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Cardiovascular Research 2007 76(3):430-441; doi:10.1016/j.cardiores.2007.08.009
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Copyright © 2007, European Society of Cardiology

Voltage-dependent Na+ channel phenotype changes in myoblasts. Consequences for cardiac repair*

Ramón Martínez-Màrmola,b, Miren Davide, Rosario Sanchesb, Meritxell Roura-Ferrera,b, Nùria Villalongaa,b, Eleonora Sorianellob,c, Susan M. Webbd, Antonio Zorzanob,c, Anna Gumàb, Carmen Valenzuelae and Antonio Felipea,b,*

aMolecular Physiology Laboratory, Universitat de Barcelona, Spain
bDepartament de Bioquímica i Biología Molecular, Universitat de Barcelona, Spain
cParc Cientific de Barcelona, Universitat de Barcelona, Spain
dDepartament de Endocrinologia i Medicina, Hospital Sant Pau, Universitat Autonoma de Barcelona, Spain
eInstituto de Farmacología y Toxicología, CSIC/UCM, Universidad Complutense de Madrid, Spain

*Correspondence author. Molecular Physiology Laboratory, Departament de Bioquímica i Biología Molecular, Universitat de Barcelona, Avda Diagonal 645, E-08028 Barcelona, Spain. Tel.: +34 934034616; fax: +34 934021559. afelipe{at}ub.edu

Received 22 March 2007; revised 8 August 2007; accepted 17 August 2007


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Appendix A
 Appendix A
 References
 
Objective Cellular cardiomyoplasty using skeletal myoblasts is a promising therapy for myocardial infarct repair. Once transplanted, myoblasts grow, differentiate and adapt their electrophysiological properties towards more cardiac-like phenotypes. Voltage-dependent Na+ channels (Nav) are the main proteins involved in the propagation of the cardiac action potential, and their phenotype affects cardiac performance. Therefore, we examined the expression of Nav during proliferation and differentiation in skeletal myocytes.

Methods and results We used the rat neonatal skeletal myocyte cell line L6E9. Proliferation of L6E9 cells induced Nav1.4 and Nav1.5, although neither protein has an apparent role in cell growth. During myogenesis, Nav1.5 was largely induced. Electrophysiological and pharmacological properties, as well as mRNA expression, indicate that cardiac-type Nav1.5 accounts for almost 90% of the Na+ current in myotubes. Unlike in proliferation, this protein plays a pivotal role in myogenesis. The adoption of a cardiac-like phenotype is further supported by the increase in Nav1.5 colocalization in caveolae. Finally, we demonstrate that the treatment of myoblasts with neuregulin further increased Nav1.5 in skeletal myocytes.

Conclusion Our results indicate that skeletal myotubes adopt a cardiac-like phenotype in cell culture conditions and that the expression of Nav1.5 acts as an underlying molecular mechanism.

KEYWORDS Sodium channels; Cardiomyoplasty; Myogenesis; Skeletal myoblast; Cardiac repair


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 Appendix A
 Appendix A
 References
 
Cellular therapy is a promising strategy for heart failure [1–3]. Following myocardial infarction, the muscle becomes a non-contractile fibrous scar. Myocardium cannot fully regenerate because cardiac muscle cells do not re-enter the cell cycle. Unlike cardiomyocytes, skeletal myocytes proliferate and differentiate, thereby generating electrically excitable fibres [3]. Skeletal myoblasts, cardiomyocytes, and bone marrow mononuclear cells have been transplanted to replace lost myocardial tissue [1–4]. Autologous transplantation into the infarcted myocardium improves myocardial performance both in vitro and in vivo [1–3]. However, the mechanisms underlying improved cardiac function remain uncertain.

Muscle excitability is generated by the incorporation of ion channels into the surface membrane [5]. Voltage-dependent sodium channels (Nav) are the main proteins involved in propagating action potential in nerve and muscle [6,7]. Nav gene family comprises different members (Navl.l–Navl.9, NavX), and based on their sensitivity to tetrodotoxin (TTX), they are separated into TTX-sensitive and TTX-resistant [6,8]. Cardiac and skeletal muscles express different genes. While adult skeletal muscle expresses predominantly a TTX-sensitive channel (Navl.4), adult cardiomyocytes mostly present a TTX-resistant protein (Navl.5) [9,10]. However, development and muscle denervation regulate their expression [5,11–13]. For instance, TTX-sensitive currents increase and TTX-resistant channels decrease with age in skeletal muscle, but denervation counteract this phenotype [5,12,13]. Nav1.5 accounts for ~90% of the Na+ currents in the adult mammalian heart, thus determining major electrophysiological and pharmacological properties. The relative participation of other isoforms is uncertain [5,11].

The success of cellular cardiomyoplasty based on skeletal myocytes would be strongly dependent upon the acquisition of the Nav cardiac-like phenotype. The ability of skeletal myoblast to myoregenerate, as well as their resistance to ischemia, allows these cells to grow quickly within the infarcted area and generate myofibers [1–4]. Pre-treating myoblasts before transplantation has been proposed to increase cardiac-like properties [14,15]. By using neonatal-derived skeletal myoblasts, we demonstrate that myotubes improve the Nav cardiac-like phenotype in vitro. Since myoblast proliferation and differentiation are important in graft generation, we characterized Nav expression in skeletal myoblasts in culture. Proliferation increased Navl.4 and Nav1.5 expression. However, neither protein has an apparent role in the cell cycle progression. Myotube differentiation induces Navl.5, which triggers a cardiac-like phenotype. Navl.5 plays a pivotal role in myogenesis. Furthermore, neuregulin-1, a growth factor that regulates differentiation and which has been implicated in the protection and survival of cardiac and skeletal muscle, further increased the adoption of a cardiac-like Nav phenotype. Our results suggest that myoblasts originating from an early state of differentiation and treated in culture prior to transplantation, adopt more optimal electrical cardiac phenotypes.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 Appendix A
 Appendix A
 References
 
2.1 Cell culture
The rat skeletal muscle-derived L6E9 cell line was kindly provided by Dr. B. Nadal-Ginard (Cardiovascular Institute and Center for Cardiovascular Health, Mount Sinai School of Medicine, NY). Myoblasts were grown in Dulbecco's modified medium (DMEM) supplemented with 10% fetal bovine serum (FBS), antibiotics, 2 mM L-glutamine, and 25 mM Hepes (pH7.4). Pre-confluent myoblasts (70–80%) were induced to differentiate by lowering FBS to a final concentration of 2%. In some experiments, aqueous solutions of TTX (Alomone) were simultaneously added to the media. Recombinant neuregulin 1 containing the bioactive EGF domain, heregulin-β1 (NRG), was from PromoKine.

Primary culture of adult human skeletal muscle myoblasts were obtained from a deltoid muscle biopsy. The sample was minced and cultured as previously described [16]. Myoblast were cultured in 75% DMEM and 25% M199 medium, supplemented with 10% FBS, 2 mM glutamine, antibiotics, 0.25 µg/ml fungizone, 10 µg/ml insulin, 10 ng/ml epidermal growth factor, and 25 ng/ml fibroblast growth factor. To obtain highly purified myoblasts, primary cultures were sorted for the early surface marker CD56 by immunomagnetic selection [16]. CD56-positive cells were seeded in the medium containing 15% FBS until ~80% confluence. Myogenesis was induced by lowering FBS to 2%.

2.2 Proliferation assay
DNA synthesis was determined by the incorporation of 3H-thymidine (GE Healthcare) into DNA. L6E9 cells (2x104) were seeded in 24-well plates without FBS, supplemented with 0.2% BSA, for 36 h. Cells were further cultured for 24 h in the absence (resting) or presence (proliferation) of 10% FBS with/without aqueous solutions of TTX. Finally, this was replaced by the same medium containing 1 µCi/ml [3H]-thymidine. After three hours, cells were fixed in 70% methanol, washed three times in ice-cold 10% trichloracetic acid, and solubilized in 1% SDS and 0.3% NaOH. The content of the well was used for counting radioactivity.

2.3 RNA isolation, RT-PCR analysis and real-time PCR
Total RNA from rat tissues (brain, heart, and skeletal muscle) and L6E9 cells was isolated using the Tripure reagent (Roche Diagnostics). All aspects of this investigation conformed to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). RNA was treated with DNaseI and PCR controls were performed in the absence of reverse transcriptase. cDNA synthesis was performed with transcriptor reverse transcriptase (Roche) with a random hexanucleotide and oligo dT, according to the manufacturer's instructions. Once cDNA was synthesized, the conditions were set for further PCR: 94 °C for 1 min, 1 min at the corresponding annealing temperature (see Table 1 in Supplementary Material (SM)), and 72 °C for 1 min. These settings were applied for 30 cycles.

Real-time PCR was performed using a LightCycler machine with LightCycler FastStart DNA MasterPLUS SYBR Green I (Roche). The reactions for Nav1.4 and Nav1.5 (specific Nav1.5 settings in parenthesis) were performed as follows: 95 °C for 5 s, 61 °C (58 °C) for 6 s, 72 °C for 9 s (11 s) and 85 °C for 2 s, preceded by 10 min at 95 °C and followed by 10 s at 95 °C and 30 s at 65 °C. 18 S was included as an internal reference. The chosen forward and reverse PCR primers, PCR annealing temperatures, PCR product length and accession numbers are shown in SM Table 1. The real-time PCR efficiency (E) of one cycle in the exponential phase was calculated according to the equation: E=10(–1/slope). The normalized Nav1.5/Nav1.4 ratio was calculated as follows: Ratio=(1+E) {Delta}Ct (Nav1.5)/(1+E) {Delta}Ct (Nav1.4), where Ct signifies the threshold cycle.

2.4 Membrane protein extracts and Western blot
Cells were washed in cold phosphate-buffered saline (PBS) and lysed on ice with lysis solution (1% NP40, 10% glycerol, 50 mmol/L HEPES pH 7.5, 150 mmol/L NaCl), supplemented with 1 µg/ml aprotinin, 1 µg/ml leupeptin, 86 µg/ml iodoacetamide and 1 mM phenylmethylsulfonyl fluoride as protease inhibitors. To obtain enriched membrane preparations, homogenates were centrifuged at 3000 g for 10 min and the supernatant was further centrifuged at ~150,000 g for 90 min. The pellet was resuspended in 30 mM HEPES (pH 7.4). To concentrate enriched membrane proteins from L6E9 cells, Microcon centrifugal filter devices (Amicon) were used.

Crude membrane protein samples were separated on 7.5% SDS-PAGE, transferred to nitrocellulose membranes (Immobilon-P, Millipore) and blocked in 5% dry milk-supplemented 0.2% Tween 20 PBS. Filters were immunoblotted with antibodies against Nav1.5 (1/200, Alomone), myosin heavy chain (MHC; 1/20, MF20-anti-meromyosin — from Developmental Studies Hybridoma Bank), myogenin (1/20, F5D, DSHB), Na/K ATPase {alpha}1-subunit (1/100, {alpha}6F, DSHB), caveolin 3 (1/5000, BD Biosciences) and pan-caveolin (1/200, BD Biosciences). As a loading and transfer control, we used a monoclonal anti-β-actin antibody (1/5000, Sigma).

2.5 Immunolocalization, nuclei staining and fusion index
A Monoclonal Anti-pan-Sodium Channel (1/100, clone K58/35, Sigma) was used. Anti-caveolin polyclonal (1/250), which recognizes caveolin isoforms 1, 2, and 3, was from BD Biosciences. CY2 and CY3 secondary antibodies (1/250) were from Amershan.

L6E9 cells fixed with 4% paraformaldehyde in PBS for 10 min were further permeabilized with 1% Triton X-100/PBS for 10 min. After 60 min incubation with blocking solution (10% goat serum/5% non fat dry milk/0.75% Triton X-100/PBS), cells were reacted with antibodies in 10% goat serum/0.75% Triton X-100/PBS for 90 min. Cells were further incubated with CY3 anti-rabbit antibody in PBS. CY2 anti-mouse was used for caveolin visualization. For nuclei staining, fixed and permeabilized cells were stained with DAPI (50 ng/ml) in PBS for 2 min. Under a fluorescent microscope, several randomly chosen fields were photographed, and the nuclei per cell were counted. The fusion index was calculated as follows: Fusion Index (%)=(number of nuclei in myotubes/total number of nuclei in the field)*100.

2.6 Electrophysiology
L6E9 cells were held at –120 mV and Na+ currents generated by 10 ms square pulses from –90 mV to +60 mV in 10 mV steps. Normalized conductance was set by using Boltzmann's equation: G/Gmax=1/(1+exp((V1/2–V)/k)). Vh is the voltage at which the current is half activated and k is the slope factor of the activation curve. To calculate voltage-dependent inactivation, cells were held at –120 mV and pre-pulses (2.75 s) to various potentials, ranging from –140 mV to +10 mV in 10 mV steps, followed by a test pulse of –10 mV during 80 ms were applied. Steady-state inactivation plots were fitted to a Boltzmann equation: I/Imax=1/(1-exp ((V1/2–V)/k)). Currents were recorded at room temperature (21–23 °C) using the whole cell patch-clamp technique with an Axopatch 1C amplifier (Axon Instruments). Micropipettes were pulled from borosilicate glass capillary tubes (GD-1; Narishige) using a P-87 puller (Sutter Instrument). The intracellular pipette solution contained (in mM) 5 NaCl, 5 KCl, 130 CsF, 1 MgCl2, 4 MgATP, 10 HEPES-K, and 5 EGTA and adjusted to pH 7.2 with CsOH. The bath solution contained (in mM) 140 NaCl, 5 KCl, 1.8 CaCl2, 1 MgCl2, 10 HEPES-Na, 10 glucose, 20 TEA, and 2 CoCl2 and adjusted to pH 7.4 with NaOH. TTX (Alomone) was dissolved in water. Micropipette resistance was set at 1–2 M{Omega}. Data analysis was performed using pClamp 9.0.1 and Origin 7.0.3 (Microcal Software).


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 Appendix A
 Appendix A
 References
 
3.1 Voltage-dependent Nav1.4 and Nav1.5 generate inward Na+ currents in L6E9 myoblasts
Na+ currents were evoked in L6E9 myoblasts (Fig. 1). Cells were held at –120 mV and 10 ms square pulses were applied from –90 to +60 mV in 10 mV steps (Fig. 1A). The threshold potential was ~–60 mV, and the I/V relationship exhibited a peak at –20 mV (Fig. 1B). Normalized conductance versus test potential indicated that the half-activation voltage was –36.8±2.2 mV with a k slope value of 7.4±0.4 mV (Fig. 1C). Steady-state availability curves were obtained by plotting current data versus test potential (Fig. 1C). The Vh was –80.9±2 mV with a slope factor of 6.6±0.2 mV. L6E9 cells expressed TTX-sensible and TTX-resistant channels (Fig. 1D). IC50 values were 0.8 nM and 1 µM for the high and the low-affinity, respectively. Na+ currents evoked (–20 mV) in the presence of 100 nM TTX indicated that the TTX-sensitive current was about 40% of the total.


Figure 1
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  		Fig. 1 L6E9 myoblasts express voltage-dependent Na+ channels. (A) Representative traces of Na+ currents. Cells were held at –120 mV and pulse potentials were applied. (B) Current intensity as a function of membrane potential. L6E9 cells were held at –120 mV and currents were evoked by 10 ms square pulses from –90 mV to +60 mV in 10 mV steps. (C) Normalized conductance (•) and Steady-state availability ({circ}) curves plotted against test potentials. To calculate steady-state inactivation cells were held at –120 mV and pre-pulses (2.75 s) to various potentials, ranging from –140 mV to +10 mV in 10 mV steps, followed by a test pulse of –10 mV during 80 ms were applied. (D) Dose-dependent inhibition curves of the Na+ current by TTX. Currents were evoked at –20 mV from a holding potential of –120 mV during a pulse potential of 10 ms. The percentage of inhibition was calculated by comparing the current at a given concentration of toxin versus that obtained in its absence. Values represent the mean±SEM of at least 6 independent experiments. (E) mRNA expression of Nav1.1, Nav1.3, and Nav1.6 in the rat brain, but not in L6E9 cells. (F) Nav1.4 and Nav1.5 mRNA expression. RT-PCR reactions were set as described in the Materials and methods section with oligonucleotides described in SD Table 1. Rat skeletal muscle and heart were used as positive controls. PCR reactions were performed in the presence (+RT) and absence of the RT reaction (–RT). PCR products were run in a 2% agarose gel. (G) Nav1.4 and Nav1.5 mRNA expression levels quantified by real-time PCR. For each primer set, a standard curve was made and the slope factor calculated. The corresponding real-time PCR efficiency (E) of one cycle in the exponential phase was calculated according to the equation: E=10(–1/slope). The normalized Nav1.4 and Nav1.5 expression were calculated as follows: ((1+E) {Delta}Ct(GOI))/((1+E) {Delta}Ct(control)), where Ct represents the threshold cycle; GOI, gene of interest; control, 18S. ***p<0.001 vs. Nav1.4 (Student's t test). (H) Representative confocal images of Nav1 immunofluorescence in L6E9 myoblasts. Left panel, DAPI staining of nuclei; Center panel, Nav1 expression; Right panel, overlay of both images.

 
Several Nav may be responsible for Na+ currents in L6E9 [8]. Nav1.1, Nav1.3, Nav1.4, Nav1.5 and Nav1.6 mRNA expression were analyzed, with only Nav1.4 and Nav1.5 being detected (Fig. 1E and F). Brain, heart and skeletal muscle were used as controls. Nav1.5 expression was 3-fold that of Nav1.4 (Fig. 1G). Five hundred µg of membrane protein was necessary to detect Nav1.5 by Western blot (not shown). As such sample amounts render analysis difficult, we conducted immunodetection assays. Although several antibodies were used, the monoclonal anti-pan Nav (Sigma, clone K58/35) yielded the best results (Fig. 1H).

3.2 Myoblast proliferation induces Nav1.4 and Nav1.5
Proliferation led to a 3-fold increase in Na+ currents in L6E9 cells (Fig. 2A). Current intensity versus test potential plot indicated that currents shared voltage dependence (Fig. 2 B). Vh and k values of activation (Fig. 2C) and inactivation (Fig. 2D) were similar both in resting and proliferating cells. Half-activation voltages and slope values were –32.5±1.8 mV and 9.2±0.7 mV, and –36.8±2.2 mV and 8.3±0.3 mV for resting and proliferating myoblasts, respectively (SM Table 2). Similarly, Vh and the slope factor of the availability curves were –84.8±2.2 mV and 5.9±0.3 mV, and –81.2±2.1 mV and 6.4±0.3 mV in resting and proliferating cells, respectively (SM Table 2). Proliferation triggered a 3- to 4-fold transient increase in Nav1.4 and Nav1.5 mRNA expression (Fig. 2E) and the Nav1.5/Nav1.4 ratio remained constant (Fig. 2F). Since K+ channels play a role during proliferation [17], we assessed whether Nav channels would also be involved. Fig. 2G shows that TTX did not inhibit the proliferation-dependent 3H-thymidine incorporation into myoblasts indicating that Nav have no apparent role.


Figure 2
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  		Fig. 2 Myoblast proliferation induced Nav channels. (A) Representative traces of Na+ currents. Resting cells were incubated for 24 h in the absence (resting) or presence of FBS (proliferation). Currents were elicited by a depolarizing step as indicated. (B) Current intensity versus voltage relationship of Na+ currents. Cells were held at –120 mV and pulse potentials as described in Fig. 1 were applied. (C) Normalized conductance versus test potential plot. (D) Steady-state inactivation curves. A double-pulse protocol was applied as described in Fig. 1. Symbols for B–D are: {circ}, resting; •, proliferation. (E) Proliferation induces Nav1.4 and Nav1.5 mRNA in L6E9 myoblasts. {circ}, Nav1.4; •, Nav1.5. Samples were collected after the addition of FBS and real-time PCR analysis was performed at the indicated times. (F) Nav1.4/Nav1.5 mRNA ratio. The normalized Nav1.4 and Nav1.5 expression levels were calculated following the equation in Fig. 1. (G) TTX did not inhibit proliferation in L6E9 cells. Proliferating myoblasts were cultured in the presence of TTX (0.1 and 100 µM). Values represent the mean±SEM of 3 different assays, each done in triplicate.

 
3.3 Myogenesis differentially regulates voltage-dependent Na+ channels
SM Fig. 1 shows that myoblasts formed myotubes, with a fusion index over 60%. Concomitantly, while myogenin was early induced, MHC increased steadily. Na/K ATPase {alpha}1 subunit and β-actin controls remained constant [18].

Pulse potentials evoked Na+ currents in 96 h-myotubes (Fig. 3A, SM Table 2). Currents were 4-fold higher in myotubes than in myoblasts and the peak current density steadily increased during myogenesis (Fig. 3B and C). The activation threshold shifted to –70 mV in myotubes (Fig. 3D). Vh shifted –11 mV (–48.2±3.4 vs. –36.8±2.2 mV) in myotubes (Fig. 3E) with no changes in the k slope (6.6±0.9 vs. 7.4±0.4 mV for myotubes and myoblasts, respectively). In addition, the half-inactivation voltage (Fig. 3F) was also more electronegative (–92.6±2.4 vs. –80.9±2.0 mV for myotubes and myoblasts, respectively). In order to differentiate between Nav1.4 (TTX-sensible) and Nav1.5 (TTX-resistant), currents were elicited in the presence of 100 nM TTX (Fig. 3G). While the contribution of TTX-sensitive component in myoblasts comprised about 40% of the total, this value decreased to 16% in myotubes (Fig. 3H).


Figure 3
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  		Fig. 3 Myogenesis induced Nav currents. (A) Na+ currents in myotubes. Currents were elicited in 96-h differentiated myotubes by depolarizing steps as indicated. (B) Representative traces of Na+ currents in myoblasts and myotubes. (C) Current density plotted as a function of differentiation time. *p<0.05 vs. 0 h, Student's t test. (D) Current intensity versus voltage relationship of Na+ currents. Pulse potentials as described in panel A were applied. (E) Normalized conductance of Na+ currents in myotubes versus test potential. (F) Steady-state inactivation curves. Dashed lines in B, E and F represent myoblasts. Symbols for D–F: {circ}, myoblasts; •, myotubes. (G) Peak current plotted as a function of membrane potential in the presence of 100 nM TTX. The TTX-sensitive component was calculated by comparing the current at a given potential in the presence of the toxin versus that obtained in its absence. Symbols are: •, overall intensity; {circ}, TTX-resistant; {Delta}, TTX-sensitive. (H) Major components of Na+ currents in myoblasts (0 h) and myotubes (96 h). The peak current intensity (–20 mV) was considered 100%. TTX-sensitive and TTX-resistant components are calculated as shown in panel G. Open bars, TTX-resistant; closed bars, TTX-sensitive. Values represent the mean±SEM of at least 6 independent cells.

 
Nav1.4 and Nav1.5 mRNA expression was analysed during myogenesis (Fig. 4A). Unlike Nav1.4, Nav1.5 increased steadily during differentiation and myotubes expressed much more Nav1.5 (Fig. 4B). Myotubes expressed more Nav protein (Fig. 4 F and K). Although the antibody recognized both Nav1.4 and Nav1.5, biophysical and pharmacological properties, and mRNA expression indicated that the increase in the Nav signal stemmed from selective Nav1.5 induction.


Figure 4
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  		Fig. 4 Myogenesis specifically induces Nav1.5 and co-localizes the channel with caveolin. (A) Real-time PCR was performed as described in Fig. 1. Differentiation was induced and samples were collected at the indicated times. Symbols are: {circ}, Nav1.4; •, Nav1.5. (B) Nav1.4/Nav1.5 mRNA ratio. The normalized Nav1.4 and Nav1.5 expression levels were calculated following the equation in Fig. 1. Values represent the mean±SEM of 4 different assays, each done in triplicate. (C) Expression of myogenic markers. MHC, myosin heavy chain; Na/K, {alpha}1 subunit of Na/K ATPase; Cav3, caveolin 3; Cav, overall caveolin (1–3); β-act, β-actin. (D-M) Immunofluorescence of Nav1 and caveolin in L6E9 cells. (D–H) myoblasts (0 h). (I–M) myotubes (96 h). Nav1, Nav1.X; cav, overall caveolins; merge, overlay of signals.

 
3.4 Myogenesis-induced Nav1.5 efficiently targets to caveolae
Nav target to caveolae in heart playing a mayor role in the cardiovascular system [19,20]. Myogenesis induced MHC and caveolin 3, as previously reported (Fig. 4C) [18,21]. However, caveolin 1 down-regulates during myogenesis [22]; thus, we used an anti-pan caveolin antibody. Fig. 4C shows that the overall caveolin abundance (cav1-3) remained constant. While Nav weakly co-localized with caveolin in myoblasts (Fig. 4F–H), myogenesis-induced Nav1.5 strongly co-localized with caveolin in myotubes (Fig. 4K–M).

3.5 Nav1.5 plays a pivotal role during myogenesis
L6E9 cells were induced to differentiate in the presence of TTX. TTX (100 nM), which inhibits Nav1.4, did not modify the expression of MHC and caveolin 3 (Fig. 5A and B). Surprisingly, in 2 out of 3 experiments, myogenin was induced. In contrast, 100 µM TTX, which blocks Nav1.5, decreased MHC, caveolin 3, and myogenin. Again, the overall caveolin abundance did not change. In addition, 100 µM TTX generated fewer and smaller myotubes (Fig. 5C) and the fusion index was affected (Fig. 5D). While myotubes accounted for ~70% of the culture, 100 µM TTX reduced this value to <50%. Our results demonstrated that TTX, at doses which inhibited Nav1.5, impaired the myogenesis of L6E9 myoblasts.


Figure 5
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  		Fig. 5 Nav1.5 is involved in myogenesis. (A) Protein expression of myogenic markers was impaired by the presence of TTX. MHC, myosin heavy chain; Cav3, caveolin 3; Cav, overall caveolin (cav1-3); Myog, myogenin; β-act, β-actin. (B) Percentage of expression. At least 4 Western blots were performed and the relative abundances of MHC, Cav3 and Myogenin were analyzed. (C) Phase images of differentiated L6E9 in the absence or presence of 100 µM TTX. (D) Fusion index in the absence or presence of 100 µM TTX. Open bars, no TTX; grey bars, 0.1 µM TTX; closed bars, 100 µM TTX. Values represent the mean±SEM of at least 4 independent experiments. *p<0.05; **p<0.01; ***p<0.001 vs. control (0 TTX), Student's t test.

 
3.6 Neuregulin 1 further induced L6E9 myogenesis, concomitantly with an increase in Nav1.5
Since myoblasts may be preconditioned prior to transplantation, we sought to determine whether NRGs, the growth factors involved in myocyte survival and differentiation [23,24], improved the cardiac-like phenotype. Cells were induced to differentiate and NRG (3 nM) was added 24 h later and studies were carried out after 2 days of NRG treatment [23]. NRG-treated cells achieved higher levels of differentiation, although 100 µM TTX inhibited myogenesis (Fig. 6A and B). In addition, NRG further increased Nav1.5 expression but had no effect on Nav1.4 (Fig. 6C). Moreover, we further confirmed the role of Nav1.5 (Fig. 6D). TTX diminished NRG-induced MHC and caveolin 3 expression. NRG did not alter the myogenin expression, indicating that its induction occurred before the addition of NRG (Fig. 6D). These results were further confirmed by immunofluorescence of caveolin 3 (SM Fig. 2). Caveolin 3 immunoreactivity increased in the presence of NRG while TTX diminished the signal.


Figure 6
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  		Fig. 6 Neuregulin-dependent myogenesis further induced Nav1.5 in L6E9 cells. Myoblasts were induced to differentiate by serum depletion in the presence or absence of 100 µM TTX. NRG was added (+NRG) 24 h after the cells were changed to the low serum medium, and studies were carried out after 2 days of NRG treatment (72 h of differentiation). (A) Phase images of L6E9 cells. (B) Fusion index of L6E9 cells in the presence (closed bars) or absence (open bars) of NRG, (C) Nav1.4 (open bars) and Nav1.5 (closed bars) mRNA expression. Values represent the mean±SEM of at least 3 independent experiments. *p<0.05; **p<0.01; ***p<0.001 (Student's t test). (D) Expression of myogenic markers. Protein extracts were analyzed by Western blot. MHC, myosin heavy chain; Cav 3, caveolin 3; Myog, myogenin; Na/K, {alpha}1 subunit of Na/K ATPase; β-act, β-actin.

 
3.7 Differential expression of Nav1.4 and Nav1.5 during proliferation and differentiation in human skeletal muscle cells
To further confirm the data of L6E9 cells, we performed a serial of experiments in primary human skeletal muscle cultures (SM Fig. 3). Human myoblasts were induced to proliferate during 24 h and to differentiate for 5 days. MHC expression increased in myotubes (SM Fig. 3A). While proliferation induced Nav1.4 and Nav1.5, differentiation specifically increased Nav1.5 (SM Fig. 3B–D). Unlike neonatal L6E9 cells, Nav1.4 mRNA was much more abundant than Nav1.5 in adult human myoblasts (SM Fig. 3C and D). While the Nav1.4/ Nav1.5 ratio ({Delta}Ct (Nav1.5–Nav1.4)) remained constant during proliferation, myogenesis triggered an important decrease (SM Fig. 3C and D). However, Nav1.4 was still predominant in adult skeletal myotubes as previously described [12]. Furthermore, immunocytochemistry further confirmed the Nav1.5 accumulation in myotubes (SM Fig. 3E).


   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
Appendix A
 Appendix A
 References
 
The use of skeletal myoblasts for cardiac repair in patients with myocardial infarction has been an active field of research in recent years [1–4]. Skeletal myoblasts form contractile myofibers and the acquisition of a cardiac phenotype is crucial to successful engraftment. The expression of cardiac proteins, such as ion channels, is important for the establishment of electromechanical junctions. Therefore, an understanding of the molecular events underlying engraftment is essential to improving this cellular therapy.

Since Nav are the main proteins involved in the propagation of cardiac action potential [6,7], we analyzed the physiological regulation of Nav during proliferation and myogenesis in skeletal myoblasts. The neonatal skeletal muscle cell line L6E9 expresses Nav1.4 and Nav1.5. Unlike C2C12 cell line, the cardiac-like Nav1.5 expression was strong in L6E9 myoblasts [14]. L6E9 cells do not contract in vitro, suggesting an early state of differentiation [25]. This is relevant to our results. Mature skeletal muscle expresses Na+ channels that are sensitive to TTX [12], which is in agreement with our human skeletal muscle cell cultures. However, embryonic muscles synthesize TTX-insensitive isoforms [8,12]. In this context, myoblasts preconditioned in culture express more TTX-resistant cardiac-like Nav. Differentiated C2C12 myotubes express a low Nav-cardiac phenotype but Nav1.5 slightly increases (~30%) and electrophysiological parameters shift to more hyperpolarised values in conditioned cardiac culture media [14]. However, L6E9 myotubes augmented more than 8 fold Nav1.5 and this was further increased by the presence of NRG. Our data indicate that by using weakly differentiated skeletal myoblasts (i.e., foetal or neonatal), cells achieved higher levels of the cardiac phenotype.

Unlike cardiomyocytes, skeletal myocytes proliferate. We describe, for the first time, Nav expression during skeletal myocyte proliferation. We found a mRNA increase concomitantly with a raise in Na+ conductance. This could be a consequence of changes in the membrane potential during the cell cycle progression, in which K+ channels are involved [17]. However, unlike K+ channels, Na+ channels do not play any apparent role.

TTX-resistant Na+ channels account for over 90% of Na+ currents in adult cardiac myocytes. The remaining 10% are generated by neural Nav1.X, skeletal muscle Nav1.4, and probably other NavX [5,11]. This is similar to what we found in myotubes. Na+ currents in L6E9 myoblasts have Vh around –35 mV, which is higher than the value of Nav1.4 (–25 mV), but lower than that of Nav1.5 (–40 mV). Previous works demonstrate that C2C12 cells only achieved a Nav-like current phenotype (–43 mV) by culturing myotubes in conditioned cardiac culture media [14]. However, L6E9 myotubes expressed Na+ channels that were half maximally open at a membrane potential around –48 mV, quite similar to Nav1.5 [9,10]. Comparable analyses reflected similar inactivation rates, whereby cardiac Nav1.5 and L6E9 myotubes share more hyperpolarised values [9].

Localization of ion channels is also necessary for proper electrical signalling [19,20]. While cardiac-Nav target to caveolin-rich domains [19], their localization in skeletal muscle remained unknown. Myogenic-induced Nav1.5 co-localized with caveolin, which places the channels in close proximity to signalling molecules. Thus, the β-adrenergic receptor stimulation of cardiac sodium channels is associated with caveolae in the sarcolemma, a localization which improves cardiac performance [19].

Unlike proliferation, Nav1.5 is involved in myogenesis, in which K+ channels play a pivotal role [18,26–28]. Nav are responsible for the initiation and propagation of action potentials [6,7]. Therefore, changes in membrane potential may be preceded by Nav functions. Low concentrations of TTX induce myogenin and Nav currents [29,30]. In this scenario, myogenin promotes myogenesis [29,31]. Our data in no way challenge this result. TTX (100 nM) did not inhibit myogenesis, but rather increased myogenin ~50%. However, higher doses of TTX decreased differentiation.

Cardiomyoplasty with cells expressing distinct electrophysiological properties may cause side effects [1–4]. Thus, preconditioning myoblasts appears to improve cardiac phenotype acquisition [14,15]. However, the percentage of cells that survive and differentiate following transplantation remains very low [3]. We found that NRG, a survival factor involved in cardiomyocyte and skeletal myocyte differentiation [23,24], further induced the Nav cardiac-phenotype in vitro. Alternative approaches, such as intramyocardial injection in conjunction with coronary artery bypass or combined cardiomyoplasty with bone marrow cells, increases the graft survival [4,32]. Coronary artery bypass and revascularization with bone marrow cells improve ischemic area perfusion and cardiac function [3,32,33]. In this context, our NRG results have physiological relevance. Similar to what occurs in coronary bypasses, NRG increases glucose uptake [23]. In addition, NRG stimulates the survival of neurons surrounding cortical brain injuries in vivo [34].

Cardiomyoplasty is not only feasible, but has been associated with improved cardiac function. While the Myoblast Autologous Grafting in Ischemic Cardiomyography (MAGIC) trial failed to reach its main objectives, the experience partially reversed remodeling indicating that additional experimental studies are still warranted in order to address important issues central to improving this cellular therapy. Therefore, knowledge of the molecular events surrounding myoblast engraftment is crucial.


   Appendix A
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Appendix A
Appendix A
 References
 
Supplementary Material


   Appendix A
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Appendix A
 Appendix A
References
 
Supplementary data
Supplementary data associated with this article can befound, in the onlineversion, at doi:10.1016/j.cardiores.2007.08.009.


Figure 1
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  		Supplemental Figure 1 Serum depletion induces myogenesis in L6E9 myoblasts. Sub-confluent myoblasts were induced to differentiate by serum depletion with images and samples obtained at different times. (A) Images show that L6E9 mononucleated myoblasts differentiated into multinucleated myotubes. (B) Fusion index. Cells were fixed in glutaraldehyde and nuclei were stained with Giemsa. The fusion index was calculated as the number of multinucleated cells in a fixed area. At least 10 different fields were randomly analyzed from each plate. Values represent the mean±SEM of at least 3 independent experiments. (C) Western blot analysis of myogenic markers. Myog, myogenin; MHC, myosin heavy chain; Na/K, {alpha}1 subunit of the Na+/K+ ATPase; β-act, β-actin. Four independent experiments were analyzed; representative filters are shown.

 


Figure 2
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  		Supplemental Figure 2 Neuregulin-1-induced myogenesis is inhibited by the presence of TTX. Myoblasts were induced to differentiate by serum depletion in the presence or absence of 100 µM TTX. 3 nM NRG was added (+NRG) 24 h after the cells were changed to the low serum medium, and studies were carried out after 2 days of NRG treatment (72 h of differentiation). Cells were fixed and nuclei staining (blue) and caveolin 3 detection (green) were performed as described in Materials and methods section.

 


Figure 3
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  		Supplemental Figure 3 Expression of Nav1.4 and Nav1.5 during proliferation and differentiation in human skeletal muscle cells. A deltoid muscle biopsy from a 60 year-old woman was performed and skeletal muscle cells were isolated. To study proliferation, myoblasts (40–60% of confluence) were cultured in medium without FBS, supplemented with 0.2 % BSA, for 36 h. Cells were further cultured for 24 h in the absence or presence of 10% FBS. Pre-confluent myoblasts (70–80%) were induced to differentiate during 5 days by lowering FBS to a final concentration of 2%. (A) Myosin Heavy Chain (MHC) expression in 5-days-differentiated myotubes. (B) Nav1.4 and Nav1.5 mRNA expression by Real-time PCR. Open bars, Nav1.4; Closed bars, Nav1.5. (C) Nav1.4/Nav1.5 relative abundance during proliferation. Open bars, resting cells; closed bars, proliferating cells. (D) Nav1.4/Nav1.5 relative abundance during differentiation. Open bars, myoblasts; closed bars, myotubes. Left panels represent Ct values; right panels represent differences between Nav1.4 and Nav1.5 ({Delta}Ct (Nav1.5–Nav1.4)). Ct means the threshold cycle. Note that low Ct values indicate high mRNA expression. Values are mean±SEM of four independent experiments. *p<0.05; ***p<0.001; ns, not significant (Student's t test). (E) Immunofluorescence Nav1 detection in myoblasts (0 days) and myotubes (5 days). Cells were fixed and nuclei staining (blue) and Nav1 detection (green) were performed as described in Materials and Methods. Left panels, overlay of phase and nuclei; right panels, overlay of nuclei and Nav1.

 

   Acknowledgements
 
Supported by the Ministerio de Educación y Ciencia (MEC), Spain (BFI2002-00764 and BFU2005-00695 to AF; SAF2005-00489 to AG; SAF2004-06856, CAM GR/SAL/0854/2004 and FIS RD06/0014/0006 to CV). RMM, RS and NV hold fellowships from the MEC and MRF from the Generalitat de Catalunya. We thank the editorial assistance of the Language Advisory Service (University of Barcelona). RMM and MD contributed equally. CV and AF contributed equally.


   Notes
 
* AF designed research. AZ coordinated human biopsies and contributed with reagents. SMW collected human samples. ES isolated human myoblasts. RMM, MD and CV performed electrophysiology. RMM, RS, MRF, NV undertook research. RS and AG performed NRG experiments. RMM, MD, CV and AF analysed data. AF and CV wrote the paper. Back


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Appendix A
 Appendix A
 References
 

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