française contre les Myopathies, by the European Union Grant HPRN-CT-2002-00331, and by the Department of Anesthesia, Kantonsspital Basel. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§§ To whom correspondence should be addressed. Tel.: 41-61-265-2373; Fax: 41-61-265-3702; E-mail: susan.treves{at}unibas.ch.

1 The abbreviations used are: CCD, central core disease; MH, malignant hyperthermia; RYR, ryanodine receptor; [Ca2+]i, intracellular calcium concentration; IL-6, interleukin-6; NF-{kappa}B, nuclear factor {kappa}B; JNK, c-Jun N-terminal kinase; FITC, fluorescein isothiocyanate; ELISA, enzyme-linked immunosorbent assay; ANOVA, analysis of variance; NF-AT, nuclear factor of activated T cells. Back


   ACKNOWLEDGMENTS
 
We greatly appreciate the co-operation of the MHS and CCD patients. We also thank Evgueni Voronkov for expert technical assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Isaacs, H., Heffron, J. J., and Badenhorst, M. (1975) J. Neurol. Neurosurg. Psychiatry 38, 1177–11786[Abstract/Free Full Text]
  2. Shy, G. M., and Magee, K. R. (1956) Brain 79, 160
  3. Shuaib, A., Paasuke, R. T., and Brownell, K. W. (1987) Medicine (Baltimore) 66, 389–396[Medline] [Order article via Infotrieve]
  4. Greenfield, J. G., Cornman, T., and Shy, G. M. (1958) Brain 81, 461[Free Full Text]
  5. Hayashi, K., Miller, R. G., and Brownell, A. K. (1989) Muscle Nerve 12, 95–102[CrossRef][Medline] [Order article via Infotrieve]
  6. Sewry, C. A., Müller, C. R., Davis, M., Dwyer, J. S. M., Dove, J., Evans, G., Schröder R., Fürst, D., Helliwell, T., Laing, N., and Quinlivan, R. C. M. (2002) Neuromusc. Disorders 12, 930–938[CrossRef][Medline] [Order article via Infotrieve]
  7. Denborough, M. (1998) Lancet 352, 1131–1136[CrossRef][Medline] [Order article via Infotrieve]
  8. Larach, M. G., Localio, A. R., Allen, G. C., Denborough, M. A., Ellis, F. R., Gronert, G. A., Kaplan, R. F., Muldoon, S. M., Nelson, T. E., Ording, H., Rosenberg, H., Wand, B. E., and Wedel, D. J. (1994) Anesthesiology 80, 771–779[CrossRef][Medline] [Order article via Infotrieve]
  9. MacLennan, D. H., and Phillips, M. S. (1992) Science 256, 789–794[Abstract/Free Full Text]
  10. Jurkat-Rott, K., McCarthy, T., and Lehmann-Horn, F. (2000) Muscle Nerve 23, 4–17[CrossRef][Medline] [Order article via Infotrieve]
  11. McCarthy, T. V., Quane, K. A., and Lynch, P. J. (2000) Hum. Mut. 15, 410–417[CrossRef][Medline] [Order article via Infotrieve]
  12. Zorzato, F., Fujii, J., Otsu, K., Phillips, M., Green, N. M., Lai, F. A., Meissner, G., and MacLennan, D. H. (1990) J. Biol. Chem. 265, 2244–2256[Abstract/Free Full Text]
  13. Takeshima, H., Nishimura, S., Matsumoto, T., Ishida, H., Kangawa, K., Minamino, N., Matsuo, H., Ueda, M., Hanaoka, M., Hirose, T., et al. (1989) Nature 339, 439–445[CrossRef][Medline] [Order article via Infotrieve]
  14. Du, G. G., Sandhu, B., Khanna, V. K., Guo, X. H., and MacLennan D. H. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 16725–16730[Abstract/Free Full Text]
  15. Gao, L., Balshaw, D., Xu, L., Tripathy, A., Xin, C., and Meissner, G. (2000) Biophys. J. 79, 828–840[Medline] [Order article via Infotrieve]
  16. Chen, S. R., Li, P., Zhao, M., Li, X., and Zhang, L. (2002) Biophys. J. 82, 2436–2447[Medline] [Order article via Infotrieve]
  17. Lynch, P. J., Tong, J., Lehane, M., Mallet, A., Giblin, L., Heffron, J. J., Vaughan, P., Zafra, G., MacLennan, D. H., and McCarthy, T. V. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 4164–4169[Abstract/Free Full Text]
  18. Scacheri, P. C., Hoffman, E. P., Fratkin, J. D., Semino-Mora, C., Senchak, A., Davis, M. R., Laing, N. G., Vedanarayanan, V., and Subramony, S. H. (2000) Neurology 55, 1689–1696[Abstract/Free Full Text]
  19. Monnier, N., Romero, N. B., Lerale, J., Nivoche, Y., Qi, D., MacLennan, D. H., Fardeau, M., and Lunardi, J. (2000) Hum. Mol. Genet. 9, 2599–2608[Abstract/Free Full Text]
  20. Tilgen, N., Zorzato, F., Halliger-Keller, B., Muntoni, F., Sewry, C., Palmucci, L. M., Schneider, C., Hauser, E., Lehmann-Horn, F., Muller, C. R., and Treves, S. (2001) Hum. Mol. Genet. 10, 2879–2887[Abstract/Free Full Text]
  21. Tong, J., McCarthy, T. V., and MacLennan, D. H. (1999) J. Biol. Chem. 274, 693–702[Abstract/Free Full Text]
  22. Censier, K., Urwyler, A., Zorzato, F., and Treves, S. (1998) J. Clin. Invest. 101, 1233–1242[Medline] [Order article via Infotrieve]
  23. Wehner, M., Rueffert, H., Koenig, F., Meinecke, C. D., and Olthoff, D. (2003) Cell Calcium 34, 163–168[CrossRef][Medline] [Order article via Infotrieve]
  24. Herrmann-Frank, A., Richter, M., and Lehmann-Horn, F. (1996) Biochem. Pharmacol. 12, 149–155
  25. Yang, T., Ta, T. A., Pessah, I., and Allen, P. (2003) J. Biol. Chem. 278, 25722–25730[Abstract/Free Full Text]
  26. Avila, G., O'Brien, J. J., and Dirksen, R. T. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 4215–4220[Abstract/Free Full Text]
  27. Avila, G., O'Connell, K. M. S., and Dirksen, R. T. (2003) J. Gen. Physiol. 121, 277–286[Abstract/Free Full Text]
  28. Delling, U., Tureckova, J., Lim, H. W., DeWindt, L. J., Rotwein, P., and Molkentin, J. D. (2000) Mol. Cell. Biol. 20, 6600–6611[Abstract/Free Full Text]
  29. Constantin, B., Cognard, C., and Raymond, G. (1996) Cell Calcium 19, 365–374[CrossRef][Medline] [Order article via Infotrieve]
  30. McKay, I., and Cidlowski, J. A. (1999) Endocr. Rev. 20, 435–459[Abstract/Free Full Text]
  31. Horsley, V., Jansen, K. M., Mills, S. T., and Pavlath, G. K. (2003) Cell 113, 483–494[CrossRef][Medline] [Order article via Infotrieve]
  32. Febbraio, M., and Pedersen, B. K. (2002) FASEB J. 16, 1335–1347[Abstract/Free Full Text]
  33. Sugiura T., Harigai, M., Kawaguchi, Y., Takagi, K., Fukasawa, C., Ohsako-Higami, S., Ohta, S., Hara, M., and Kamatani, N. (2002) Int. Immunol. 178, 917–924
  34. Reid, M. B., and Li, Y. P. (2001) Acta Physiol. Scand. 171, 225–232[CrossRef][Medline] [Order article via Infotrieve]
  35. Gelfand, J. A., and Dinarello, C. A. (2001) in Harrison's Principles of Internal Medicine (Braunwald, E., Fauci, A. S., Kasper, D. L., Hauser, S. L., Longo, D. L., and Jameson, J. L., eds) 15th Ed., pp. 90–94, McGraw Hill Inc., New York
  36. Fischer-Lougheed, J., Liu, J. H., Espinos, E., Mordasisi, D., Bader, C. R., Belin, D., and Bernheim, L. (2001) J. Cell Biol. 153, 677–685[Abstract/Free Full Text]
  37. Treves, S., Pouliquin, P., Moccagatta, L., and Zorzato, F. (2001) Cell Calcium 31, 1–12
  38. Girard, T., Urwyler, A., Censier, K., Müller, C. R., Zorzato, F., and Treves S. (2001) Hum. Mut. 18, 357–358
  39. Goncharova, E. J., Kam, Z., and Geiger, B. (1992) Development 114, 173–178[Abstract]
  40. Fleischer, S., and Inui, M. (1989) Annu. Rev. Biophys. Biophys. Chem. 18, 333–364[CrossRef][Medline] [Order article via Infotrieve]
  41. Rios, E., and Pizarro, G. (1991) Physiol. Rev. 71, 849–908[Free Full Text]
  42. Benacquista, B. L., Sharma, M. R., Samso', M., Zorzato, F., Treves, S., and Wagenkecht, T. (2000) Biophys. J. 78, 1349–1358[Medline] [Order article via Infotrieve]
  43. Balciunaite, G., Keller, M., Balciunaite, E., Piali, L., Zuklys, S., Mathieu, Y. D., Boyd, R., Sussman, D. J., and Hollander, G. A. (2002) Nat. Immunol. 3, 1102–1108[CrossRef][Medline] [Order article via Infotrieve]
  44. Zhao, F., Li, P., Chen, S. R., Louis, C. F., and Fruen, B. R. (2001) J. Biol. Chem. 276, 13810–13816[Abstract/Free Full Text]
  45. Santana, L. F., Chase, E. G., Votaw, V. S., Nelson, M. T., and Greven, R. (2002) J. Physiol. 544, 57–69[Abstract/Free Full Text]
  46. Girard, T., Cavagna, D., Padovan, E., Spagnoli, G., Urwyler, A., Zorzato, F., and Treves, S. (2001) J. Biol. Chem. 276, 48077–48082[Abstract/Free Full Text]
  47. Gonzalez, E., Messi, M. L., Zheng, Z., and Delbono, O. (2003) J. Physiol. 552, 833–844[Abstract/Free Full Text]
  48. Heinrich, P. C., Behrmann, I., Müller-Newen, G., Schaper, F., and Graeve, L. (1998) Biochem. J. 334, 297–314
  49. Sehgal, P. B., Grieninger, G., and Tosato, G. (1989) Ann. N. Y. Acad. Sci. 557, 1–583
  50. Kami, K., Morikawa, Y., Sekimoto, M., and Senba, E. (2000) J. Histochem. Cytochem. 48, 1203–1213[Abstract/Free Full Text]
  51. Zorzato, F., Yamaguchi, N., Xu, L., Meissner, G., Müller, C. R., Pouliquin, P., Muntoni, F., Sewry, C., Girard, T., and Treves, S. (2003) Hum. Mol. Genet. 12, 379–388[Abstract/Free Full Text]
  52. Zumwalt, J. W., Thunstrom, B. J., and Spangelo, B. L. (1999) Endocrinology 140, 888–896[Abstract/Free Full Text]
  53. Kamdar, S. J., Fuller, J. A., and Evans, R. (1997) Exp. Cell Res. 232, 439–442[CrossRef][Medline] [Order article via Infotrieve]
  54. Friday, B. B., and Pavlath, G. K. (2001) J. Cell. Science 114, 303–310[Abstract]
  55. Gallant, E. M., and Lentz, E. R. (1992) Am. J. Physiol. 262, C422–C426

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Appl. Physiol.Home page
B. T. Corona, C. Rouviere, S. L. Hamilton, and C. P. Ingalls
Eccentric contractions do not induce rhabdomyolysis in malignant hyperthermia susceptible mice
J Appl Physiol, November 1, 2008; 105(5): 1542 - 1553.
[Abstract] [Full Text] [PDF]

Home page
 Proc. Natl. Acad. Sci. USAHome page
E. Zvaritch, F. Depreux, N. Kraeva, R. E. Loy, S. A. Goonasekera, S. Boncompagni, A. Kraev, A. O. Gramolini, R. T. Dirksen, C. Franzini-Armstrong, et al.
An Ryr1I4895T mutation abolishes Ca2+ release channel function and delays development in homozygous offspring of a mutant mouse line
PNAS, November 20, 2007; 104(47): 18537 - 18542.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
T. Yang, E. Esteve, I. N. Pessah, T. F. Molinski, P. D. Allen, and J. R. Lopez
Elevated resting [Ca2+]i in myotubes expressing malignant hyperthermia RyR1 cDNAs is partially restored by modulation of passive calcium leak from the SR
Am J Physiol Cell Physiol, May 1, 2007; 292(5): C1591 - C1598.
[Abstract] [Full Text] [PDF]

Home page
 Hum Mol GenetHome page
H. Zhou, N. Yamaguchi, L. Xu, Y. Wang, C. Sewry, H. Jungbluth, F. Zorzato, E. Bertini, F. Muntoni, G. Meissner, et al.
Characterization of recessive RYR1 mutations in core myopathies
Hum. Mol. Genet., September 15, 2006; 15(18): 2791 - 2803.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
J. R. Lopez, N. Linares, I. N. Pessah, and P. D. Allen
Enhanced response to caffeine and 4-chloro-m-cresol in malignant hyperthermia-susceptible muscle is related in part to chronically elevated resting [Ca2+]i
Am J Physiol Cell Physiol, March 1, 2005; 288(3): C606 - C612.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/42/43838    most recent
M403612200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ducreux, S.
Right arrow Articles by Treves, S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ducreux, S.
Right arrow Articles by Treves, S.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement