Alterations of excitation-contraction coupling and excitation coupled Ca(2+) entry in human myotubes carrying CAV3 mutations linked to rippling muscle

Abstract

Rippling muscle disease is caused by mutations in the gene encoding caveolin-3 (CAV3), the muscle-specific isoform of the scaffolding protein caveolin, a protein involved in the formation of caveolae. In healthy muscle, caveolin-3 is responsible for the formation of caveolae, which are highly organized sarcolemmal clusters influencing early muscle differentiation, signalling and Ca(2+) homeostasis. In the present study we examined Ca(2+) homeostasis and excitation-contraction (E-C) coupling in cultured myotubes derived from two patients with Rippling muscle disease with severe reduction in caveolin-3 expression; one patient harboured the heterozygous c.84C>A mutation while the other patient harbored a homozygous splice-site mutation (c.102+ 2T>C) affecting the splice donor site of intron 1 of the CAV3 gene. Our results show that cells from control and rippling muscle disease patients had similar resting [Ca(2+) ](i) and 4-chloro-m-cresol-induced Ca(2+) release but reduced KCl-induced Ca(2+) influx. Detailed analysis of the voltage-dependence of Ca(2+) transients revealed a significant shift of Ca(2+) release activation to higher depolarization levels in CAV3 mutated cells. High resolution immunofluorescence analysis by Total Internal Fluorescence microscopy supports the hypothesis that loss of caveolin-3 leads to microscopic disarrays in the colocalization of the voltage-sensing dihydropyridine receptor and the ryanodine receptor, thereby reducing the efficiency of excitation-contraction coupling.

Figure 1

Western blot analysis of skeletal muscle proteins in muscle biopsies from the two RMD patients harboring CAV3 mutations. Proteins (30 µg) in the total muscle homogenate (CAV3, α1.1 DHPR, RyR1 and glycogen phosphorylase) or total SR fraction (20 µg) (SERCA2 and calsequestrin) were blotted onto nitrocellulose and probed with the indicated antibodies as specified in the Methods section. The relative expression levels of the immunopositive bands in the biopsy from the RMD patient harboring the c.84C>A mutation (left) and the homozygous splice-site mutation c.102+ 2T>C (right) were compared to that of control biopsies that were considered 100%; intensity values were estimated by densitometric analysis of the indicated number of blots and normalized with respect to the band intensity of glycogen phosphorylase (total homogenate) or calsequestrin (SR). Bars represent mean ± SEM of n experiments; ^*P<0.0001.

Figure 2

Characterization of cytoplasmic Ca²⁺ homeostasis in myotubes with CAV3 deficiency. Calcium imaging was performed in fura-2 loaded myotubes as described in the Methods section. A: Mean (± SEM of n = 58 and 92 for control and RMD, respectively) resting [Ca²⁺] (expressed as fluorescence intensity ratio 340/380 nm) was not different in control and RMD myotubes. B: Mean (± SEM) peak Ca²⁺ increase induced by the addition of 100 mM KCl (inducing depolarization) and 600 µM 4-chloro-m-cresol (4-cmc, RyR1 agonist) in the presence of Krebs-Ringer medium (KR) containing 100 µM La³⁺. Open boxes, control myotubes; gray boxes, myotubes from CAV3 deficient myotubes (n = 7–15 measurements). C: Traces showing fura-2 change in fluorescence (ratio 340/380 nm) of individual myotubes from a control and a CAV3 mutation-bearing patient in response to 100 mM KCl and 600 µM 4-cmc.

Figure 3

Voltage-dependence of Ca²⁺ transients in control and RMD myotubes. Cells were patch-clamped and held at a holding potential (V_H) of − 80 mV. A: Paired sample traces of current (lower trace) and Ca²⁺ release recordings (upper trace) at different test potentials (from − 40 to 0 mV) in a fluo-3 loaded control myotube. Depolarizations (50 msec) to increasing membrane potentials activated Ca²⁺ release from the SR.B: Comparison of Ca²⁺ release during a depolarizing step to − 20 mV in control (black) and Cav-3 deficient RMD (red) myotubes. Linescan images and line profiles show the reduced Ca²⁺ transient amplitude in caveolin-3 deficient RMD myotubes at same trigger voltage when compared with control. C: Summary of the voltage-dependence of Ca²⁺ release in control (WT, n = 10) and caveolin-3-deficient RMD myotubes (n = 8). Ca²⁺ transient amplitudes have been normalized to the maximal release amplitude in each cell. Membrane potentials at half-maximal activation (V_1/2) indicate a right shift of the voltage-dependence in RMD myotubes (P<0.05).

Figure 4

TIRF measurements of Ca²⁺ influx induced by 60 mM KCl in human myotubes. A: Myotubes from a control patient were visualized by brightfield (top left panel), with a surface reflection interference contrast (SRIC) filter to visualize and fix the focal plane of the coverglass/cell membrane interphase (top central panel). Next panels show pseudocolored ratiometric images (peak fluorescence after addition of KCl/ resting fluorescence) of fluo-4 fluorescence changes at the indicated time-points after application of KCl. Fluorescence was monitored through a 60 × TIRF objective and analyzed using Metamorph as detailed in the Methods section. Bar indicates 30 µm. B: Representative traces from ECCE showing changes in fluo-4 fluorescence in a myotube from a control individual (_____) and a patient harbouring a CAV3 mutation (…….) stimulated with 60 mM KCl in the presence of 2 mM Ca²⁺ or myotubes from a control in the absence of added Ca²⁺ and in the presence of 100 µM La³⁺ (.-.-.-.-.). TIRF measurements were performed as indicated in the Methods section in myotubes pretreated with 100 µM ryanodine. C: Bar graph depicting mean (± SEM) peak increase of fluo-4 fluorescence induced by 60 mM KCl in control and caveolin-3 deficient myotubes in the presence of 2 mM Ca²⁺. Insert shows the mean (± SEM) peak fluo-4 fluorescence increase of control myotubes in the presence of 100 µM La³⁺ (gray bar) or 2 mM Ca²⁺ (empty bar).

Figure 5

Coimmunolocalization of the α1.1 subunit DHPR and RyR1 by immunofluorescence analysis by TIRF microscopy in human myotubes from a control individual. Myotubes were visualized using an inverted Nikon TE2000 TIRF microscope equipped with a CFI Plan Apochromat 100 × TIRF objective (1.49 NA). Left panel shows photomicrograph of cells through a SRIC filter; central left panel shows the same cells excited with a Sapphire laser at 488 nm (α1.1 subunit of the DHPR; green fluorescence); central right panel shows photomicrograph of the same cells excited at 405 and visualized through a BrightLine CH 427 filter (RyR; dark blue fluorescence). Right panel, merged images using the “color-combine” option included in the Metamorph software package. Arrows indicate overlapping pixels (light blue). Bar indicates 10 µm.