Attenuated Ca2+ release in a mouse model of limb girdle muscular dystrophy 2A
© DiFranco et al. 2016
Received: 23 November 2015
Accepted: 30 January 2016
Published: 24 February 2016
Mutations in CAPN3 cause limb girdle muscular dystrophy type 2A (LGMD2A), a progressive muscle wasting disease. CAPN3 is a non-lysosomal, Ca-dependent, muscle-specific proteinase. Ablation of CAPN3 (calpain-3 knockout (C3KO) mice) leads to reduced ryanodine receptor (RyR1) expression and abnormal Ca2+/calmodulin-dependent protein kinase II (Ca-CaMKII)-mediated signaling. We previously reported that Ca2+ release measured by fura2-FF imaging in response to single action potential stimulation was reduced in old C3KO mice; however, the use of field stimulation prevented investigation of the mechanisms underlying this impairment. Furthermore, our prior studies were conducted on older animals, whose muscles showed advanced muscular dystrophy, which prevented us from establishing whether impaired Ca2+ handling is an early feature of disease. In the current study, we sought to overcome these matters by studying single fibers isolated from young wild-type (WT) and C3KO mice using a low affinity calcium dye and high intracellular ethylene glycol-bis(2-aminoethylether)-n,n,n′,n′-tetraacetic acid (EGTA) to measure Ca2+ fluxes. Muscles were subjected to both current and voltage clamp conditions.
Standard and confocal fluorescence microscopy was used to study Ca2+ release in single fibers enzymatically isolated from hind limb muscles of wild-type and C3KO mice. Two microelectrode amplifier and experiments were performed under current or voltage clamp conditions. Calcium concentration changes were detected with an impermeant low affinity dye in the presence of high EGTA intracellular concentrations, and fluxes were calculated with a single compartment model. Standard Western blotting analysis was used to measure the concentration of RyR1 and the α subunit of the dihydropyridine (αDHPR) receptors. Data are presented as mean ± SEM and compared with the Student’s test with significance set at p < 0.05.
We found that the peak value of Ca2+ fluxes elicited by single action potentials was significantly reduced by 15–20 % in C3KO fibers, but the kinetics was unaltered. Ca2+ release elicited by tetanic stimulation was also impaired in C3KO fibers. Confocal studies confirmed that Ca2+ release was similarly reduced in all triads of C3KO mice. Voltage clamp experiments revealed a normal voltage dependence of Ca2+ release in C3KO mice but reduced peak Ca2+ fluxes as with action potential stimulation. These findings concur with biochemical observations of reduced RyR1 and αDHPR levels in C3KO muscles and reduced mechanical output. Confocal studies revealed a similar decrease in Ca2+ release at all triads consistent with a homogenous reduction of functional voltage activated Ca2+ release sites.
Overall, these results suggest that decreased Ca2+ release is an early defect in calpainopathy and may contribute to the observed reduction of CaMKII activation in C3KO mice.
KeywordsCalpain Skeletal muscle C3KO Ca2+ release Excitation-contraction coupling RyR1 DHPR Calpainopathy Limb girdle muscular dystrophy Calpain Skeletal muscle C3KO Ca2+ release Excitation-contraction coupling RyR1 DHPR Calpainopathy
Calpain 3 (CAPN3) is the muscle-specific member of a family of proteolytic enzymes commonly referred to as calpains . Mutations in CAPN3 lead to limb girdle muscular dystrophy type 2A (LGMD2A), a disease characterized by progressive muscle weakness and wasting . CAPN3 is present in several skeletal muscle cell fractions including the cytosolic, the myofibrillar, and the membrane fraction . In the membrane fraction, CAPN3 is highly concentrated at the triads, which are sites of contact between transverse tubules (T-tubules) and the terminal cisternae of the sarcoplasmic reticulum (SR). The interaction between voltage-sensitive calcium (Ca2+) channels (or dihydropyridine receptors (DHPR)) in the T-tubules with the SR’s Ca2+ release channels (or ryanodine receptors (RyR1)) is responsible for the excitation-contraction (E-C) coupling process .
The CAPN3 knockout mouse is used to model LGMD2A. Muscles of this mouse replicate many features of the LGMD2A phenotype such as reduced fiber diameter, mitochondrial abnormalities, and reduced muscle growth following a bout of atrophy [5–7]. In addition, there is loss of RyR1 from triad fractions, and concomitant loss of Ca2+/calmodulin-dependent protein kinase II (CaMKII) signaling.
We and others have demonstrated that CAPN3 can bind triad components such as RyR1 [3, 8], aldolase , and calsequestrin , but none of these associated proteins appear to be CAPN3 substrates . On the contrary, we observed reductions in RyR1 levels in muscles from calpain-3 knockout (C3KO) mice and LGMD2A patients with decreased or absent CAPN3 . Thus, CAPN3 appears to play a stabilizing (non-proteolytic) role for RyR1 at the triad, but the manner in which it accomplishes this task is still undefined.
Ca2+ triggers muscle contraction and regulates several signaling pathways that subsequently control downstream gene expression necessary for proper muscle remodeling, which in turn allow muscles to match gene expression with functional demands placed on them [10–12]. Such feedback leads to muscular plasticity allowing the muscle to meet metabolic demands and load-bearing capacity. We previously showed that isolated muscle fibers from C3KO release less Ca2+ upon activation than wild-type mice (C57BL) (WT) fibers but saw no difference in the rate of Ca2+ uptake . These observations led us to hypothesize that impaired Ca2+ handling might cause deficits in downstream signaling pathways reliant on Ca2+. Subsequently, we showed abnormal CaMKII signaling, blunted gene expression, and impaired muscle adaptation in muscles lacking CAPN3 . These studies revealed that the pathophysiological mechanisms underlying LGMD2A involve impaired Ca2+-mediated signaling and a weakened muscle adaptation response .
One important question in LGMD2A pathophysiology is whether the altered Ca2+ handling is an early pathological feature of calpainopathy. Our previous Ca2+ release studies were performed in aged C3KO mice (1 year old) using the permeant form of fura2-FF (fura2-FF AM, Teflabs, Texas, USA) in conjunction with video microscopy and a limited electrophysiological approach. While the affinity of this dye is appropriate to study fast Ca2+ release in skeletal muscle fibers, it is well known that permeant Ca2+ dyes can be internalized into organelles, thus confounding the interpretation of the data. In order to determine if intrinsic impairments in the Ca2+ release mechanisms are observed as an early feature of disease, here we used the impermeant form of the low affinity Ca2+ indicator Oregon Green BAPTA-5N (OGB-5N), in conjunction with large intracellular ethylene glycol-bis(2-aminoethylether)-n,n,n′,n′-tetraacetic acid (EGTA) concentrations (an approach that allows for accurate estimations of Ca2+ release fluxes in skeletal muscle fibers [13–15] and compared the properties of Ca2+ fluxes in single fibers of flexor digitorum brevis (FDB) and interosseous (IO) muscles between young adult C3KO and WT mice). The fibers were subjected to various stimulation paradigms under current and voltage clamp conditions, and OGB-5N fluorescence was measured using space-averaged and high-spatial-resolution detection approaches [16–18].
Overall, the results reveal that the magnitude of action potential which evoked Ca2+ release is indeed reduced in FDB fibers from young C3KO mice and that this impairment is consistent with a uniform reduction of the number of release sites per triad, since the kinetics and voltage dependence of the Ca2+ release process remains intact.
Animal handling and muscle fiber isolation
Geometrical parameters and membrane capacitance of isolated WT and C3KO fibers used for electrophysiological experiments
Surface area (cm2 × 10−4)
Membrane capacitance (μF/cm2)
Electrophysiology and calcium release measurements
Membrane potential was measured and controlled with a two microelectrode amplifier. Membrane potential was maintained at −90 mV in both current and voltage clamp experiments. Low resistance electrodes (~8 MΩ) were used. Both electrodes were filled with a solution (“internal solution”, see below) containing 250 μM of the low affinity calcium dye OGB-5N and a high concentration of EGTA (30 mM) semi-saturated with Ca2+ (2:1 [EGTA]:[Ca2+]). This mixture allowed for: (a) fixing the resting myoplasmic [Ca2+]concentration in all fibers to ~80 nM (a value close to the Kd of EGTA for Ca2+ at pH = 7.4), (b) recording fluorescence transients that approximate the Ca2+ release flux from the SR to the myoplasm, and (c) preventing contraction and the ensuing optical artifacts. After impalement of both electrodes, a 20-min period was allowed for equilibration between the pipette solution and the myoplasm. This assures that all measurements were performed in the presence of similar intracellular concentrations of the components of the internal solution (i.e., Ca2+ dye and EGTA). Calcium release was elicited by either action potentials (under current clamp conditions) or voltage pulses (under voltage clamp conditions). Action potentials were triggered by brief (0.5 ms) current pulses (15 % above threshold). Single current pulses or 1-s trains of current pulses (10, 20, 50, and 100 Hz) were used. The optical setup consisted of an inverted microscope equipped with (a) a standard epifluorescence attachment, (b) a custom-made confocal spot detection system, (c) a two-axis stage with 20 nm resolution scanning ability, (d) a CCD camera, and (e) two photodiode-based light detection systems [23–25].
Two fluorescence illumination/detection paradigms were used: wide-field (global) and confocal. Global illumination/detection allowed us to record space-averaged (multi-sarcomeric) Ca2+ release elicited by single and repetitive stimulation pulses (under current clamp conditions) and/or voltage pulses (voltage clamp conditions). For action potential-elicited Ca2+ release, fibers were rested 30 or 120 s, respectively, between single and repetitive current stimulations. For voltage clamp experiments, pulses of 20 ms in duration and amplitudes of −20 to 180 mV (with 10 mV increments) with respect to the resting potential were used, and fibers were rested 60 s between stimuli. Confocal illumination/detection was used to record spatially resolved (intra-sarcomeric) Ca2+ release elicited by single current pulses with 60 s rests between stimuli. For global detection, fluorescence was excited and recorded from a ~20-μm disk area at the center of the fiber, nearby the microelectrodes. For confocal recordings, a diffraction limited illumination spot (~0.8 μm FWHM) was used and the fiber was moved 200 nm stepwise [18, 25].
Ca2+ release signals are reported in terms of either dimensionless fluorescence (ΔF/F), or the underlying Ca2+ release fluxes (μM/ms), which are calculated using the single compartment model previously reported [15, 26] with modifications. The current model includes reaction equation for the following components: (a) 900 μM parvalbumin (Ca2+ binding: k on = 0.025 μM ms−1, k off = 0.7 ms−1; Mg2+ binding: k on = 1.5 × 10−5 μM ms−1, k off = 0.003 ms−1); (b) 240 μM troponin (Ca2+ binding: k on = 0.15 μM ms−1, k off = 0.45 ms−1); (c) 5 mM total adenosine 5′-triphosphate (ATP) (Mg2+ binding; k on = 0.013 μM ms−1, k off = 0.15 ms−1); (d) 30 mM EGTA (Ca2+ binding: k on = 0.0105 μM ms−1; k off = 0.00075 ms−1); (e) 250 μM OGB-5N (Ca2+ binding: k on = 0.157 μM ms−1; k off = 9.42 ms−1); (f) 5 mM total [Mg2+] (resting free [Mg2+] = 0.15 mM); and (g) 15 mM Ca2+ (resting free [Ca2+] = 0.075 μM).
Our model allows us to calculate action potential (AP)-evoked time-dependent changes in free [Ca2+] and [Mg2+]. We do not report these values since, as discussed elsewhere [15, 26], in fibers equilibrated with 15 mM EGTA, this is the dominant buffer and the free [Ca2+] transients are significantly depressed with respect to those under physiological conditions. The advantage of our approach is that movement is blocked, and [Ca2+] release fluxes can be accurately estimated [15, 26].
For current clamp experiments, we measured the amplitude of the action potentials and fluorescence transients and the depolarization at the end of the trains. For voltage clamp experiments, the peak and steady-state values of Ca2+ release transients were plotted as a function of the voltage; the data were fitted to Boltzmann functions of the form: ΔF/F = ΔF/F max / (1 + exp((V − V 1/2)/k)), where k and V 1/2 are the slope (i.e., the e-fold change) and the midpoint of the distribution, respectively.
All experiments were performed at room temperature (21–22 °C).
Three solutions were used (mM): (a) internal solution: 75 aspartate, 5 ATP di-sodium, 5 phospho-creatine di-Tris, 5 reduced glutathione, 5 MgCl2, 30 EGTA, 15 Ca(OH)2, 20 3-(n-morpholino)propanesulfonic acid, 4-morpholinepropanesulfonic acid (MOPS), ph = 7.4 with KOH. The free [Mg+2] concentration in this solution was estimated (using published Mg2+ binding constants for ATP and parvalbumin) to be 0.15 mM. (b) Tyrode: 150 NaCl, 4 KCl, 2 CaCl2, 1 MgCl2, 10 MOPS, 10 glucose, pH = 7.4 with NaOH. (c) tetraethylammonium (TEA)-Cl: 150 TEA-OH, 10 CsCl, 2 CaCl2, 1 MgCl2, 10 MOPS, pH = 7.4 with HCl.
Osmolality of solutions was 300 ± 5 mOsmol/kg H2O. The internal solution was used in all experiments, Tyrode was used for current clamp experiments, and TEA-Cl was used for voltage clamp experiments. For voltage clamp experiments, tetrodotoxin (TTX) (200 nM), nifedipine (20 μM), 9-anthracene carboxylic acid (200 μM), and TEA (150 mM) were used to block sodium, calcium, chloride, and potassium currents, respectively.
Muscle extract preparation for Western blotting
For Western blot analysis of total muscle extracts, FDB muscles were dissected and homogenized in a Dounce homogenizer in 30 volumes of reducing sample buffer (80 mM Tris, pH 6.8, 0.1 M dithiothreitol, 2 % SDS, and 10 % glycerol) with protease inhibitors cocktail (Sigma), boiled for 2 min and centrifuged at 12,000g ×10 min × 4 °C. The following antibodies were used for Western blot analyses: anti-calpain3 12A2 (Novocastra), anti-RyR1 (Thermo), anti-vinculin (Sigma), and anti-DHPRα (Thermo).
Voltage, current, and optical data were filtered at 10, 5, and 2 kHz, respectively. Data were digitized at 30 μs/point using a National Instruments board (PCI-6221). Electrical stimulation, illumination, and data acquisition were under computer control, using a custom program written in LabView.
Data are reported as mean ± SEM. Means calculated from population data were compared using the Student’s t test. Significance was set at p < 0.05.
Calcium release in response to single stimulation is smaller in C3KO fibers
Frequency dependence of Ca2+ release in C3KO and WT fibers
C3KO fibers display a similar pattern of release as WT fibers in response to stimulation paradigms (Fig. 3e–h). As expected from single stimulation experiments (Fig. 1), the response to the first pulse of each train was smaller in C3KO fibers. In addition, for all frequencies, the maximum fluorescence change and the inter-stimulus fluorescence value at the end of the trains was also smaller in the C3KO fiber than those from the WT fiber (Fig. 3a–d). Moreover, the fluorescence decay after the trains was slower in C3KO fibers. While this was not further studied, it may result from a difference in SERCA activity between the two strains.
APs recorded from WT and C3KO fibers in response to tetanic stimulation
Amplitude of the first and last action potentials (AP) elicited in WT and C3KO fibers by 1 s trains of pulses at various frequencies
Action potential amplitude (mV)
The voltage dependence of Ca2+ release is not altered in C3KO
Parameters used to fit the voltage dependence of peak and steady-state Ca2+ release in WT and C3KO fibers to Boltzmann distributions
V 1/2 (mV)
Peak release (mean ± SEM)
Steady-state release (mean ± SEM)
The k and V 1/2 parameters used to fit the voltage dependence of Ca2+ release from WT fibers (and C3KO fibers) reported here are in good agreement with those previously reported by us and others for FDB and interosseous fibers [20, 27–29].
DHPR and RYR1 concentrations are reduced in C3KO muscles
Ca2+ release in C3KO is homogeneously reduced at all triads
While the biochemical data presented here are consistent with the above Ca2+ release studies, they do not allow us to predict alterations in the spatial distribution (i.e., the topology) of the voltage-dependent release sites among the triads.
We finally used the fluorescence transients occurring at T locations (triads) to calculate the Ca2+ fluxes that would produce such transients. It should be noticed that these calculations allow estimating the actual maximal Ca2+ flux as it occurs at the sites of release (i.e., a voxel of ~1 μm3, as estimated from the xy and z resolutions of our confocal microscope ). The average Ca2+ flux transients are shown in Fig 10d, e for WT and C3KO data, respectively. For WT fibers (Fig. 10d), Ca2+ release flux at T locations (300 ± 13.5 μM/ms) is significantly larger than the 220 ± 7.4 μM/ms in C3KO fibers. Overall, these calculations demonstrate the Ca2+ release fluxes calculated at the triads are larger than those calculated from global recordings, which are space-averaged.
In the current study, we conducted a detailed analysis of Ca2+ release in a mouse model of LGMD2A and conclude that the absence of CAPN3 leads to impaired Ca2+ handling as an early feature of disease. We combined electrophysiological and optical methods to measure Ca2+-dependent fluorescence changes (ΔF/F transients) in isolated FDB fibers, loaded with a low affinity Ca2+ dye and high EGTA concentrations and maintained in current or voltage clamp conditions. Ca2+ fluxes were calculated from ΔF/F transients recorded using space-averaged (or global) and space-resolved (confocal) detection techniques.
We conducted the studies here on young mice because the phenotype of C3KO mice is progressive as the mice age . Since our previous Ca2+ measurements were performed in aged mice, we were concerned that the defects in Ca2+ release might have been a consequence of structural abnormalities that accumulated over time. The data generated here showed that Ca2+ release was reduced and the expression level of both RyR and αDHPR were decreased in young C3KO mice compared to age-matched WT mice. Thus, these changes appear to be largely independent of the degenerative processes that occur in the pathogenesis of C3KO muscles during disease progression.
Action potential evoked Ca2+ release is reduced in C3KO mice
In this study, we found that the Ca2+ fluxes elicited by single action potentials in C3KO fibers are significantly smaller than those from age-matched WT mice. These changes in amplitude are not accompanied by alterations in the kinetics of the flux transients. The limitation seen in Ca2+ release in response to single stimulation is more pronounced during repetitive stimulation, which is the physiological pattern used by the CNS to recruit mechanical output of muscles. Also, this attenuation in Ca2+ release increases with frequency, and this characteristic is expected to impact muscle mechanical output and fatigue resistance. A detailed comparative analysis of the electrophysiological data (Figs. 1, 2, 3, 4, and 5), obtained in all the regimes of stimulations used, demonstrated that differences in Ca2+ release in fibers from both genotypes of mice was not due to differences in the ability to evoke an AP.
Since electrode recordings only report membrane potential changes at the surface membrane, while Ca2+ release is controlled by the T-tubules, we measured the membrane capacitance of the fibers under experimentation. We found that alterations in Ca2+ flux in C3KO fibers was not due to differences in T-tubule-to-surface-membrane-area ratio, since the specific membrane capacitance of these fibers was not significantly different from that of WT fibers (Table 1). These data allow us to eliminate any gross structural alterations of the T-tubule system as an explanation for the data in the C3KO mice. In addition, the diameter of the limited number of C3KO fibers used for electrophysiological studies did not differ significantly from that of WT fibers (Table 1). Nevertheless, fiber diameter measurements in FDB cross-sections (Fig. 8) demonstrate that C3KO fibers are significantly smaller than WT ones, consistent with our published work and the known phenotype of LGMD2A patients. Overall, these results clearly indicate that one or more steps of the excitation contraction coupling process, beyond the action potential itself, is likely responsible for the blunted Ca2+ release in C3KO and are consistent with our observation of reduced RyR concentrations in the absence of CAPN3.
Voltage dependence of the E-C coupling mechanism in C3KO
Since action potential generation and waveform are not altered in C3KO, we explored the possibility that the voltage dependence of the E-C coupling process is impaired in C3KO fibers. In this case, we evaluated the features of ΔF/F since our model does not incorporate explicit formalisms to predict the waveforms of Ca2+ release in response to voltage pulses. Nevertheless, the analysis above demonstrates that the use of either ΔF/F or Ca2+ flux transients leads to concurrent conclusions.
We found no voltage dependence alteration in either the peak or steady-state of ΔF/F transients (and thus in Ca2+ release) in response to voltage pulses. The slope and the midpoint voltage for activation of Ca2+ release from WT and C3KO are indistinguishable from each other. Instead, only a depression of both the peak and steady-state Ca2+ fluxes was found. Both values were reduced by ~20 % with respect to control values; a depression similar to that found in AP evoked Ca2+ fluxes. These results demonstrate that the voltage-sensing step of the E-C coupling process, i.e., the response of the αDHPR to membrane depolarization, is not altered in C3KO, while Ca2+ release is scaled down. These findings point towards possible alterations in the gain of the transduction process or the release process itself as the cause of the impaired Ca2+ handling.
Ca2+ release is homogenously reduced in all triads of C3KO mice
Although it is not clear why the concentrations of both DHPR and the RyR1 are significantly reduced in C3KO mice (Fig. 9), our biochemical findings predict that there is a reduced number of release sites and/or an abnormal DHPR-RyR1 stoichiometry at the triads; this concurs with our functional data. One potential consequence of reduced expression of either, or both, RyR1 and αDHPR is an uneven distribution of functional triads along the sarcomeres. However, when we tested this possibility by means of a high temporal and spatial resolution method to detect submicron Ca2+-dependent fluorescence changes [18, 25], we found a stereotypical pattern of decreased Ca2+ release along the sarcomeres of C3KO with no sign of altered or failing triads along the stretches scanned (Figs. 9 and 10). Our confocal measurements concur with the capacitance measurements and suggest that the T-tubule system architecture is normal in C3KO mice. In addition, since the voltage dependence of the Ca2+ release is unaltered in C3KO mice, we speculate that the 1:4 RyR1:αDHPR stoichiometry to the release sites is also normal, and accordingly, our data can be explained if the number of (normal) release sites per triad is uniformly reduced along the sarcomeres.
Ca2+ release and content of calpain, αDHPR, and RyR
One explanation for why the absence of CAPN3 leads to reductions of both DHPR and RyR1 may be related to the fact that the total amount of DHPR is tightly regulated in adult fibers  and is related to the level of RyR1 . We have shown that calpain 3 maintains the RyR1 complex at the triad , thus the reduced RyR1 concentration is likely the explanation for the reduction in αDHPR.
The disparity between the reductions of Ca2+ fluxes (14–20 %) and the levels of both αDHPR (40 %) and RyR (60 %) is intriguing. In principle, the large reduction in αDHPR and RyR1 predicts a larger reduction in Ca2+ fluxes. Possible explanations are that not all the αDHPR and RyR1 participate in Ca2+ release in WT animals or alternatively that the gain of Ca2+ release is in fact increased in C3KO mice, thus partially compensating for the reduced number of release sites. Another possibility is that the impaired Ca2+ release at the triads of C3KO mice leads to a reduced Ca2+-dependent inactivation of the RyR, which partially compensates for the downregulation of RyR expression. Moreover, it is possible that the content of RyR1 and αDHPR is dependent on fiber diameter (i.e., smaller fibers are presumably more altered). Since the C3KO electrophysiological data were obtained from a population of relatively large (and uniform) diameter fibers, it is possible that their content of RyR1 and αDHPR is not as low as expected from biochemical data.
Pathophysiological mechanisms in LGMD2A
In prior studies, we and others [7, 8] demonstrated that CAPN3 may play a structural role in maintaining the integrity of the triad-associated protein complex . Importantly, we also showed that in LGMD2A patients, the concentration of RyR1 was decreased compared to healthy controls . Decreased Ca2+ in skeletal muscle can affect several downstream signaling pathways that link muscle contractile activity with gene expression. This link is extremely important as it allows muscle to adapt as needed to the level and type of activity necessary, for example, by gradual transition of muscle fiber phenotype (i.e., fast or slow) or by switching the metabolic profile (i.e., oxidative vs. glycolytic metabolism). One such signaling pathway is mediated by CaMKII, which activates the transcription factor MEF2, to enhance gene expression of slow-type genes and facilitate the transition from fast to slow fiber phenotype. CaMKII signaling and several aspects of muscle adaptation are blunted in C3KO muscles  (and data not shown).
These results suggest that decreased Ca2+ release may be an early defect in the pathogenic cascade of LGMD2A and that reduced Ca2+ release precedes impaired CaMKII signaling. The results are important because they establish key features of the pathogenic cascade in LGMD2A. Future studies are necessary to elucidate the precise role of CAPN3 at triads and its role in Ca2+ handling in skeletal muscle.
Ca2+/calmodulin-dependent protein kinase II
gene coding for calpain-3
full-duration at half-maximum
ethylene glycol-bis(2-aminoethylether)-n,n,n′,n′-tetraacetic acid
flexor digitorum brevis muscle
limb girdle muscular dystrophy type 2A
3-(n-Morpholino)propanesulfonic acid, 4-morpholinepropanesulfonic acid
Oregon Green BAPTA-5N
ryadodine receptor (intracellular Ca2+ release channel)
wild-type mice (C57BL)
alpha subunit of the dihydropyridine receptor or voltage-dependent Ca2+ channel (Cav1.1)
This work was supported by funding from the National Institute of Arthritis, Musculoskeletal and Skin Diseases (NIAMS) for a Wellstone Cooperative Muscular Dystrophy Center (U54AR052646-Sweeney), a P30 Muscular Dystrophy Core Center (NIAMS-P30AR057230-Spencer), RO1s AR048177 (Spencer), AR063710 (Heiny/Vergara), and AR041802 (Hamilton/Vergara), and an R21 (AR067422-DiFranco). Funding was also provided by grants from the Muscular Dystrophy Association (Spencer and Vergara).
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