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. 2007 Feb 22;580(Pt 3):745–754. doi: 10.1113/jphysiol.2006.124198

Loss of caveolin-3 induced by the dystrophy-associated P104L mutation impairs L-type calcium channel function in mouse skeletal muscle cells

Harold Couchoux 1, Bruno Allard 1, Claude Legrand 1, Vincent Jacquemond 1, Christine Berthier 1
PMCID: PMC2075458  PMID: 17317753

Abstract

Caveolins are membrane scaffolding proteins that associate with and regulate a variety of signalling proteins, including ion channels. A deficiency in caveolin-3 (Cav-3), the major striated muscle isoform, is responsible for skeletal muscle disorders, such as limb-girdle muscular dystrophy 1C (LGMD 1C). The molecular mechanisms leading to the muscle wasting that characterizes this pathology are poorly understood. Here we show that a loss of Cav-3 induced by the expression of the LGMD 1C-associated mutant P104L (Cav-3P104L) provokes a reduction by half of the maximal conductance of the voltage-dependent L-type Ca2+ channel in mouse primary cultured myotubes and fetal skeletal muscle fibres. Confocal immunomiscrocopy indicated a colocalization of Cav-3 and Cav1.1, the pore-forming subunit of the L-type Ca2+ channel, at the surface membrane and in the developing T-tubule network in control myotubes and fetal fibres. In myotubes expressing Cav-3P104L, the loss of Cav-3 was accompanied by a 66% reduction in Cav1.1 mean labelling intensity. Our results suggest that Cav-3 is involved in L-type Ca2+ channel membrane function and localization in skeletal muscle cells and that an alteration of L-type Ca2+ channels could be involved in the physiopathological mechanisms of caveolinopathies.

Caveolins are integral membrane proteins associated with cholesterol-rich omega-shaped plasma membrane pits called caveolae (Parton & Richards, 2003; Cohen et al. 2004). The mammalian caveolin protein family comprises caveolin-1 and -2, expressed in many non-muscle cells, and the muscle-specific caveolin-3 (Cav-3) (Way & Parton, 1995; Tang et al. 1996). In recent years, point mutations and deletions in the human CAV3 gene have been shown to be responsible for several muscle disorders (Cohen et al. 2004; Fulizio et al. 2005). One of the best characterized mutations of Cav-3 consists of the P104L substitution (Cav-3P104L), associated with limb-girdle muscular dystrophy 1C (LGMD 1C) (Minetti et al. 1998; Betz et al. 2001). Cav-3P104L exerts a dominant negative effect presumably through the formation of aggregates with the wild-type Cav-3 which accumulate in the Golgi complex and are degraded through a proteasome-dependent mechanism (Galbiati et al. 1999a,b; Sunada et al. 2001; Carozzi et al. 2002).

How a defective Cav-3 expression leads to muscle disorders remains a mystery, largely because of the poor understanding of the role played by Cav-3 in normal skeletal muscle function and the controversial nature of results described in the literature. Results obtained using antisense inhibition of Cav-3 expression in cultured C2C12 skeletal muscle cells suggest a role for Cav-3 in promoting myoblast fusion (Galbiati et al. 1999a) whereas opposite conclusions were drawn from results obtained on muscle cell lines derived from Cav-3-null mice and Cav-3-over-expressing mice (Volonte et al. 2003). Also, recent results obtained on zebrafish embryos after Cav-3 knockdown suggested that Cav-3 is required for correct muscle intracellular organization and myogenesis (Nixon et al. 2005). Along this line, immunolabelling experiments revealed a close association of Cav-3 with the T-tubule network in developing and mature skeletal muscle fibres (Parton et al. 1997; Ralston & Ploug, 1999; Galbiati et al. 2001). Additionally, T-tubule structure was shown to be altered in Cav-3-deficient muscles suggesting that the protein may play a role in normal T-tubule network organization (Galbiati et al. 2001; Minetti et al. 2002).

Although multiple molecular investigations have shown that Cav-3 may associate with a number of structural or signalling proteins including dystroglycan (Song et al. 1996; Sotgia et al. 2000), M-phosphofructokinase (Scherer & Lisanti, 1997), dysferlin (Matsuda et al. 2001) or nitric oxide synthase (Feron et al. 1996; Venema et al. 1997), there is still no explanation as to how Cav-3 deficiency leads to the progressive and irreversible skeletal muscle wasting that characterizes muscular caveolinopathies. Alteration of intracellular Ca2+ handling is the source of various cell disorders and is likely to be involved in a number of the most devastating degenerative muscle pathologies (Mallouk et al. 2000; Rizzuto & Pozzan, 2003). Yet, up to now, the consequences of a down-regulation of Cav-3 on skeletal muscle Ca2+ homeostasis have not been investigated. Of interest, we previously showed that depletion of membrane cholesterol in fetal skeletal fibres induced a disruption of caveolae and T-tubules and a marked reduction in the L-type Ca2+ channel function of the dihydropyridine receptor (DHPR), a critical element in the regulation of intracellular Ca2+ levels in skeletal muscle (Pouvreau et al. 2004b). This altered L-type voltage-dependent Ca2+ channel function caused by cholesterol depletion could have resulted from the associated loss of Cav-3.

Here, we investigated the properties of voltage-dependent Ca2+ channels in mouse primary cultured myotubes and in fetal muscle fibres expressing the LGMD 1C-associated Cav-3P104L mutant.

We demonstrate that the loss of Cav-3 induced by expression of the Cav-3P104L mutant induces (i) a drastic reduction in the maximum conductance of the voltage-activated L-type Ca2+ channel in both models, and (ii) a decrease in the labelling intensity of the pore-forming alpha1S subunit of the DHPR protein (Cav1.1).

Part of this work has been published in abstract form (Couchoux et al. 2006).

Methods

All experiments were carried out at room temperature (20-22°C) unless otherwise stated.

Isolation and culture of skeletal muscle cells

A suspension of muscle cells was first obtained from ribcages of 18-day-old mouse fetuses (Swiss OF1, Charles River, France) using enzymatic treatment, as previously described (Pouvreau et al. 2004b). Briefly, pregnant mice were killed by cervical dislocation and the fetuses were extracted after laparotomy and killed by decapitation, in accordance with local ethical guidelines laid down by the French Directives for Care of Laboratory Animals (decree 87–848). The two halves of the ribcage of each fetus were dissected in normal Tyrode solution containing (mm): NaCl 140, KCl 5, CaCl2 2.5, MgCl2 1 and Hepes-NaOH 10; pH 7.4. The tissues were incubated at 37°C for 15 min in normal Tyrode solution containing 3 mg ml−1 collagenase (type I, Sigma) and 1 mg ml−1 trypsin (type III, Sigma). Fibres were isolated by trituration with a sterile pipette and then allowed to sediment for 10 min according to a procedure described by Rahkila et al. (1996), plated on poly-d-lysine-coated dishes and cultured in survival medium (Dulbecco's modified Eagle's medium (DMEM) with 2% decomplemented horse serum).

For primary cultured skeletal myotubes, the supernatant from the sedimentation step was twice dispersed in dishes coated with calf skin collagens (Calbiochem) and allowed to sediment for 15 min to remove fibroblasts. Myoblasts from the supernatant were grown in proliferation medium consisting of DMEM with 10% fetal bovine serum, 10% horse serum and 2% chick embryo extract until transfection. All cells were cultivated at 37°C in 5% CO2.

Transfection

Plasmids encoding yellow fluorescent protein (YFP)-tagged wild-type Cav-3 (Cav-3wt–YFP) and Influenza Hemaglutinin epitope (HA)-tagged Cav-3P104L were generously provided by R. Parton (University of Queensland, Australia). YFP-tagged Cav-3P104L (Cav-3P104L–YFP) was generated by subcloning into enhanced YFP-encoding vector (pEYFP; Clontech) using Kpn I/BamHI restriction sites.

Fetal fibres were transfected after 2 days in survival medium using CombiMag (OzBiosciences) combined with FuGene 6 (Roche Diagnostics). Primary myoblasts were transfected at ∼70% confluency using the same method and were allowed subsequently to differentiate by switching to DMEM with 10% fetal bovine serum. Immunostaining and electrophysiology experiments were performed 5 days after transfection.

Immunofluorescence staining

Cells were fixed with 1% paraformaldehyde–0.01% glutaraldehyde in PBS (10 min) and permeabilized with 1% Triton X-100–50 mm glycine in PBS (5 min). Non-specific sites were blocked with 3% normal goat serum (NGS)–0.1% bovine serum albumin in PBS (30 min) and endogenous biotins were saturated using the Avidin/Biotin Blocking Kit (Vector Laboratories). Cells were then incubated for 2 h with anti-Cav-3 rabbit polyclonal antibody (PA1-066, Affinity BioReagents) or anti-Cav1.1 mouse monoclonal antibody (mAb427, Chemicon International) diluted 1: 1500 and 1: 500, respectively, in PBS with 1% NGS. Cells were subsequently incubated for 30 min with biotin-conjugated goat anti-rabbit or goat anti-mouse IgG (Jackson Immunoresearch Laboratories) diluted 1: 1000 in PBS with 1% NGS and treated for 30 min with Cy3- or Alexa488-conjugated streptavidin (Jackson Immunoresearch Laboratories or Molecular Probes, respectively) diluted 1: 1000 in PBS. For sequential labelling, cells were first labelled with the anti-Cav-3 primary antibody then labelled with the anti-Cav1.1 primary antibody after performing a mouse non-specific sites blocking step (M.O.M. blocking kit, Vector Laboratories). Preparations were all mounted in antifading medium (Vectashield) and examined using an LSM 510 confocal fluorescence microscope (Zeiss). Excitation was performed with argon and helium–neon lasers at 488 or 533 nm and optical sections were collected using a 63 × oil immersion objective and appropriate band pass filters.

Image analysis

Cav1.1 labelling was quantified by numerical analysis of Z-series of confocal images taken from Cav-3P104L–YFP transfected primary cultures after immunolabelling of Cav1.1. The confocal Z-step size was set to 0.5 μm. We rigorously ensured that settings (including photomultiplier gain, offset, aperture and laser power) remained identical to allow comparison of the labelling intensity between different samples.

Field-specific analysis was performed by measuring Cav1.1 immunoreactivity of Cav-3P104L–YFP transfected myotubes and of non-transfected control myotubes present on the same RGB image. RGB images corresponding to each optical section of a given Z-stack were analysed using the Image J software (National Institutes of Health, Bethesda, MA, USA). Analysis was performed after extracting layers of each RGB image: the red layer corresponded to the Cav1.1 labelling (Cy3 image) and the green layer corresponded to the Cav-3P104L–YFP signal. In order to quantify the labelling relative to the cell surface area, the cell outer limits had to be estimated from the fluorescence signals. In control myotubes this was achieved on the Cy3 image whereas in Cav-3P104L–YFP transfected myotubes the Cav-3P104L–YFP signal was used because of the very weak, unreliable Cav1.1 labelling in some of these myotubes. Non-labelled regions that clearly corresponded to nuclei were excluded from the area measurement.

Within the thus selected regions, the Cav1.1 immunoreactivity of Cav-3P104L–YFP-expressing myotubes and control myotubes was quantified on the Cy3 image by measuring the sum of the values of the pixels (integrated density) after optimally setting the threshold to eliminate background signal.

Mean Cav1.1 labelling intensity was determined for each control and transfected myotube present on a given image by normalizing the integrated density to the cell area. These measurements were performed on all images of the confocal stack and seven different stacks of images were analysed.

Electrophysiology

Membrane currents were recorded in the whole-cell configuration on cultured myotubes and fetal fibres using a patch-clamp amplifier (RK 400, Bio-Logic). Pipettes were filled with a solution containing (mm): caesium aspartate 110, TEA chloride 20, MgCl2 2, MgATP 5, EGTA 10 and Hepes 10; pH 7.2. The bath solution contained (mm): TEA MeSO3 140, CaCl2 10, MgCl2 2, TTX 0.001 and Hepes 10; pH 7.2. Voltages were not corrected for liquid junction potentials, which were calculated to be < 5 mV with the solutions used. Data acquisition and generation of command voltage pulses were done using the pClamp9 software driving the Digidata 1322A A/D and D/A converter (Axon Instruments). Cell capacitance, used to calculate the density of currents (A/F), was determined by integration of a control current trace obtained with a 10 mV depolarizing pulse from −80 mV. Leak currents were subtracted from all recordings using a 10 mV depolarizing pulse from the holding potential supposing a linear evolution of leak current with depolarization. The amplitude of the currents was measured at the peak of the currents and individual curves of the voltage dependence of ICa,L density were fitted with eqn (1):

graphic file with name tjp0580-0745-m1.jpg (1)

where I(V) is the density of the current measured, V is the test pulse, Gmax is the maximum conductance per capacitance, Vrev is the apparent reversal potential, V0.5 is the half-activation voltage and k is a steepness factor. Individual curves of the voltage dependence of the normalized conductance were obtained by dividing I(V) by Gmax(V − Vrev).

Statistics

Non-linear least-squares fits were performed using a Marquadt–Levenberg algorithm routine included in Microcal Origin. Data values are presented as means ± s.e.m. In the figures, values of standard errors are in some cases smaller than data symbols and are therefore not apparent. Data were statistically analysed using unpaired Student's t test. Values were considered significant when P < 0.05.

Results

Cav-3P104L expression leads to the loss of endogenous Cav-3 in mouse primary cultured myotubes

We first examined Cav-3 imunolocalization in both control and Cav-3P104L-expressing primary myotubes. The immunolabelling patterns shown in Fig. 1 are representative of results obtained in several independent experiments (control, n = 5; Cav-3P104L-transfected myotubes, n = 6).

Figure 1. Cav-3P104L–YFP expression leads to the loss of Cav-3 in primary cultured myotubes.

Figure 1

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A, in control myotubes, Cav-3 immunolabelling strongly outlines the surface plasma membrane. It also localizes intracellularly in small dot-shaped clusters (arrowheads) as best seen on the partial 2 × enlargement (inset). B, Cav-3P104L–YFP accumulates preferentially around the nuclei in vesicular structures (top panel). Cav-3 immunolabelling is greatly decreased, if not totally absent, in the Cav-3P104L–YFP-expressing myotube (arrow) whereas membrane staining is seen at the extremity of a nearby non-transfected myotube (bottom panel). Scale bars, 10 μm.

In control myotubes (Fig. 1A), Cav-3 labelling (red) was intense at the surface plasma membrane and was also characteristically present more discretely throughout the cell where it exhibited a punctate or reticular pattern consistent with the developing T-tubule membrane system (Flucher et al. 1991; Parton et al. 1997).

As shown in Fig. 1B, Cav-3P104L-expressing myotubes displayed a different pattern of immunostaining (bottom panel). YFP-tagged Cav-3P104L (top panel) localized in perinuclear vesicles probably belonging to the Golgi apparatus. The corresponding Cav-3 labelling indicated that endogenous Cav-3 no longer reached the sarcolemma, and was usually barely detected (Fig. 1B). Our data thus confirm that expression of the mutant Cav-3P104L in primary myotubes induces a drastic loss of endogenous membrane-associated Cav-3, as previously shown in skeletal muscle of adult transgenic mouse and LGMD-1C patients (Minetti et al. 1998; Sunada et al. 2001). Golgi-associated mutant or endogenous caveolin was not detected by the PA1-066 antibody, probably because of epitope-masking of the corresponding N-terminal epitope when in the Golgi compartment as previously suggested (Luetterforst et al. 1999).

The amplitude of the L-type Ca2+ current is reduced in Cav-3P104L-expressing mouse primary cultured myotubes

The properties of voltage-dependent Ca2+ channels were studied in control and Cav-3P104L-expressing myotubes. Two types of voltage-dependent Ca2+ currents typical of cultured skeletal myotubes were recorded. T-type calcium current (ICa,T), displaying fast activating and inactivating kinetics, developed for depolarizations within the −40 to 0 mV range and L-type Ca2+ current (ICa,L), displaying slow activating and inactivating kinetics, was recorded for depolarizations to values more positive than −20 mV (Fig. 2A). We found that the amplitude of ICa,L was strongly reduced in Cav-3P104L-expressing myotubes as compared to control myotubes whereas ICa,T remained unchanged (Fig. 2A and B). For each cell, the current–voltage relationship of ICa,L was fitted using eqn (1) (see Methods). Mean values for Gmax, V0.5, Vrev and k were 197 ± 12 S F−1, 9 ± 0.8 mV, 65 ± 1.6 mV and 5 ± 0.4 mV for control myotubes (n = 40) and 88 ± 13 S F−1, 11 ± 1.9 mV, 55 ± 2 mV and 5 ± 0.5 mV for Cav-3P104L-expressing myotubes (n = 21), respectively (Fig. 2C). In Cav-3P104L-expressing myotubes, there was a mean reduction of 55% of Gmax and a 10 mV shift of Vrev towards negative potentials (P < 0.0001 and P < 0.001, respectively). The slight shift of Vrev was probably due to unmasking of a contaminating outward current component as a consequence of the decreased amplitude of the inward Ca2+ current. In myotubes transfected with YFP alone, Gmax (191 ± 10 S F−1, n = 6) was not significantly changed. Finally, we found that the mean capacitance was not significantly different in control myotubes (218 ± 20 pF, n = 40) and in Cav-3P104L-expressing myotubes (229 ± 27 pF, n = 34).

Figure 2. ICa,L is reduced in Cav-3P104L-expressing primary cultured myotubes.

Figure 2

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A, representative currents elicited by applying 500 ms duration voltage pulses from a holding potential of −80 mV to the indicated values in the two types of cells. B, mean current–voltage relationships established for the fast transient current (ICa,T) and for the slow activating current (ICa,L) in 40 control and 21 Cav-3P104L-expressing myotubes. C, mean current–voltage relationships of isolated ICa,L on the same control and mutant myotubes. The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details).

Expression of Cav-3P104L in myotubes alters the distribution and mean intensity of L-type voltage-dependent Ca2+ channel immunolabelling

A possible reason for the drastic reduction in the amplitude of ICa,L induced by Cav-3P104L expression is that loss of Cav-3 leads to decreased expression or defective targeting of the DHPR.

To explore this possibility, control and Cav-3P104L-expressing myotubes were immunostained with anti-Cav-3 and anti-Cav1.1 antibodies. Representative images are shown in Figs 3 and 4. In control myotubes (Fig. 3) both proteins essentially colocalized; this was particularly apparent at the level of the surface plasma membrane. In the internal cellular compartment, such a colocalization is also visible although it appears more restricted to specific areas (arrowheads). The Cav1.1 labelling corresponded to an extended array of regularly distributed dots at the level of which Cav-3 labelling was, although not systematically, also concentrated. As for Fig. 1, these labelled internal structures probably belong to the nascent T-tubule system as previously documented (Flucher et al. 1991, 1993; Parton et al. 1997).

Figure 3. Double immunolabelling reveals a strong colocalization of Cav-3 and Cav1.1 in primary cultured myotubes.

Figure 3

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Cav-3 (red) and Cav1.1 (green) strongly colocalize as clearly shown on the merged image. The colocalization is particularly pronounced at the surface plasma membrane, where both proteins accumulate (arrows). Additionally, the Cav1.1 labelling appears characteristically concentrated within internal punctate structures distributed almost uniformly throughout the cell and probably belonging to the developing T-tubule network. Cav-3 labelling appears generally weaker in this internal compartment except for specific areas, where it colocalizes with Cav1.1 (arrowheads). Scale bar, 10 μm.

Figure 4. Localization and labelling intensity of Cav1.1 are altered in Cav-3P104L-expressing myotubes.

Figure 4

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A, in this representative Cav-3P104L–YFP expressing myotube (green), Cav1.1 labelling (red) appears much dimmer than in the non-transfected neighbouring myotubes and is also partly redistributed to the perinuclear region where it colocalizes with Cav-3P104L, as shown on the merged image (arrow). Scale bar, 10 μm. B, numerical image analysis of Cav1.1 labelling intensity. Field-specific analysis was performed on confocal Z-stacks by measuring Cav1.1 immunoreactivity of control myotubes and Cav-3P104L-expressing myotubes present on the same image. Mean labelling intensity appeared to be reduced by 66% in Cav-3P104L-expressing myotubes (P < 0.0001).

In Cav-3P104L-expressing myotubes, Cav1.1 labelling was clearly reduced (Fig. 4A). In addition, part of the residual labelling appeared to redistribute around the nuclei, probably within the Golgi compartment, as indicated by the colocalization with Cav-3P104L–YFP (merged image).

Cav1.1 labelling intensity (mean pixel value per pixel of cell area) was measured in individual control and in Cav-3P104L-expressing myotubes using numerical analysis of confocal images. As shown in Fig. 4B, the mean Cav1.1 labelling intensity in Cav3P104L-expressing myotubes (23 ± 2.4 mean pixel value per pixel of cell area, n = 56) was significantly reduced by 66% as compared to control myotubes (68.2 ± 3.3 mean pixel value per pixel of cell area, n = 134; P < 0.0001). Identical results were obtained if, in order to limit any bias introduced by variability within the different sampled images, the labelling densities were compared on pairs of control and Cav-3P104L-expressing myotubes present in the same image field (data not shown).

Cav-3P104L expression in fetal skeletal muscle fibres also leads to the loss of endogenous Cav-3 and to a reduction of ICa,L amplitude

As Cav-3 is thought to be involved in muscle developmental processes, it is possibile that the reduction of the voltage-dependent Ca2+ channel function in Cav-3P104L-expressing cultured myotubes could result from abnormal myotube differentiation. To test this, we explored channel function in fetal skeletal muscle fibres maintained in survival medium and transfected with YFP-tagged Cav-3P104L. This cell system, although not fully mature, possesses a well-differentiated skeletal muscle phenotype and is suitable for transfection.

As shown in Fig. 5A (top panel), Cav-3 immunolabelling was present at the level of both the surface and T-tubule membrane in the control fibres (cell on the right) whereas the labelling intensity was strongly decreased in the Cav-3P104L-expressing fetal fibres (cell on the left) indicating that, as in cultured myotubes, expression of the mutant Cav-3P104L induced an important loss of endogenous Cav-3. It is also worth noting that in fetal fibres Cav-3P104L–YFP also accumulates in the perinuclear region. We then measured Ca2+ currents in Cav-3P104L-expressing fetal fibres. In survival medium, ICa,T vanished after a few days in culture whereas robust ICa,L could still be recorded (Fig. 5B, left and middle traces). This current was typical of fetal skeletal muscle fibres (Pouvreau et al. 2004b) with slow kinetics of activation, a threshold at −10 mV, a peak at +20 mV and Vrev close to +80 mV (Fig. 5C). Its amplitude was strongly decreased in Cav-3P104L-expressing fibres as compared to control fibres. Mean values for Gmax, V0.5, Vrev and k were 203 ± 11 S F−1, 10 ± 0.9 mV, 77 ± 1.1 mV and 4 ± 0.2 mV in control fibres (n = 31) and 85 ± 18 S F−1, 16 ± 3.5 mV, 63 ± 5 mV and 4 ± 0.7 mV in Cav-3P104L-expressing fibres (n = 8), respectively. Gmax was significantly reduced by 58% (P < 0.0001) and Vrev and V0.5 were significantly shifted by −14 mV (P < 0.0001) and +6 mV (P < 0.02), respectively, in Cav-3P104L-expressing fibres as compared to controls. The voltage shift of V0.5 is clearly shown in the right panel of Fig. 5C, where the mean normalized conductance–voltage relationship in control and Cav-3P104L-expressing fetal fibres is presented. Fitting the individual normalized conductance–voltage relationship with a Boltzman equation gave mean values for V0.5 and k of 12 ± 4.7 mV and 3.4 ± 2 mV in control fibres and of 17.7 ± 2.4 mV and 3.2 ± 0.6 mV in Cav-3P104L-expressing fetal fibres, respectively. The mean capacitance of Cav-3P104L-expressing fibres (182 ± 13 pF, n = 8) did not significantly differ from that of control fibres (175 ± 21 pF, n = 31).

Figure 5. ICa,L is altered by Cav-3P104L expression but is not affected by wild-type Cav-3 expression in fetal fibres.

Figure 5

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A, immunolocalization of Cav-3 in fetal fibres expressing Cav-3P104L–YFP (top panel) and YFP-tagged wild-type Cav-3 (WT-Cav-3, bottom panel). YFP-tagged Cav-3P104L preferentially accumulates around nuclei (*), as shown in this representative transfected fibre in which almost no endogenous Cav-3 is revealed by anti-Cav-3 antibody immunolabelling. Conversely, immunolabelling clearly revealed endogenous Cav-3 in the nearby non-transfected fibre. YFP-tagged wild-type Cav-3 appears concentrated at the surface plasma membrane as well as in discrete intracellular clusters, where it is also detected by immunolabelling. Scale bar, 20 μm, applies to all images. B, representative ICa,L recordings, elicited by applying 500 ms duration voltage pulses from a holding potential of −80 mV to a potential of +30 mV. C, left panel, mean current–voltage relationships established for the current in 31 control, eight Cav-3P104L and 16 wild-type Cav-3-expressing fetal fibres. The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details). The right panel presents the mean normalized conductance–voltage relationship in control (n = 31) and Cav-3P104L-expressing (n = 8) fetal fibres.

Over-expression of wild-type Cav-3 in fetal skeletal muscle fibres does not affect ICa,L

We also tested the effect of over-expression of wild-type Cav-3 on the functional expression of the Ca2+ channels. For this, expression of a YFP-tagged wild-type Cav-3 was achieved after transfection of fetal muscle fibres in survival medium. Figure 5A (bottom panel) shows that under these conditions YFP-tagged wild-type Cav-3 appears concentrated at the surface plasma membrane as well as in discrete intracellular clusters presumably corresponding to T-tubules. The measured ICa,L yielded similar properties in Cav-3 over-expressing fibres and in control ones (Fig. 5B and C). Mean values for the parameters fitted from the individual current–voltage curves in Cav-3 over-expressing fibres for Gmax, V0.5, Vrev and k were 188 ± 16 S F−1, 8 ± 1.6 mV, 72 ± 1.4 mV and 4 ± 0.3 mV, respectively (n = 16). The only difference as compared to the control fibres was a slight 5 mV shift of Vrev towards negative potentials (P = 0.01).

Discussion

The present study demonstrates that expression of the LGMD-1C-associated Cav-3P104L mutant in skeletal muscle cells induces a very significant alteration of both the ICa,L and the cellular distribution of the corresponding channels in mouse skeletal muscle cells. Altogether, our results strongly suggest that a marked alteration of the function of the L-type voltage-dependent Ca2+ channel results from Cav-3 deficiency in skeletal muscle. Such a deficiency, revealed by an almost complete loss of immunolabelling, was induced here by the expression of Cav-3P104L mutant in transfected primary cultured myotubes and fetal fibres, thereby reproducing the pathological loss of caveolin observed in skeletal muscle of LGMD-1C patients and transgenic mice bearing the P104L mutation (Minetti et al. 1998; Sunada et al. 2001). Upon expression of Cav-3P104L in cultured myotubes and fetal fibres, mutant as well as endogenous Cav-3 did not reach the sarcolemma and we observed a strong intracellular accumulation of Cav-3P104L, presumably at the level of the Golgi apparatus, consistent with previous observations in fibroblasts, C2C12 and LGMD-1C biopsies (Minetti et al. 1998; Galbiati et al. 1999b; Carozzi et al. 2002).

The reduction in ICa,L amplitude in Cav3P104L-expressing myotubes (55%) was accompanied by a similar reduction in the Cav1.1-labelling intensity (66%). It is thus likely that the reduced ICa,L mainly results from a reduced number of channels rather than from an alteration of the functional properties of the channel. It was recently suggested that Cav-3 has a role in trafficking since its down-regulation was shown to lead to Golgi accumulation of dysferlin (Hernandez-Deviez et al. 2006). Trafficking and/or membrane anchoring of channels such as store-operated Ca2+ channels were moreover found to be dependent on caveolin-1 (Brazer et al. 2003). In this respect, the partial Golgi redistribution of the Cav1.1 labelling and its marked reduction upon Cav-3P104L expression suggest that Cav-3 may also act as a post-Golgi trafficking partner for the DHPR. Cav-3 could exert such a role from the earliest stages of muscle differentiation as both proteins display similar temporal appearance patterns, as shown in cultured muscle cells. Several studies have indeed demonstrated that the mRNA and corresponding L-type voltage-dependent Ca2+ channel protein appear at a detectable level as myoblasts fuse into myotubes (Romey et al. 1989; Varadi et al. 1989; Cognard et al. 1993; Kyselovic et al. 1994); likewise, Cav-3 expression was reported to be strongly induced upon differentiation of cultured mouse skeletal muscle cells (Way & Parton, 1995; Song et al. 1996; Tang et al. 1996; Parton et al. 1997).

Cav-3 is also believed to participate in biogenesis of skeletal muscle T-tubules. This hypothesis first originated from ultrastuctural and immunofluorescence studies (Ishikawa, 1968; Franzini-Armstrong, 1991; Parton et al. 1997; Ralston & Ploug, 1999). Furthermore, skeletal muscles from both CAV-3 knock-out mice and Cav-3P104L-expressing LGMD-1C patients were reported to harbour T-tubule structural abnormalities, although not identical ones (Galbiati et al. 2001; Minetti et al. 2002). Although Cav-3 does not appear to be indispensable for the formation of the T-tubule network, these data suggest that Cav-3 may play a role in its proper organization. It is thus possible that the observed impairment of the DHPR Ca2+ channel function induced by Cav3P104L expression could result from an altered T-tubule biogenesis. In order to test this, the mean electrical capacitance, which is a reliable index of the total membrane surface, was determined and found to be not significantly altered in Cav-3P104L-expressing myotubes. This indicates that Cav-3P104L expression did not alter the amount of the electrically active T-tubular membrane but rather specifically targeted the DHPR. This was further confirmed by results obtained in fetal muscle fibres in which a comparable decrease in maximal conductance of the L-type Ca2+ channel was observed upon Cav-3P104L expression. Under these conditions, and in contrast to cultured myotubes, Cav-3P104L expression was achieved at a maturation stage at which well-formed T-tubules are already present (Takekura et al. 2001) so that an effect related to the disruption of T-tubules organization could hardly be raised. Besides, membrane capacitance was also found to be unchanged in Cav-3P104L-expressing fetal fibres. Furthermore, the lack of effect of Cav-3P104L expression on ICa,T in primary cultured myotubes provides a critical internal control in this set of experiments and further argues in favour of a selective effect of the loss of Cav-3 on the DHPR.

Our immunolabelling experiments show that Cav-3 and DHPR proteins colocalize at both the surface and T-tubular membranes, in agreement with previous studies (Jorgensen et al. 1989; Parton et al. 1997; Ralston & Ploug, 1999). However, it is not entirely clear whether Cav-3 directly interacts physically with DHPR and/or indirectly serves to organize ion channel regulating molecules or the membrane environment for proper ion channel targeting. A number of other ion channels have been shown to be specifically targeted to caveolin-rich microdomains in different tissues (Martens et al. 2004). In cardiac muscle, Cav-3 has been shown to co-immunoprecipitate with the cardiac isoform of the pore-forming subunit of the L-type Ca2+ channel and to play a role in the β2-adrenergic response (Balijepalli et al. 2006). In endo-thelial cells, the Ca2+-activated K+ channel was found to directly interact with caveolin-1 (Wang et al. 2005) and caveolin-1 was also found to bind to the store-operated Ca2+ TRPC1 channels (Brazer et al. 2003). In our study, no reliable quantitative comparison of the Cav1.1 labelling between control and Cav-3P104L-expressing fetal fibres could be achieved, because of the much lower transfection efficiency in this system. We thus cannot rule out the possibility that, in this cell system, the observed reduction in L-type Ca2+ channel conductance would result from mechanism(s) other than decreased expression, membrane mistargeting or anchoring of the channel protein. Along this line, Western blot analysis showed that the amount of Cav1.1 was unchanged in fully matured Cav-3-null mouse muscles (Galbiati et al. 2001). Hence these data suggest that, unlike in myotubes, the decrease of Ca2+ channel conductance in the more differentiated fetal fibres does not result from a reduced density of the protein but rather is primarily due to an altered function of the L-type Ca2+ channel associated with the loss of Cav-3. In agreement with this hypothesis, we found that the voltage dependence of the Ca2+ current was significantly shifted towards positive membrane potentials in Cav-3P104L-expressing fetal fibres but not in myotubes. This result thus strengthens the possibility that the reduced amplitude of ICa,L in Cav-3P104L-expressing cells mainly results from a disruption of Ca2+ channel function in fetal fibres but from a reduced number of channels in myotubes. Finally, our observation that Cav-3 over-expression in fetal fibres did not have any drastic effect on ICa,L may indicate that, under physiological conditions, the available amount of endogenous Cav-3 already allows optimal operating of the L-type Ca2+ channel.

Our results provide important clues to explain the physio-pathological consequences of Cav-3 deficiency in skeletal muscle. A reduction of function of the DHPR may be associated with muscle weakness since it is directly involved in excitation–contraction coupling (Rios & Pizarro, 1991). This may account for the muscle weakness which characterizes LGMD-1C (Woodman et al. 2004). The disease is also associated with progressive muscle wasting (Woodman et al. 2004), and, as suspected for a number of muscular dystrophies (Rizzuto & Pozzan, 2003), chronic intracellular Ca2+ overload may act in this case as the primary process of necrosis. Along this line, transgenic mice expressing Cav-3P104L were shown to over-produce NO (Sunada et al. 2001) which in turn causes a redox-sensitive continuous leak of Ca2+ from the sarcoplasmic reticulum (Pouvreau et al. 2004a). In this way, the reduction in ICa,L possibly together with a reduction in the voltage-controlled calcium release in Cav-3 deficient muscle may be seen as a compensatory or protective mechanism to preserve normal Ca2+ levels.

Acknowledgments

We are very grateful to Dr R.G. Parton (University of Queensland, Australia) for the gift of the wild-type and P104L caveolin cDNAs and to Dr L.E. Espinosa (University of Marseille, France) for his expert advice and help regarding image analysis. This work was supported by grants from Centre National de Recherche Scientifique (CNRS), Agence Nationale de la Recherche, Université Lyon 1 and Association Française contre les Myopathies.

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