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Originally published In Press as doi:10.1074/jbc.M405054200 on August 18, 2004

J. Biol. Chem., Vol. 279, Issue 45, 47092-47100, November 5, 2004
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Involvement of Inositol 1,4,5-Trisphosphate in Nicotinic Calcium Responses in Dystrophic Myotubes Assessed by Near-plasma Membrane Calcium Measurement*

Olivier Basset{ddagger}, François-Xavier Boittin{ddagger}, Olivier M. Dorchies{ddagger}, Jean-Yves Chatton§, Cornelis van Breemen¶, and Urs T. Ruegg{ddagger}||

From the {ddagger}Pharmacology Laboratory, School of Pharmacy, University of Lausanne-Geneva, 1211 Geneva, Switzerland, the §Department of Physiology and Cellular Imaging Facility, University of Lausanne, 1005 Lausanne, Switzerland, and the Department of Pharmacology and Therapeutics, University of British Columbia, Vancouver, British Columbia V6T 1Z1, Canada

Received for publication, May 6, 2004 , and in revised form, August 17, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In skeletal muscle cells, plasma membrane depolarization causes a rapid calcium release from the sarcoplasmic reticulum through ryanodine receptors triggering contraction. In Duchenne muscular dystrophy (DMD), a lethal disease that is caused by the lack of the cytoskeletal protein dystrophin, the cytosolic calcium concentration is known to be increased, and this increase may lead to cell necrosis. Here, we used myotubes derived from control and mdx mice, the murine model of DMD, to study the calcium responses induced by nicotinic acetylcholine receptor stimulation. The photoprotein aequorin was expressed in the cytosol or targeted to the plasma membrane as a fusion protein with the synaptosome-associated protein SNAP-25, thus allowing calcium measurements in a restricted area localized just below the plasma membrane. The carbachol-induced calcium responses were 4.5 times bigger in dystrophic myotubes than in control myotubes. Moreover, in dystrophic myotubes the carbachol-mediated calcium responses measured in the subsarcolemmal area were at least 10 times bigger than in the bulk cytosol. The initial calcium responses were due to calcium influx into the cells followed by a fast refilling/release phase from the sarcoplasmic reticulum. In addition and unexpectedly, the inositol 1,4,5-trisphosphate receptor pathway was involved in these calcium signals only in the dystrophic myotubes. This surprising involvement of this calcium release channel in the excitation-contraction coupling could open new ways for understanding exercise-induced calcium increases and downstream muscle degeneration in mdx mice and, therefore, in DMD.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Duchenne muscular dystrophy (DMD)1 is an X-linked disease that affects about 1 in 3500 males. DMD results in progressive muscle degeneration (1) that ultimately leads to premature death by respiratory or cardiac failure during the third decade. Pharmacological interventions such as glucocorticoids improve the quality of life of patients (2). DMD is caused by the absence of dystrophin, a 427-kDa protein localized under the plasma membrane (3). In normal skeletal muscle fibers, dystrophin is associated with a glycoprotein complex and provides a linkage between the extracellular matrix and the cytoskeleton. This complex tends to stabilize the plasma membrane (4). Numerous studies have shown that the absence of dystrophin in DMD impairs the stability of the plasma membrane, resulting in a greater fragility toward mechanical stress and increased permeability to calcium (Ca2+) (5–9). Indeed, it has been proposed that an alteration of Ca2+ homeostasis might be responsible for the muscle degeneration that occurs in muscle fibers from DMD patients or in those of the mouse model of DMD, the mdx mouse (10–12). Elevations of cytosolic or nearplasma membrane Ca2+ concentrations in myotubes and skeletal muscle fibers from DMD patients or mdx mice have been reported (13, 14). How this increased entry of Ca2+ affects the local concentration of Ca2+ in subcellular compartments and whether this process is involved in the development of the disease is still unclear (15). However it has already been suggested that altered Ca2+ homeostasis and fiber degeneration in mdx mice is "use-dependent" (16–18). In skeletal muscle cells and myotubes, it has been shown that activation of nicotinic receptors leads to plasma membrane depolarization, which triggers a voltage-gated Ca2+ channel opening. Contraction is then triggered by the release of Ca2+ from the sarcoplasmic reticulum through the opening of ryanodine receptors, which are activated by voltage-gated Ca2+ channels (19). However, although the role of ryanodine receptors and voltage-gated Ca2+ channels are well established in excitation-contraction coupling, the skeletal muscle sarcoplasmic reticulum possesses another type of Ca2+ release channel, IP3 receptors (20–22). In many cells, increases in intracellular Ca2+ are mediated by the inositol 1,4,5-trisphoshate pathway in the following manner. The activation of numerous G-protein-coupled receptors triggers hydrolysis of phosphatidylinositol 4,5-bisphosphate to generate IP3, which triggers the opening of IP3 receptors, and intracellular Ca2+ increases (23). Furthermore, IP3 receptors may also be involved in the control of Ca2+ influx through plasma membrane store-operated Ca2+ channels (24–27).

In skeletal muscle cells, IP3 receptors are localized in the nuclear envelope as well as in the sarcoplasmic reticulum (20, 22). In addition, it has been suggested that the intracellular messenger IP3 may contribute to excitation-contraction coupling (28). Indeed, it has been shown that IP3 can potentiate depolarization-induced Ca2+ release or produce the contraction of intact or skinned skeletal muscle fibers (29–31). These results suggest that IP3 may play a role in excitation-contraction coupling in skeletal muscle cells. Furthermore, it has also been shown that the plasma membrane depolarization that activates voltage-gated Ca2+ channels can induce IP3 production and IP3 receptor activation in myotubes with a mechanism involving a G-protein (20, 32, 33).

The mechanisms involved in Ca2+ responses during physiological stimulation (using nicotinic receptor agonists) have not been studied in dystrophic myotubes to date. Here, we have studied the IP3 receptor pathway in dystrophic myotubes using plasma membrane-targeted aequorin (SNAP-25 aequorin) for measuring the near-plasma membrane Ca2+ concentration. We show here that SNAP-25 aequorin is a reliable tool to measure Ca2+ increases occurring just below the plasma membrane. We also show that nicotinic receptor stimulation triggers nearplasma membrane Ca2+ increases that depend on IP3 receptor activation in myotubes derived from mdx mice only, thus showing a non-common pathway for excitation-contraction coupling.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture—Cultures of purified myoblasts were prepared in Petri dishes (Falcon; BD Sciences) and maintained at 37 °C in a water-saturated atmosphere of 95% air and 5% CO2.

The cell cultures were obtained as described previously (34) with minor modifications. Briefly, 3-week-old mdx C57Bl/10 and control C57Bl/10 mice were killed by cervical dislocation, extensor digitorum longus (EDL) muscle was aseptically removed bilaterally, cleaned of its tendinous ends and adhering connective tissue, and minced into small pieces of ~1 mm3. Individual muscles from two mice were pooled. After the muscle tissues were rinsed in DMEM (Invitrogen), digestion was performed by three successive rounds of incubation in 10 ml of DMEM supplemented with 10 mM HEPES (pH 7.4), 10% fetal calf serum (Invitrogen), and 0.15% Pronase (Roche Applied Science) at 37 °C. After the first incubation (5 min), the tissues were triturated with a serologic pipette, and the supernatant, containing mainly blood cells and connective debris, was discarded. The second and third incubations lasted 20 min each with frequent triturations. The supernatants were collected, pooled, filtered through a 40-µm mesh cell strainer (Falcon; BD Biosciences), and centrifuged at 300 x g for 10 min. The cell pellets were resuspended in growth medium (see below), and the suspensions were plated onto collagen type I-coated (1 µg/cm2; Sigma) 100-mm diameter Petri dishes, one dish per original muscle. Growth medium was composed of a 1:1 (v/v) mixture of DMEM and MCDB202 (CryoBioSystem) supplemented with 20% fetal calf serum, 2% Ultroser® SF (Biosepra SA), antibiotics (Ciproxin; Bayer), and NaHCO3 from concentrated solution (Invitrogen) to get 2.6 g/liter in complete medium. Myoblasts were plated on coated dishes and grown for ~2 days in growth medium. For the experiments, cells were transfected (see below) and induced to fuse by changing the growth medium to differentiation medium (DMEM supplemented with 1.7% fetal calf serum, 3.3% horse serum (Sigma), 10 µg/ml insulin (Fluka), and Ciproxin).

Transfection—Myoblasts were plated at 12,000 cells/cm2 on 13-mm Thermanox coverslips (Nalge Nunc International) in four-well plates. When 80–90% confluent, growth medium was removed and replaced with a serum-free medium, Opti-MEM I (Invitrogen). Cells were transfected overnight using LipofectAMINE 2000 (Invitrogen) at a ratio of 1 µg of DNA per 2 µl of transfection reagent. The DNA-LipofectAMINE 2000 complex was prepared in Opti-MEM I medium. After overnight incubation, this medium was replaced by a differentiation medium. Myotubes were used 3 or 4 days after differentiation.

Plasmids—The aequorin plasmids were gifts from T. Pozzan, Padova, Italy. Cells were transfected with a pcDNAI expression vector containing a cDNA encoding either green fluorescent protein (GFP) for localization or wild type aequorin for Ca2+ measurement, both fused with the SNAP-25 targeting sequence to measure subsarcolemmal calcium (pm[Ca2+]) (35) or with cytosolic aequorin to measure cytosolic calcium concentrations ([Ca2+]c) (13). The IP3 sponge plasmid was a gift from H. L. Roderick and M. D. Bootman, Cambridge, United Kingdom. Cells were co-transfected with a pdc515 expression vector (Microbix Biosystems Inc.) containing a cDNA encoding enhanced GFP, the high affinity IP3 sponge (36), and a pcDNAI expression vector containing a cDNA encoding the SNAP-25 aequorin.

Confocal Microscopy—Mdx myotubes were grown in plastic culture dishes and transfected with a pcDNAI expression vector containing the cDNA encoding a GFP-tagged SNAP-25 targeting sequence. Confocal imaging was performed on a living myotube on a LSM 510 Meta confocal scanner mounted on an upright Axioskop 2 FS microscope (Carl Zeiss). Fluorescence was excited at 488 nm, and emission was detected at 505–530 nm using a 40 x 0.8 numerical aperture water immersion objective (Achroplan, Carl Zeiss). Simultaneous differential interference contrast images of cells were recorded on the transmitted light detector with the polarized 488-nm light used for GFP fluorescence excitation.

Intracellular Calcium Measurement—After 3 or 4 days of differentiation, subsarcolemmal Ca2+ concentration was determined in a population of myotubes as described previously with minor modifications (37). Briefly, the SNAP-25 aequorin was reconstituted in a Ca2+-free physiological salt solution (PSS) (145 mM NaCl, 5 mM KCl, 1 mM MgCl2, 5 mM HEPES, and 10 mM glucose, pH 7.6) containing 0.1 mM EGTA and 5 µM coelenterazine (Calbiochem) for 1 h before the experiment to decrease the [Ca2+] within the cell. Cells were superfused at a rate of 1 ml/min in a custom-made 0.5-ml chamber thermostated at 37 °C (MecaTest, Geneva, Switzerland). Emitted luminescence was detected at 466 nm with a photomultiplier apparatus (EMI 9789A, Electron Tubes Limited, United Kingdom) and recorded every second using a computer photon-counting board (EMI C660) as described previously (38). The relationship between recorded counts and [Ca2+] is shown in Equation 1 (39),

(Eq. 1)
where L represents the recorded photons/s and Lmax represents the remaining photons that correspond to the total light output during the whole experiment minus the photons emitted up to the measured point. KTR and KR are the parameters of the aequorin and n is the number of Ca2+ binding sites. At 37 °C, KTR = 120, KR = 1.01E + 07 M-1, and n = 2.99. Total light output was obtained by exposing cells to 10 mM CaCl2 after permeabilization with 100 µM digitonin to consume all the aequorin.

45Ca2+ Influx Measurement—Influx measurement was performed using 45Ca2+ (5–50 mCi/mg; PerkinElmer Life Sciences) as described with minor modifications (40). Briefly, myotubes were washed twice and preincubated at 37 °C for 10 min in 250 µl of the PSS solution described above also containing 0.12 mM CaCl2 and, eventually, the indicated inhibitor. 45Ca2+ influx was initiated by incubation with 200 µl of PSS containing 1.2 mM CaCl2 alone or with 100 µM carbachol (CCh) and 0.4 mCi of 45Ca2+ for 10 min at 37 °C. The cells were subsequently washed four times with ice-cold PSS containing no CaCl2 but 0.1 mM EGTA (to remove extracellularly bound 45Ca2+). Cells were lysed with 250 µl of 1% SDS (w/v). The radioactivity of the lysate was measured by scintillation counting (Packard 460C, Zurich, Switzerland). Values are expressed in counts per minute.

Data Analysis—Data analysis was performed using the software GraphPad Prism (GraphPad Software, San Diego, CA) and Matlab (The MathWorks Inc, Natick, MA). Results are expressed as means ± S.E. Statistical significance between the different values was assessed with the unpaired Student's t test or one-way analysis of variance test followed by a Dunnett test. p > 0.05 was considered not significant; p values ≤ 0.05 were considered significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
As the removal of intracellular Ca2+ is required for the complete reconstitution of aequorin (41), SNAP-25 aequorin-expressing cells were preincubated in Ca2+-free PSS containing 0.1 mM EGTA for 1 h before performing the experiment. For each experiment the basal level of luminescence was recorded for 1 min before performing the experiment. The stimulus applied was PSS containing 1.2 mM Ca2+ and the test compounds.

SNAP Aequorin Localization—To investigate the localization of the SNAP-25 aequorin, cells were transfected with cDNA encoding the GFP fused with the SNAP-25 targeting sequence. Fig. 1 shows a representative confocal section of a transfected living mdx myotube after 3 days of differentiation. SNAP-25-GFP-mediated fluorescence (Fig. 1A) appeared to be mainly localized at the sarcolemma when compared with the localization assessed with differential interference contrast imaging (Fig. 1, B and C). However, some SNAP-25-GFP fluorescence was detected in the cytosol, which could correspond to a non-palmitoylated protein (35). Similar results were obtained with control C57 myotubes (data not shown).



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  		FIG. 1.
SNAP-25 aequorin localization in mdx myotubes. Shown is a confocal section of a living mdx myotube expressing the GFP protein tagged with the SNAP-25 targeting sequence, indicating SNAP-25 aequorin localization. A, GFP fluorescence. B, differential interference contrast. C, superposition of panels A and B.

 
To further investigate the localization of the SNAP-25 aequorin, cells were incubated for 10 min with the high affinity Ca2+ chelator BAPTA-AM to buffer cytosolic Ca2+. Cytosolic Ca2+ responses assessed with cytosolic aequorin triggered by CCh and Ca2+ readdition were completely inhibited in cells loaded with BAPTA-AM (Fig. 2A). In contrast, in cells transfected with SNAP-25 aequorin the maximal Ca2+ responses and time to peak values (17.50 ± 2.0 s and 20.17 ± 2.7 s) were not significantly reduced by BAPTA-AM (Fig. 2B). These results indicate that Ca2+ responses measured with SNAP-25 aequorin were due to Ca2+ increases in a restricted area under the plasma membrane, presumably where BAPTA-AM had very limited access. Moreover, as shown in Fig. 2C, the Ca2+ peaks triggered by these stimuli were 10 times larger when measured with SNAP-25 aequorin than with cytosolic aequorin (4.97 ± 0.66 and 0.53 ± 0.04 µM, respectively). Altogether, these results indicate that SNAP-25 aequorin was mainly localized at the plasma membrane. Therefore the probe appears to be a reliable tool for measuring near-plasma membrane Ca2+ concentrations.



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  		FIG. 2.
BAPTA-AM effect on the Ca2+ response. A and B, representative traces of the global Ca2+ concentration in a population of EDL mdx myotubes showing the effect of a 10 µM BAPTA-AM preincubation compared with non-treated myotubes. Confluent myoblasts were transfected with cytosolic aequorin (A) and SNAP-25 aequorin (B). Myotubes were used at days 3 or 4 after differentiation. Arrows indicate the beginning of stimulation with CCh. C, summarizing histogram; a cross indicates myoblasts transfected with SNAP-25 aequorin. Cytosolic Ca2+ response with BAPTA-AM is not shown because of complete abolition. The bar graphs correspond to mean [Ca2+] values ± S.E. (n ≥ 6; ns, not significant (p > 0.05); *** corresponds to a significant inhibition (p < 0.001) using an unpaired Student's t test).

 
CCh-induced Ca2+ Responses in Control and Dystrophic Cells—To test the hypothesis that the IP3 receptor pathway could be involved in acetylcholine receptor activation, we measured the effects of the nicotinic agonist carbachol (100 µM) and the effects of the potent IP3 receptor inhibitor, 2-aminoethoxydiphenyl borate (2-APB) (42), on subsarcolemmal Ca2+ transients in both control and dystrophic myotubes. As shown in Fig. 3, 100 µM CCh together with 1.2 mM Ca2+ triggered peak pm[Ca2+] increases of 1.06 ± 0.11 µM for control cells and 4.97 ± 0.66 µM for dystrophic cells, representing a 4.5-fold increase. When cells were incubated for 10 min with 75 µM 2-APB, the CCh-induced Ca2+ response was not significantly changed in control myotubes but was significantly decreased in dystrophic myotubes (from 4.97 ± 0.66 to 1.63 ± 0.21 µM). These results therefore suggest that the Ca2+-release pathways involved in CCh responses are different between control and dystrophic myotubes and that the IP3 pathway may be involved in CCh-induced Ca2+ responses. We then investigated the pathway involved in dystrophic myotubes.



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  		FIG. 3.
CCh-induced Ca2+ response in control and dystrophic cells. Shown is a summarizing histogram showing the effect of 1.2 mM Ca2+ readdition together with 100 µM CCh and 75 µM 2-APB in control C57 and dystrophic myotubes. The bar graphs correspond to mean pm[Ca2+] values ± S.E. (n ≥ 4; ns, not significant (p > 0.05); *** corresponds to a significant inhibition (p < 0.001) using an unpaired Student's t test). Plain columns represent results from the control cells; hatched columns represent results from the dystrophic cells.

 
Ca2+ Increases Triggered by CCh Are Mediated by Nicotinic Receptors—The application of CCh together with Ca2+ triggered pm[Ca2+] increases of a maximal amplitude of 4.97 ± 0.66 µM. As shown in Fig. 4, C and E, preincubation of cells for 10 min with atropine (100 µM), a selective muscarinic receptor antagonist, did not affect the maximal amplitude of the CCh-induced Ca2+ peak. On the other hand, 10 min of incubation with the highly selective nicotinic receptor antagonist d-tubocurarine (10 µM) decreased CCh-induced Ca2+ responses from 4.97 ± 0.66 µM in non-treated cells to 1.47 ± 0.03 µM in the presence of the compound (Fig. 4, D and E). Average CCh responses in the presence of d-tubocurarine were not statistically different from the Ca2+ responses triggered by Ca2+ readdition alone. Altogether, these results indicate that CCh-induced Ca2+ increases were mediated by the activation of nicotinic receptors.



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  		FIG. 4.
Nicotinic receptors are involved in the CCh-induced response. A–D, representative traces of subsarcolemmal Ca2+ increases in a population of EDL-derived mdx myotubes showing the effect of 1.2 mM Ca2+ readdition (A), 1.2 mM Ca2+ readdition together with 100 µM CCh (B), 100 µM atropine (C), and 10 µM d-tubocurarine (D). Arrows indicate the beginning of stimulation with CCh. E, summarizing histogram; the bar graphs correspond to mean pm[Ca2+] values ± S.E. (n ≥ 6; ns, not significant (p > 0.05); * and ** correspond to a significant inhibition (p < 0.05 and 0.01 and p < 0.001, respectively) using a one-way analysis of variance test followed by a Dunnett test to compare all of the columns with the 1.2 mM Ca2+ and 100 µM CCh bar).

 
Initial Ca2+ Influx Induced by CCh—Ca2+ increases triggered by CCh may rely on Ca2+ influx through plasma membrane channels. Fig. 5A shows that CCh-induced Ca2+ increases were triggered by an influx through plasma membrane Ca2+ channels. Indeed, incubation for 10 min with 1 mM cadmium, a non-selective Ca2+ channel inhibitor (43), drastically decreased the CCh-induced Ca2+ response (from 4.97 ± 0.66 µM for non-treated cells to 0.46 ± 0.02 µM for treated cells). Moreover, CCh had no effect on the pm[Ca2+] when added to a Ca2+-free solution, whereas the addition of CCh together with Ca2+ led to a Ca2+ response (Fig. 5B). These results strongly suggest that the Ca2+ responses were triggered at least in part by Ca2+ influx through plasma membrane Ca2+ channels.



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  		FIG. 5.
Initial Ca2+ influx induced by CCh. A, histogram showing the effect of cadmium (1 mM) on the CCh-induced Ca2+ response (n ≥ 6; *** indicates significant inhibition (p < 0.001) using unpaired Student's t test). B, representative trace of a subsarcolemmal Ca2+ response induced by 100 µM CCh in a population of EDL-derived mdx myotubes showing the effects of a Ca2+ omission and the readdition of 1.2 mM Ca2+. C, histogram illustrating the effect of nifedipine (1 µM) on subsarcolemmal Ca2+ response induced by 100 µM CCh (n ≥ 6; * indicates significant inhibition (p < 0.05) using an unpaired Student's t test). D, histogram showing the effect of 100 µM CCh and 1 µM nifedipine on 45Ca2+ influx. The bar graph corresponds to the means of 45Ca2+ influx values ± S.E. (n = 8; *** corresponds to a significant difference (p < 0.001) using an unpaired Student's t test).

 
We then tested the effect of nifedipine (1 µM), a potent and selective blocker of L-type voltage-gated Ca2+ channels, on the CCh-induced Ca2+ response. Results indicated that L-type voltage-gated Ca2+ channel activation was involved in the response triggered by CCh (Fig. 5C). To confirm that this response was truly due to influx, we used the 45Ca2+ influx technique. As shown in Fig. 5D, Ca2+ influx was increased from 638 ± 51 cpm/well with Ca2+ alone to 910 ± 69 cpm/well with 100 µM CCh. Moreover, when cells were incubated with 1 µM nifedipine and both Ca2+ and CCh, the Ca2+ influx was decreased (591 ± 13 cpm/well). These results confirm that CCh triggered Ca2+ influx through voltage-gated L-type Ca2+ channels.

Quick Sarcoplasmic Reticulum Ca2+ Refilling and Ca2+ Release—Results from experiments described above have shown that in dystrophic myotubes CCh-induced Ca2+ responses were at least in part due to Ca2+ influx. To go further, we tested the effect of the sarcoplasmic-endoplasmic reticulum Ca2+ ATPase (SERCA) blocker thapsigargin (44). As shown in Fig. 6, incubation for 10 min with 1 µM thapsigargin decreased the CCh-induced Ca2+ response from 4.97 ± 0.66 to 1.92 ± 0.24 µM for the thapsigargin-treated cells, which is very similar to the response triggered by Ca2+ alone (1.84 ± 4.4 µM). Altogether these results show that the CCh-induced Ca2+ transients are due to both Ca2+ influx and Ca2+ release from the sarcoplasmic reticulum.



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  		FIG. 6.
Quick sarcoplasmic reticulum Ca2+-refilling and Ca2+-release. A, representative trace of subsarcolemmal Ca2+ response in a population of EDL mdx myotubes showing the effect of 1 µM thapsigargin on the CCh-induced Ca2+ response. Arrow indicates the beginning of stimulation with CCh. B, summarizing histogram. The bar graphs correspond to mean of pm[Ca2+] values ± S.E. (n = 12; ns, not significant (p > 0.05); *** corresponds to a significant difference (p < 0.001) using an unpaired Student's t test).

 
Involvement of the IP3 Pathway in CCh-induced Response— Results shown in Fig. 3 suggest that IP3 pathway may be involved in CCh-induced Ca2+ response in dystrophic myotubes. To test the involvement of this pathway, we investigated the effect of U73122 [GenBank] , a potent phospholipase C (PLC) inhibitor (45). Indeed, PLC is required for IP3 formation through phosphatidylinositol 4,5-bisphosphate cleavage. As shown in Fig. 7, incubation of the cells with 20 µM U73122 [GenBank] decreased the CCh-induced Ca2+ peak amplitude from 4.97 ± 0.66 µM for non-treated cells to 2.76 ± 0.19 µM for treated cells. This result suggests that the PLC/IP3 pathway may be involved in the CCh-mediated Ca2+ response. Moreover, we tested the effect of xestospongin D, a potent blocker of IP3 receptors, at a very low concentration (0.1 µM) (46). Preincubation of the cells for 10 min with this compound decreased the CCh-induced Ca2+ responses from 4.97 ± 0.66 to 2.44 ± 0.34 µM.



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  		FIG. 7.
Involvement of the IP3 pathway in CCh-induced Ca2+ response. A–D, representative traces of subsarcolemmal Ca2+ response in a population of EDL mdx myotubes showing the effect of 20 µM U73122 [GenBank] (A), 75 µM 2-APB (B), 0.1 µM xestospongin D (Xesto D) (C), and an IP3 sponge co-transfected with SNAP-25 aequorin (D). Arrows indicate beginning of stimulation with CCh in the presence of the compounds indicated. E, summarizing histogram. The bar graphs correspond to mean pm[Ca2+] values ± S.E. (n ≥ 6; *** corresponds to a significant inhibition (p < 0.001) as assessed by a one-way analysis of variance test followed by a Dunnett test to compare all the column with the 1.2 mM Ca2+ and 100 µM CCh bar).

 
Because 2-APB and xestospongin D may have nonspecific effects (47, 48) we co-transfected cells with both a high affinity IP3 sponge and SNAP-25 aequorin. This IP3 sponge has been shown to behave as an IP3 chelator and to inhibit IP3-mediated Ca2+ responses (36). Again, the CCh-induced Ca2+ peak was strongly decreased in co-transfected cells (from 4.97 ± 0.66 to 0.72 ± 0.13 µM) and was below that of the response due to Ca2+ alone. Altogether, these results clearly show that the IP3 pathway was involved in CCh-induced Ca2+ response in our mdx myotubes.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study we used SNAP-25, a plasma membrane protein, to target the Ca2+ sensitive photoprotein aequorin to the subsarcolemmal area of both control and dystrophic myotubes. Because of the complexity (different degrees of maturation) and density (heavy cytoskeleton, actin, and myosin) of skeletal muscle cells, we first checked the localization of the probe. This was investigated by confocal microscopy using SNAP-25-GFP on living mdx myotubes. Results suggest that this probe and, therefore, also SNAP-25 aequorin was preferentially localized at the sarcolemma. Confocal sections of mdx myotubes also showed some cytosolic fluorescence that corresponds to non-targeted protein or, most probably, non-palmitoylated SNAP-25. Indeed, working with live cells indicates that proteins are continuously expressed, and it has been shown previously that the SNAP-25 protein must be palmitoylated on cysteine residues to be targeted correctly to the inner leaflet of the plasma membrane (35). In our case, the cytosolic fluorescence is likely to be due to non-mature SNAP-25-GFP protein.

Localization of SNAP-25 aequorin was also assessed with functional experiments. Indeed, BAPTA-AM pretreatment of the cells did not significantly decrease CCh-induced Ca2+ responses measured with SNAP-25 aequorin but completely suppressed those detected with cytosolic aequorin. The fact that the time needed to reach the peak was also unaltered by BAPTA-AM pretreatment suggests that this finding is due to the inability of BAPTA to chelate Ca2+ in the space below the plasma membrane. These results clearly indicate that SNAP-25 aequorin allows us to measure Ca2+ in a restricted area under the plasma membrane that may correspond to the "fuzzy space" described in other cell types (49, 50). In this restricted area BAPTA-AM appears to have little or no access. These results also confirm that SNAP-25 aequorin was mainly localized at the plasma membrane and that it is a reliable tool for measuring near-plasma membrane Ca2+ concentration in these myotubes. Ca2+ increases in response to CCh were much bigger when measured under the plasma membrane rather than in the cytosol. This is very likely due to the fact that SNAP-25 aequorin is very close to Ca2+ channels and also that Ca2+ increases may occur in a physically restricted area, which prevents diffusion of Ca2+ in the bulk cytosol.

CCh-induced pm[Ca2+] increases were 4.5 times bigger in dystrophic myotubes than in control myotubes. This result is in agreement with previous results obtained in muscle fibers that also showed a subsarcolemmal Ca2+ overload due to enhanced Ca2+ influx in dystrophic fibers as compared with control (14). Moreover, we found that 2-APB, an IP3 receptor inhibitor, strongly decreased the CCh-induced Ca2+ responses in dystrophic but not in control C57 myotubes. These results also suggest that the mechanisms involved in CCh-induced subsarcolemmal increases may be different between control and dystrophic myotubes.

In mdx myotubes, CCh was able to trigger Ca2+ increases only when Ca2+ was added and CCh-induced Ca2+ responses were completely blocked by cadmium ion, a non-selective Ca2+ channel blocker. This result indicates that CCh-induced Ca2+ increases after preincubation in a Ca2+-free solution are triggered by Ca2+ influx through plasma membrane Ca2+ channels. Moreover, CCh-induced Ca2+ responses were partially inhibited by nifedipine, showing that L-type voltage-gated Ca2+ channel activation are involved in CCh-induced Ca2+ responses. When cells were incubated with the sarcoplasmic-endoplasmic reticulum Ca2+ ATPase blocker thapsigargin, the CCh-induced Ca2+ responses were significantly decreased and reached the same amplitude as the response triggered by Ca2+ readdition. Altogether, these results suggest that CCh-induced Ca2+ responses are due to Ca2+ influx through plasma membrane Ca2+ channels followed by a fast Ca2+ refilling of the sarcoplasmic reticulum and a secondary Ca2+ release from the store.

In mdx myotubes, CCh-induced pm[Ca2+] increases were significantly reduced when cells were incubated with two potent IP3 receptor inhibitors (2-APB and xestospongin D) or when cells were co-transfected with an IP3 sponge together with SNAP-25 aequorin. These results demonstrate that IP3 receptors are involved in CCh-induced Ca2+ increases in dystrophic myotubes. Consistently, CCh-induced Ca2+ increases were also inhibited by the phospholipase C inhibitor U73122 [GenBank] . Altogether, these results indicate that nicotinic receptor activation triggers PLC activation and IP3 release in our dystrophic myotubes, raising a question about the mechanism of PLC activation. It has been shown recently that the activation of voltage-gated Ca2+ channels by potassium chloride depolarization can activate PLC in myotubes by a mechanism involving a G-protein (32). This could indeed be the case in our dystrophic myotubes, as CCh triggers activation of L-type voltage-gated Ca2+ channels. PLC may therefore be activated by nearplasma membrane Ca2+ increases due to Ca2+ influx occurring through nicotinic receptors and/or voltage-gated Ca2+ channels (51). Altogether, these results indicate that CCh-induced Ca2+ increases rely on IP3 receptor activation in dystrophic myotubes. On the other hand, 2-APB was without effect on CCh-induced Ca2+ responses in control myotubes, indicating that the IP3 pathway is not involved in nicotinic responses in this cell type.

However, recent studies have shown that 2-APB is also a potent inhibitor of channels activated by the depletion of the Ca2+ stores (store-operated channels or SOCs) (52, 53). Indeed, since our cells were incubated for 1 h in a Ca2+-free solution for complete aequorin reconstitution, Ca2+ stores have been depleted. These conditions raise the question about the involvement of SOCs in the CCh-induced Ca2+ responses. Our results with thapsigargin and IP3 pathway inhibitors strongly suggest that IP3 receptor-induced Ca2+ release is involved in CCh-induced Ca2+ response. However, we can not exclude the involvement of SOCs in CCh responses. Indeed, IP3 receptor activation could trigger SOCs opening, as has already been shown in numerous types of cells including skeletal muscle cells (24, 26, 54). Moreover it has been shown that SOCs, which belong to the family of the transient receptor potential channels (TRPC) (55), could be involved in the increased Ca2+ influx in dystrophic fibers (56).

In conclusion, our results indicate that IP3 receptors are involved in near-plasma membrane Ca2+ increases triggered by the activation of nicotinic receptors in dystrophic myotubes. The involvement of IP3 receptors represents a new and unexpected pathway for the excitation of dystrophic skeletal muscle cells. Indeed, our control myotubes do not display any IP3 inhibitor sensitivity. It has been shown in muscle cells that the main pathway for Ca2+ release from the sarcoplasmic reticulum and muscle contraction is the ryanodine receptor (19). Moreover, increases in IP3 receptor number and a 2- to 3-fold increase in IP3 levels have been reported in mdx myotubes (21). Altogether, these results suggest that IP3 receptors may be involved in altered Ca2+ homeostasis and the subsequent muscle degeneration that occurs in dystrophic muscle cells.


   FOOTNOTES
 
* This work was supported by Swiss National Science Foundation Grant 31-68315.02 and by grants from the Swiss Foundation for Research on Muscular Diseases and the Association Française contre les Myopathies. 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.: 4122-379-3429; Fax: 4122-379-3430; E-mail: Urs.Ruegg{at}pharm.unige.ch.

1 The abbreviations used are: DMD, Duchenne muscular dystrophy; 2-APB, 2-aminoethoxydiphenyl borate; BAPTA-AM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid tetra(acetoxymethyl)ester; CCh, carbachol; DMEM, Dulbecco's modified Eagle's medium; EDL, extensor digitorum longus; GFP, green fluorescent protein; IP3, inositol 1,4,5-trisphosphate; PLC, phospholipase C; pm[Ca2+], subsarcolemmal Ca2+ concentration; PSS, physiological salt solution; SNAP-25, synaptosome-associated protein of 25 kDa. Back


   ACKNOWLEDGMENTS
 
We thank Philippe Lhote for technical help with the cell culture and Drs. H.L. Roderick and M.D. Bootman (Calcium Group, Laboratory of Molecular Signaling, The Babraham Institute, Babraham, Cambridge, United Kingdom) and Professor T. Pozzan (University of Padova, Italy) for their generous gift of plasmids.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Emery, A. E. (1991) Neuromuscul. Disord. 1, 19-29[CrossRef][Medline] [Order article via Infotrieve]
  2. Muntoni, F., Fisher, I., Morgan, J. E., and Abraham, D. (2002) Neuromuscul. Disord. 12, Suppl. 1, S162-S165[CrossRef][Medline] [Order article via Infotrieve]
  3. Ahn, A. H., and Kunkel, L. M. (1993) Nat. Genet. 3, 283-291[CrossRef][Medline] [Order article via Infotrieve]
  4. Fabbrizio, E., Bonet-Kerrache, A., Limas, F., Hugon, G., and Mornet, D. (1995) Biochem. Biophys. Res. Commun. 213, 295-301[CrossRef][Medline] [Order article via Infotrieve]
  5. Gillis, J. M. (1996) Acta Physiol. Scand. 156, 397-406[CrossRef][Medline] [Order article via Infotrieve]
  6. Alderton, J. M., and Steinhardt, R. A. (2000) J. Biol. Chem. 275, 9452-9460[Abstract/Free Full Text]
  7. McCarter, G. C., and Steinhardt, R. A. (2000) J. Membr. Biol. 176, 169-174[CrossRef][Medline] [Order article via Infotrieve]
  8. Tutdibi, O., Brinkmeier, H., Rudel, R., and Fohr, K. J. (1999) J. Physiol. 515, 859-868[Abstract/Free Full Text]
  9. Leijendekker, W. J., Passaquin, A. C., Metzinger, L., and Ruegg, U. T. (1996) Br. J. Pharmacol. 118, 611-616[Medline] [Order article via Infotrieve]
  10. Turner, P. R., Fong, P. Y., Denetclaw, W. F., and Steinhardt, R. A. (1991) J. Cell Biol. 115, 1701-1712[Abstract/Free Full Text]
  11. Gailly, P. (2002) Biochim. Biophys. Acta 1600, 38-44[Medline] [Order article via Infotrieve]
  12. Bulfield, G., Siller, W. G., Wight, P. A., and Moore, K. J. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 1189-1192[Abstract/Free Full Text]
  13. Robert, V., Massimino, M. L., Tosello, V., Marsault, R., Cantini, M., Sorrentino, V., and Pozzan, T. (2001) J. Biol. Chem. 276, 4647-4651[Abstract/Free Full Text]
  14. Mallouk, N., Jacquemond, V., and Allard, B. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 4950-4955[Abstract/Free Full Text]
  15. Ruegg, U. T., and Gillis, J.-M. (1999) Trends Pharmacol. Sci. 20, 351-352[CrossRef][Medline] [Order article via Infotrieve]
  16. Sacco, P., Jones, D. A., Dick, J. R., and Vrbova, G. (1992) Clin. Sci. (Lond.) 82, 227-236[Medline] [Order article via Infotrieve]
  17. Vilquin, J. T., Brussee, V., Asselin, I., Kinoshita, I., Gingras, M., and Tremblay, J. P. (1998) Muscle Nerve 21, 567-576[CrossRef][Medline] [Order article via Infotrieve]
  18. De Luca, A., Pierno, S., Liantonio, A., Cetrone, M., Camerino, C., Fraysse, B., Mirabella, M., Servidei, S., Ruegg, U. T., and Conte Camerino, D. (2003) J. Pharmacol. Exp. Ther. 304, 453-463[Abstract/Free Full Text]
  19. Berchtold, M. W., Brinkmeier, H., and Muntener, M. (2000) Physiol. Rev. 80, 1215-1265[Abstract/Free Full Text]
  20. Powell, J. A., Carrasco, M. A., Adams, D. S., Drouet, B., Rios, J., Muller, M., Estrada, M., and Jaimovich, E. (2001) J. Cell Sci. 114, 3673-3683[Medline] [Order article via Infotrieve]
  21. Liberona, J. L., Powell, J. A., Shenoi, S., Petherbridge, L., Caviedes, R., and Jaimovich, E. (1998) Muscle Nerve 21, 902-909[CrossRef][Medline] [Order article via Infotrieve]
  22. Salanova, M., Priori, G., Barone, V., Intravaia, E., Flucher, B., Ciruela, F., McIlhinney, R. A. J., Parys, J. B., Mikoshiba, K., and Sorrentino, V. (2002) Cell Calcium 32, 193-200[CrossRef][Medline] [Order article via Infotrieve]
  23. Berridge, M. J. (1993) Nature 361, 315-325[CrossRef][Medline] [Order article via Infotrieve]
  24. Ma, H. T., Patterson, R. L., van Rossum, D. B., Birnbaumer, L., Mikoshiba, K., and Gill, D. L. (2000) Science 287, 1647-1651[Abstract/Free Full Text]
  25. Kiselyov, K., Mignery, G. A., Zhu, M. X., and Muallem, S. (1999) Mol. Cell 4, 423-429[CrossRef][Medline] [Order article via Infotrieve]
  26. Kiselyov, K., Xu, X., Mozhayeva, G., Kuo, T., Pessah, I., Mignery, G., Zhu, X., Birnbaumer, L., and Muallem, S. (1998) Nature 396, 478-482[CrossRef][Medline] [Order article via Infotrieve]
  27. Kiselyov, K., and Muallem, S. (1999) Trends Neurosci. 22, 334-337[CrossRef][Medline] [Order article via Infotrieve]
  28. Vergara, J., Tsien, R., and Delay, M. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 6352-6356[Abstract/Free Full Text]
  29. Talon, S., Vallot, O., Huchet-Cadiou, C., Lompre, A.-M., and Leoty, C. (2002) Am. J. Physiol. 282, R1164-R1173
  30. Talon, S., Huchet-Cadiou, C., and Leoty, C. (1999) Pflugers Arch. 438, 804-816[CrossRef][Medline] [Order article via Infotrieve]
  31. Yamaguchi, N., and Kasai, M. (1997) Biochem. Biophys. Res. Commun. 240, 772-777[CrossRef][Medline] [Order article via Infotrieve]
  32. Araya, R., Liberona, J. L., Cardenas, J. C., Riveros, N., Estrada, M., Powell, J. A., Carrasco, M. A., and Jaimovich, E. (2002) J. Gen. Physiol. 121, 3-16[Abstract/Free Full Text]
  33. Jaimovich, E., Reyes, R., Liberona, J. L., and Powell, J. A. (2000) Am. J. Physiol. 278, C998-C1010
  34. Pinset, C., and Montarras, D. (1998) in Cell Biology: A Laboratory Handbook (Celis, J., ed) Vol. 1, 2nd Ed., pp. 226-232, Academic Press, New York
  35. Brini, M., Pinton, P., Pozzan, T., and Rizzuto, R. (1999) Microsc. Res. Tech. 46, 380-389[CrossRef][Medline] [Order article via Infotrieve]
  36. Uchiyama, T., Yoshikawa, F., Hishida, A., Furuichi, T., and Mikoshiba, K. (2002) J. Biol. Chem. 277, 8106-8113[Abstract/Free Full Text]
  37. Challet, C., Maechler, P., Wollheim, C. B., and Ruegg, U. T. (2001) J. Biol. Chem. 276, 3791-3797[Abstract/Free Full Text]
  38. Kennedy, E. D., Rizzuto, R., Theler, J. M., Pralong, W. F., Bastianutto, C., Pozzan, T., and Wollheim, C. B. (1996) J. Clin. Investig. 98, 2524-2538[Medline] [Order article via Infotrieve]
  39. Alvarez, J., and Montero, M. (2002) Cell Calcium 32, 251-260[CrossRef][Medline] [Order article via Infotrieve]
  40. Passaquin, A. C., Lhote, P., and Ruegg, U. T. (1998) Br. J. Pharmacol. 124, 1751-1759[CrossRef][Medline] [Order article via Infotrieve]
  41. Marsault, R., Murgia, M., Pozzan, T., and Rizzuto, R. (1997) EMBO J. 16, 1575-1581[CrossRef][Medline] [Order article via Infotrieve]
  42. Maruyama, T., Kanaji, T., Nakade, S., Kanno, T., and Mikoshiba, K. (1997) J Biochem. 122, 498-505[Abstract/Free Full Text]
  43. Oba, T., Yamaguchi, M., Wang, S., and Johnson, J. D. (1992) Biophys. J. 63, 1416-1420[Medline] [Order article via Infotrieve]
  44. Janczewski, A. M., and Lakatta, E. G. (1993) Am. J. Physiol. 265, H517-H522[Medline] [Order article via Infotrieve]
  45. Hou, C., Kirchner, T., Singer, M., Matheis, M., Argentieri, D., and Cavender, D. (2004) J. Pharmacol. Exp. Ther. 309, 697-704[Abstract/Free Full Text]
  46. Gafni, J., Munsch, J., Lam, T., Catlin, M., Costa, L., Molinski, T., and Pessah, I. (1997) Neuron 19, 723-733[CrossRef][Medline] [Order article via Infotrieve]
  47. Peppiatt, C. M., Collins, T. J., Mackenzie, L., Conway, S. J., Holmes, A. B., Bootman, M. D., Berridge, M. J., Seo, J. T., and Roderick, H. L. (2003) Cell Calcium 34, 97-108[Medline] [Order article via Infotrieve]
  48. Smet, P. D., Parys, J. B., Callewaert, G., Weidema, A. F., Hill, E., Smedt, H. D., Erneux, C., Sorrentino, V., and Missiaen, L. (1999) Cell Calcium 26, 9-13[CrossRef][Medline] [Order article via Infotrieve]
  49. Lederer, W. J., Niggli, E., and Hadley, R. W. (1990) Science 248, 283[Free Full Text]
  50. Scriven, D., Klimek, A., Lee, K., and Moore, E. (2002) Ann. N. Y. Acad. Sci. 976, 488-499[Medline] [Order article via Infotrieve]
  51. Berridge, M. J., Bootman, M. D., and Roderick, H. L. (2003) Nat. Rev. Mol. Cell Biol. 4, 517-529[CrossRef][Medline] [Order article via Infotrieve]
  52. Pan, Z., Yang, D., Nagaraj, R., Nosek, T., Nishi, M., Takeshima, H., Cheng, H., and Ma, J. (2002) Nat. Cell Biol. 4, 379-383[CrossRef][Medline] [Order article via Infotrieve]
  53. Prakriya, M., and Lewis, R. S. (2001) J. Physiol. (Lond.) 536, 3-19[Abstract/Free Full Text]
  54. Launikonis, B. S., Barnes, M., and Stephenson, D. G. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 2941-2944[Abstract/Free Full Text]
  55. Clapham, D. E. (2003) Nature 426
  56. Vandebrouck, C., Martin, D., Colson-Van Schoor, M., Debaix, H., and Gailly, P. (2002) J. Cell Biol. 158, 1089-1096[Abstract/Free Full Text]

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