AJP - Heart BIOPAC complete lab solutions
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 292: H439-H450, 2007. First published September 15, 2006; doi:10.1152/ajpheart.00119.2006 Free Article
0363-6135/07 $8.00
This Article
Free upon publication Free Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
292/1/H439    most recent
00119.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (4)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Zebedin, E.
Right arrow Articles by Hilber, K.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Zebedin, E.
Right arrow Articles by Hilber, K.

C2C12 skeletal muscle cells adopt cardiac-like sodium current properties in a cardiac cell environment

Eva Zebedin, Markus Mille, Maria Speiser, Touran Zarrabi, Walter Sandtner, Birgit Latzenhofer, Hannes Todt, and Karlheinz Hilber

Center of Biomolecular Medicine and Pharmacology, Institute of Pharmacology, Medical University of Vienna, Vienna, Austria

Submitted 1 February 2006 ; accepted in final form 7 September 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Intracardiac transplantation of undifferentiated skeletal muscle cells (myoblasts) has emerged as a promising therapy for myocardial infarct repair and is already undergoing clinical trials. The fact that cells originating from skeletal muscle have different electrophysiological properties than cardiomyocytes, however, may considerably limit the success of this therapy and, in addition, cause side effects. Indeed, a major problem observed after myoblast transplantation is the occurrence of ventricular arrhythmias. The most often transient nature of these arrhythmias may suggest that, once transplanted into cardiac tissue, skeletal muscle cells adopt more cardiac-like electrophysiological properties. To test whether a cardiac cell environment can indeed modify electrophysiological parameters of skeletal muscle cells, we treated mouse C2C12 myocytes with medium preconditioned by primary cardiocytes and compared their functional sodium current properties with those of control cells. We found this treatment to significantly alter the activation and inactivation properties of sodium currents from "skeletal muscle" to more "cardiac"-like ones. Sodium currents of cardiac-conditioned cells showed a reduced sensitivity to block by tetrodotoxin. These findings and reverse transcription PCR experiments suggest that an upregulation of the expression of the cardiac sodium channel isoform Nav1.5 versus the skeletal muscle isoform Nav1.4 is responsible for the observed changes in sodium current function. We conclude that cardiomyocytes alter sodium channel isoform expression of skeletal muscle cells via a paracrine mechanism. Thereby, skeletal muscle cells with more cardiac-like sodium current properties are generated.

sodium channel isoform expression; myoblast transplantation

INTRACARDIAC TRANSPLANTATION of undifferentiated skeletal myoblasts is a new therapy after myocardial infarction (recently reviewed in Refs. 21, 42), and early clinical trials are promising (6, 12, 22, 37). In a cardiac environment, transplanted myoblasts retain their ability to fuse with each other and form contractile differentiated skeletal muscle tissue (11, 23, 32). The actual mechanism responsible for the improvement of heart function after myoblast transplantation, however, remains to be explored.

The fact that skeletal muscle cells have different electrophysiological properties than cardiomyocytes very likely limits the efficacy of myoblast transplantation therapy. In particular, specific skeletal muscle cell electrophysiological features (functional ion channel properties and lack of connexin43 expression) do not allow for proper cardiac-like impulse conduction (e.g., Ref. 35) and thus may prevent recovery of heart function by synchronous contraction of the skeletal graft with cardiac host tissue. Induction of cardiac-like electrophysiological properties in skeletal muscle cells, if possible, would be a strategy to overcome this problem.

Transplantation of cells with "unfitting" electrophysiological properties into the heart may also cause side effects. In fact, serious ventricular arrhythmias occur (recently reviewed in Ref. 27), and cases of sudden cardiac death have been reported after myoblast transplantation. Menasche (19) suggested that the introduction of skeletal muscle cells with unfitting electrophysiological properties into cardiac host tissue could result in heterogeneities in action potential (AP) conduction, thereby setting the stage for arrhythmias. The interesting observation that the arrhythmias following myoblast transplantation most often occur only transiently (in the initial weeks) (e.g., Refs. 7, 19, 20, 37, 41) may imply that, once transplanted into cardiac tissue, skeletal muscle cells adapt their electrophysiological properties toward more cardiac-like ones, and this may finally reduce their arrhythmogenicity.

To test whether electrophysiological parameters of skeletal muscle cells do indeed shift toward more cardiac-like ones in a "cardiac milieu," for this study we designed a simple in vitro system. We treated mouse C2C12 skeletal muscle cells with differentiation medium preconditioned by primary cardiocytes and compared their functional sodium (Na+) current properties with those of control cells. We chose to study Na+ currents for two reasons. First, Na+ currents exhibit strong and well-defined functional differences between skeletal and cardiac muscle cells (e.g., Refs. 24, 44, 45). Thus emerging adaptations of their functional properties toward more cardiac-like ones can easily be judged. Second, skeletal muscle-like Na+ current properties are not suited for the electrophysiological requirements of the heart and may disturb proper AP conduction, thereby generating arrhythmias. Namely, in the heart, long-lasting (several hundred milliseconds) and repetitive (>1 Hz) depolarization is a normal characteristic of cardiomyocyte function. Under such conditions, skeletal muscle (Nav1.4) Na+ channels [in contrast to cardiac (Nav1.5) Na+ channels] would undergo almost complete slow inactivation within a few minutes (31, 35). Consequently, the strong tendency of skeletal muscle Na+ channels to enter slow inactivation, and their tardy recovery thereof, would prevent repetitive firing of APs at rates >1 Hz (35) typical for the heart. The potential arrhythmogenicity of skeletal muscle-like Na+ current properties in the heart is supported by the fact that naturally occurring cardiac Na+ channel mutations, which enhance slow inactivation, generate life-threatening arrhythmias (43, 46).

We found that prolonged treatment of C2C12 skeletal muscle cells with medium preconditioned by primary cardiocytes significantly alters basic functional properties of Na+ currents from skeletal muscle to more cardiac-like ones. Some of the results have been published previously in abstract form (48).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
A detailed description of our MATERIALS AND METHODS is given in a previous report by our group (49).

Cell culture and conditioning. Mouse C2C12 skeletal muscle cells (CRL-1772; American Type Culture Collection, Manassas, VA) were grown and differentiated on Matrigel (Becton Dickinson, Schwechat, Austria)-coated culture dishes as previously described (49). Cardiac primary cultures for the conditioning procedure were prepared from healthy neonatal Wistar rat (2–3 days old) hearts in accordance with a protocol described by Simpson and Savion (38) and coinciding with the rules of the University Animal Welfare Committee. Cardiocytes were prepared from rat rather than smaller mouse hearts to provide sufficient cell material for the conditioning procedure. Similarly, Iijima et al. (14) and Formigli et al. (10) used cocultures of rat cardiomyocytes and mouse myoblasts.

After thoracotomy, hearts were removed, placed in a culture dish containing ice-cold growth medium (GM), and freed from connective tissue under microscopic control. Cardiac tissue was mechanically dissociated in GM and repeatedly forced through the tip of a 10-ml pipette. Thereafter, the tissue homogenate was mixed 1:1 with a "solid tissue digestor" [RPMI 1640 medium, 10% fetal calf serum, 50 U/ml penicillin, 50 µg/ml streptomycin, 5 mM L-glutamine, 278 U/mg collagenase type I (Sigma, Vienna, Austria)] to isolate cardiocytes by proteolytic digestion. This mixture was shaken for 3 h at 37°C. Afterwards, the cell suspension was centrifuged (1,200 rpm, 5 min), and the pellet was resuspended in PBS. After another centrifugation step, the pellet was resuspended in GM and consecutively filtered through a cell strainer (40 µm). The filtrate was centrifuged, and the pellet was again resuspended in GM. Finally, this cell suspension was plated on Matrigel-coated dishes. Matrigel-coating was used to inhibit fibroblast proliferation (18) and support cardiomyocyte development (2). When ~80% cell confluence was reached, GM was replaced with differentiation medium (DM).

As soon as the rat cardiac primary cultures showed "beating areas" (normally after 24–48 h in DM), which contracted spontaneously in a repetitive manner, cardiac cellular supernatant (1.5 ml collected per 35-mm dish) was collected every 48 h and strained with a sterile filter to remove cellular debris. C2C12 cultures were conditioned with cardiac cellular supernatant as follows: 48 h after induction of differentiation in C2C12 cultures, the standard DM (2 ml per 3.5-mm dish) was replaced with a mixture of 0.5 ml of fresh DM and 1.5 ml of cardiac cellular supernatant. Fresh DM was applied to guarantee a supply of nutrients. The described medium transfer procedure was performed every 48 h and continued for 10–14 days. C2C12 cells that were treated in this manner are termed "cardiac-conditioned cells" throughout the text.

With the use of the identical procedure, other C2C12 cultures were treated with DM preconditioned by 100% confluent N1E neuroblastoma cell or cardiac fibroblast cultures. In some experiments, the cardiac cellular supernatant was boiled for 5 min before transfer onto C2C12 cells.

In parallel, using the same time schedule, every 48 h other C2C12 cultures were incubated in a mixture of 0.5 ml of fresh DM and 1.5 ml of DM that had been preconditioned by differentiated C2C12 cells for 48 h. Cells treated in this manner are termed "control cells" throughout the text. This procedure guaranteed that both control and conditioned cells were supplied with fresh and preconditioned DM and thus allowed for direct comparison.

For fibroblast-conditioning experiments, cardiac fibroblasts were enriched by preplating steps performed during standard primary cardiocyte preparation as described by Rohr et al. (33). Therefore, the cell suspension obtained at the end of the isolation procedure described was preplated for 90 min on uncoated culture dishes in a 37°C CO2 incubator. Thereafter, the supernatant (containing mainly cardiomyocytes) was removed and fresh GM was added. After 3–5 days, the fibroblasts formed uniform monolayers, at which time they were split and preplated for 90 min again. To further increase the purity of the fibroblast cultures, this procedure was repeated once more. Uncoated culture dishes were used during the whole culturing period because Matrigel-coating would have inhibited fibroblast proliferation (18). After reformation of uniform monolayers, GM was replaced with DM. Finally, these cultures were used to condition C2C12 cells as described above. Other fibroblast cultures were used for immunofluorescence experiments to confirm their purity.

Identical GM and DM were used for C2C12 skeletal muscle cells, rat primary cardiocytes, and other cell types: GM consisted of Dulbecco's modified Eagle's medium (Invitrogen, Lofer, Germany) containing 4.5 g/l glucose, 4 mM L-glutamine, 50 U/ml penicillin, 50 µg/ml streptomycin, and 20% fetal calf serum (PAA Labs, Pasching, Austria). DM was identical to GM except that it contained 2% horse serum (Invitrogen) instead of 20% fetal calf serum.

Immunofluorescence and immunoblotting. These experiments were performed as described previously (49).

Semiquantitative RT-PCR. Total cell RNA was extracted using RNeasy extraction kit (Qiagen, Hilden, Germany) and reverse transcribed with AMV reverse transcriptase and oligo(dT) primers (first-strand cDNA synthesis kit; Roche, Basel, Switzerland). Primers for the PCR reaction were designed in the 3' untranslated regions of the transcripts where the isoforms share almost no homology (skeletal muscle Nav1.4 forward, 5'-TGAAGATCCCGCCTCCTGA-3'; Nav1.4 reverse, 5'-AGTTTGTCTTTGTCCTGGCTA-3'; cardiac Nav1.5 forward, 5'-CGTGCCCTGTTGTATCCTG-3'; Nav1.5 reverse, 5'-CCAGCCAGGGTTGCTCGA-3'). These primers were used in a previous study by our group (49). PCR amplification of cDNAs (25 cycles, i.e., within the linear range of these PCRs) was performed with the Expand High Fidelity PCR System (Roche). A parallel reaction was performed with isolated total cell RNA omitting the reverse transcription step to demonstrate the absence of contaminating genomic DNA. Nav1.4 and Nav1.5 semiquantitative PCRs were performed with cDNA concentrations normalized to beta-actin expression. PCR products were visualized on a 2% agarose gel. Bands were quantified by densitometric analysis with Scion Image software (Frederick, MD).

Electrophysiology. Na+ currents were recorded from differentiated spherically shaped C2C12 cells (C2C12 myoballs) at room temperature (22 ± 1.5°C) using an Axoclamp 200B patch-clamp amplifier (Axon Instruments, Union City, CA) as described previously (49). Recording was begun ~10 min after whole cell access was attained to minimize time-dependent shifts in gating. Pipettes were formed from aluminosilicate glass (AF150-100-10; Science Products, Hofheim, Germany) with a P-97 horizontal puller (Sutter Instruments, Novato, CA), heat-polished on a microforge (MF-830; Narishige, Tokyo, Japan), and had resistances between 1 and 2 M{Omega} when filled with the recording pipette solution (105 mM CsF, 10 mM NaCl, 10 mM EGTA, and 10 mM HEPES, pH 7.3). Voltage-clamp protocols and data acquisition were performed with pCLAMP 6.0 software (Axon Instruments) through a 12-bit analog-to-digital/digital-to-analog interface (Digidata 1200; Axon Instruments). Data were low-pass filtered at 2 kHz (–3 dB) and digitized at 10–20 kHz. Curve fitting was performed using Origin 5.0 software (MicroCal Software, Northampton, MA). Current-voltage relationships were fit with the function Gmax x (x – Vrev) x [1 – (1/{1 + exp[(x – V0.5)/K]})], where Gmax is the maximum conductance, Vrev is the reversal potential, V0.5 is the voltage at which half-maximum activation occurred, and K is the slope factor. Na+ current density was calculated by dividing the maximum peak inward current amplitude (normally at –25 mV) of a cell by its membrane capacitance (49). Resting membrane potentials were measured using the whole cell patch-clamp technique in the current-clamp mode. Potential values were detected shortly (1–3 s) after the whole cell configuration was established in each experiment to minimize cell dialysis effects. In addition, large myoballs were chosen for these experiments, and pipettes with higher resistances [2–3 M{Omega} when filled with a KCl (130 mM)-based pipette solution] compared with those used for the Na+ current measurements were used. Steady-state inactivation data were fit with a Boltzmann function: 1/{1 + exp[(x – V0.5)/K]}, where V0.5 is the voltage at which half-maximum inactivation occurred and K is the slope factor. FractSI values, which represent the fraction of channels that are slow inactivated, were detected by measuring the respective smallest test pulse peak currents observed following 10-s inactivating prepulses between –40 and 0 mV (followed by 20-ms periods at –140 mV to allow for recovery from fast inactivation) in each experiment. Data from experiments performed to assess tetrodotoxin (TTX) sensitivity were fit with the "two-site binding" function: y = BFormulax x/(k1 + x) + BFormulax x/(k2 + x), where BFormula and BFormula are maximum binding capacity representing the relative contributions of a TTX-sensitive and a TTX-resistant Na+ channel fraction, respectively. k1 and k2 represent the respective 50% inhibitory constant (IC50) values. Recordings were made in a bathing solution that consisted of 140 mM NaCl, 2.5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES, pH 7.4. Some experiments were executed in low-Na+ bath solution containing 15 mM Na+ to minimize current amplitudes. In such experiments, 125 mM Na+ was substituted by the impermeant monovalent cation N-methyl-D-glucamine. Chemicals were purchased from Sigma. Rapid solution changes were performed with the use of a DAD-8-VC superfusion system (ALA Scientific Instruments, Westbury, NY).

Data are expressed as means ± SE. Statistical comparisons were made using two-tailed Student's unpaired t-tests if not otherwise specified. A P < 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Differentiation of control and cardiac-conditioned cells. Differentiation of C2C12 myoblasts was induced by serum reduction (49). After 7–8 days, the cells reached a well-differentiated state, and large longitudinal myotubes and spherical myoballs containing numerous nuclei were present. Figure 1, A and B, shows that control and cardiac-conditioned cells exhibited similar differentiation, as judged from their morphology. To further confirm this, protein levels of the developmental and the adult slow isoform of the myosin heavy chain (MHC) were compared in control and cardiac-conditioned cells (Fig. 1C). Decreased or increased levels of developmental versus adult slow MHC would indicate enhanced or attenuated differentiation, respectively. In contrast, protein levels of both MHC isoforms were similar in control and cardiac-conditioned cells (Fig. 1C), suggesting that differentiation was not affected by the conditioning procedure.


Figure 1
View larger version (82K):
[in this window]
[in a new window]

  		Fig. 1. Differentiation of control and cardiac-conditioned C2C12 cells. A: transmission images of typical control (left) and cardiac-conditioned cells (right) that had been differentiated for 14 days. Treatment with cardiac cellular supernatant lasted for 12 days. Bars, 40 µm. B: fast myosin heavy chain (MHC) isoforms were detected by immunofluorescence in control (left) and cardiac-conditioned cells (right) by using monoclonal MY32 antibody (anti-neonatal/fast MHC; Biomedica, Vienna, Austria) and secondary Alexa Fluor antibody (Eubio, Vienna, Austria). Bars, 40 µm. C: immunoblot analyses of developmental (MHCdev) and adult slow (MHCslow) MHC isoforms in control (C) and cardiac-conditioned cells (T). Four independent experiments (1–4) are shown. Antibodies NCL-MHCd (Novocastra, Vienna, Austria) and NOQ7.5.4D (Chemicon, Hofheim, Germany) were used to detect developmental and adult slow MHC isoforms, respectively.

 
Functional parameters of Na+ currents in control and cardiac-conditioned cells. Figure 2A shows typical original traces of Na+ currents elicited by various depolarizing voltage steps of a control and a cardiac-conditioned cell. Figure 2B gives a summary of the current-voltage relationships for control and cardiac-conditioned cells. To obtain quantitative parameters for comparison, these were fit with a function given in MATERIALS AND METHODS. Vrev, an indicator of channel ion selectivity, was not significantly different under control and experimental conditions (Table 1). In contrast, V0.5, the voltage at which half-maximum activation occurred, was significantly shifted to a more hyperpolarized value in cardiac-conditioned cells (Table 1). These results suggest that prolonged treatment of C2C12 cells with medium preconditioned by cardiocytes affects macroscopic Na+ current activation. Cardiac-conditioned cells activate at more hyperpolarized potentials.


Figure 2
View larger version (28K):
[in this window]
[in a new window]

  		Fig. 2. Effects of "cardiac milieu" on Na+ current activation and decay. From a holding potential of –120 mV, voltage steps to various potentials were applied (see inset at top). A: original traces of inward and outward currents elicited by such depolarizing voltage steps for a typical control (top traces) and cardiac-conditioned cell (bottom traces). To reduce current amplitudes, a low-Na+ bath solution (15 mM Na+) was used. B: a summary of the current-voltage relationships for control ({square}) and cardiac-conditioned cells (bullet). Solid lines connect the data points. C: typical examples of current decay following activation by a voltage step to –20 mV of a control ({square}) and a cardiac-conditioned cell (bullet) on an expanded time scale. Arrows indicate the time points at which the decay half-times were detected.

 

View this table:
[in this window]
[in a new window]

  		Table 1. Na+ current activation and decay parameters in control and cardiac-conditioned cells

 
The Na+ current densities of control and cardiac-conditioned cells were also estimated. The respective values obtained were 73 ± 12 (n = 8) and 77 ± 13 pA/pF (n = 11), and the values are not significantly different from each other. Similarly, no significant difference exists in the cell capacitance values, which amounted to 31 ± 3 pF (n = 8) in control and 37 ± 6 pF (n = 11) in cardiac-conditioned cells. The resting membrane potentials of control and cardiac-conditioned cells were –51 ± 5 (n = 8) and –49 ± 3 mV (n = 7), respectively, and also are not significantly different from each other.

Brief depolarizations (≤50 ms) cause inactivation of Na+ channels from which they recover with a single kinetic phase whose time constant is on the order of a few milliseconds. This inactivation process is called Na+ channel "fast inactivation." The kinetics of fast inactivation are represented by the speed of the current decay following channel activation. Figure 2A shows that the current decay was very rapid in control but markedly slowed in cardiac-conditioned C2C12 cells. A quantitative kinetic value was obtained by analyzing the time period between the current peak and the time point at which the current had decayed to 50% (decay half-time, see arrows in Fig. 2C). The decay half-times were significantly increased in cardiac-conditioned compared with control cells (Table 1). These data suggest that the kinetics of fast inactivation are significantly slowed by prolonged treatment of C2C12 cells with medium preconditioned by cardiocytes.

Figure 3, A and B, shows a comparison of the voltage dependencies of fast inactivation of control and cardiac-conditioned cells. The relationship of the cardiac-conditioned cells was shifted to more hyperpolarized voltages and became less steep (Fig. 3B). Statistical analysis revealed significant differences for both the voltage at which half-maximum fast inactivation occurred (V0.5) and the slope factor K (Table 2). These parameters were obtained by fitting the data with a Boltzmann equation (see MATERIALS AND METHODS). Thus the Na+ currents of cardiac-conditioned cells fast inactivated at more hyperpolarized voltages, and their fast inactivation process was less voltage dependent than that of control cells. These data suggest that prolonged treatment of C2C12 cells with medium preconditioned by cardiocytes significantly affects the voltage dependence of Na+ current fast inactivation.


Figure 3
View larger version (28K):
[in this window]
[in a new window]

  		Fig. 3. Effects of cardiac milieu on Na+ current fast and slow inactivation. A and B: voltage dependence of fast inactivation. From a holding potential of –120 mV, 50-ms inactivating prepulses to various potentials were applied (see inset in B). Typical inward currents elicited by immediately following test pulses of a control (top traces) and a cardiac-conditioned C2C12 cell (bottom traces) are shown in A. The current peaks were plotted against the prepulse voltage in B. This plot describes a summary of the voltage dependencies of steady-state fast inactivation of control ({square}) and cardiac-conditioned cells (bullet). Solid lines represent curve fits using a Boltzmann equation (see MATERIALS AND METHODS). C and D: voltage dependence of slow inactivation. From a holding potential of –120 mV, 10-s inactivating prepulses to various potentials were applied (see inset in D). Thereafter, the channels were allowed to recover from fast inactivation during a 20-ms period at –140 mV. Typical inward currents elicited by following test pulses of a control (top traces) and a cardiac-conditioned C2C12 cell (bottom traces) are shown in C. A summary of the voltage dependencies of slow inactivation of control ({square}) and cardiac-conditioned cells (bullet) is shown in D. Solid lines represent Boltzmann fits. For comparison of inactivation parameters, see Table 2.

 

View this table:
[in this window]
[in a new window]

  		Table 2. Fast and slow inactivation parameters in control and cardiac-conditioned cells

 
Prolonged depolarizations (seconds to minutes) cause inactivation of Na+ channels from which they recover with multiple kinetic phases whose time constants range over several orders of magnitude from tens of milliseconds up to tens of seconds. These kinetic phases of recovery can be summarized by the term "slow inactivation." Figure 3, C and D, shows a striking difference between the voltage dependencies of slow inactivation in control and cardiac-conditioned cells (Fig. 3D). In cardiac-conditioned cells, the fraction of channels that could be slow inactivated (fractSI) was markedly decreased compared with the corresponding fraction in control cells (Table 2). This indicates that the Na+ currents of cardiac-conditioned cells are more resistant to the process of slow inactivation. Moreover, the relationship describing the voltage dependence of slow inactivation was less steep in cardiac-conditioned than in control cells, as indicated by an increased slope factor K in Table 2. In addition, the voltage at which half-maximum slow inactivation occurred (V0.5) was shifted to a more hyperpolarized value in cardiac-conditioned cells (Table 2). These data suggest that the voltage dependence of Na+ current slow inactivation is significantly altered by prolonged treatment of C2C12 cells with medium preconditioned by cardiocytes.

All the experimental data from cardiac-conditioned C2C12 cells presented in Tables 1 and 2 were obtained after the cells had been exposed to a 10- to 14-day conditioning procedure. This comparatively long conditioning time period was chosen because the electrophysiological properties of C2C12 cells change during differentiation before finally reaching a "mature electrophysiological state" after ~8 days (4). To test whether changes in Na+ current properties also occur on a shorter time scale, we treated other C2C12 cultures for 3 days with medium preconditioned by cardiocytes. We then tested whether those Na+ current parameters which showed the most pronounced differences between "standard" control and cardiac-conditioned cells (fast inactivation kinetics and voltage dependence of slow inactivation) were also different between "short-term" control (for further explanation, see Table 3 legend) and cardiac-conditioned cells. The Na+ current parameters of short-term conditioned cells were significantly different from the control values except for the parameter V0.5 (Table 3). Moreover, comparison with the respective data given in Tables 1 and 2 reveals that short-term (3 days) cardiac conditioning generates changes in C2C12 cell Na+ current properties that are qualitatively similar to those of standard "long-term" (10–14 days) cardiac conditioning.


View this table:
[in this window]
[in a new window]

  		Table 3. Control experiments: Na+ current decay and slow inactivation parameters

 
To exclude the possibility that the described effects of cardiac conditioning on the Na+ current properties of C2C12 cells were simply due to medium depletion by primary cardiocytes, we treated other C2C12 cultures (10–14 days) with medium preconditioned by a different cell type, N1E neuroblastoma cells. We then tested whether, like cardiac conditioning, "N1E conditioning" was also effective. If so, an unspecific medium depletion effect, rather than a specific effect of cardiocytes, would become more likely. In contrast, the Na+ current parameters of N1E-conditioned cells (Table 3) were similar to those of control cells (see Tables 1 and 2), suggesting that medium depletion by N1E cells was ineffective. In a second attempt to exclude a medium depletion effect by cardiocytes, we performed conditioning experiments (10–14 days) using 10x concentrated cardiac-conditioned medium supplemented with appropriate amounts of fresh DM. We then tested whether this conditioning procedure (in the presence of a large quantity of fresh DM) generated effects on the Na+ current parameters of C2C12 cells similar to our standard cardiac-conditioning procedure (in the presence of a large quantity of preconditioned DM). This was indeed the case, as can be recognized by comparison of the respective data given in Table 3 with those given in Tables 1 and 2. Together, these results suggest that medium depletion effects can be neglected.

Cardiac fibroblasts play an important structural and functional role in the heart (5). These cells are, besides cardiomyocytes, also normally present in primary cardiac cell cultures in significant numbers. In principle, each of the two named cell types could cause the observed effects on the Na+ current properties of C2C12 cells. The immunofluorescence experiments shown in Fig. 4A suggest that, indeed, in our primary cardiocyte cultures, cardiomyocytes (a and c) and cardiac fibroblasts (b, d, and f) do coexist. Cardiomyocytes, however, seem to account for the majority of cells (compare a with b). To find out which of the two cell types was responsible for the observed conditioning effects, we generated cardiac fibroblast cultures (Fig. 4B) and treated other C2C12 cultures for 10–14 days with medium preconditioned by fibroblasts. We then tested whether "fibroblast conditioning" affected the Na+ current parameters of C2C12 cells in a manner similar to cardiac conditioning. We found no significant difference in any Na+ current parameter between fibroblast-conditioned (Table 3) and control cells (Tables 1 and 2), suggesting that cardiac fibroblasts alone were ineffective. Thus cardiomyocytes remain as the origin for the effects of cardiac conditioning.


Figure 4
View larger version (111K):
[in this window]
[in a new window]

  		Fig. 4. Cardiomyocytes and cardiac fibroblasts in cardiac primary cultures. A: the cardiomyocyte marker beta-MHC (slow; a and c) and the fibroblast marker vimentin (b and d) were detected by immunofluorescence in an ~80% confluent standard cardiocyte culture 12 h after addition of differentiation medium (DM) with the antibodies NOQ7.5.4D (Chemicon) and anti-vimentin (V5255, Sigma), respectively. Antibody Alexa Fluor (Eubio) was used as secondary antibody. A typical cardiocyte culture 7 days after addition of DM is shown in e (transmission image) and f (vimentin staining). Bars: a and b, 300 µm; c and d, 50 µm; e and f, 1,250 µm. B: transmission image (a), as well as an appendant overview (b) and detail of vimentin staining (c) of a cardiac fibroblast culture purified by 3 preplating steps (see MATERIALS AND METHODS). beta-MHC staining failed to produce any specific signal exceeding the control signal (cells treated only with the secondary antibody) in such cultures (data not shown). This suggests that only fibroblasts were present. Bars: a and b, 300 µm; c, 50 µm.

 
Additional experiments were performed to test whether proteins contained in the medium preconditioned by cardiocytes were responsible for the described effects. Cardiac cellular supernatant was boiled to denature proteins before transfer onto C2C12 cells. The Na+ current parameters of such cardiac-conditioned cells (Table 3) were similar to those of control cells (see Tables 1 and 2), suggesting that the effect of cardiac conditioning was lost after protein denaturation.

Na+ channel isoform expression in control and cardiac-conditioned cells. Comparison of the Na+ current parameters in our control and cardiac-conditioned cells (Tables 1 and 2) with studies using heterologous expression systems to investigate functional differences between skeletal muscle Nav1.4 and cardiac Nav1.5 Na+ channels (24, 31, 36, 45) suggests that cardiac conditioning generates more cardiac-like Na+ current properties. Namely, these authors reported channel activation (36, 45), as well as fast and slow inactivation (24, 45), to occur at more hyperpolarized potentials in Nav1.5 compared with Nav1.4 channels. In addition, Nav1.5 channels exhibited slowed kinetics of fast inactivation (24, 45), a reduced voltage dependence of slow inactivation (24), and a higher resistance to undergo slow inactivation (24, 31). All these Nav1.4-to-Nav1.5 shifts in Na+ current parameters were also found in our cardiac-conditioned compared with control cells (compare values in Tables 1 and 2). This suggested that an upregulation of the expression of Nav1.5 versus Nav1.4 channels could be responsible for cardiac conditioning-induced Na+ current modulation. To test for the expected switch in Na+ channel isoform expression, we compared the TTX sensitivities of Na+ currents in control and cardiac-conditioned cells. It is well known that the currents through Nav1.5 channels are more resistant to TTX block than Nav1.4 channels; thus a shift in Na+ channel isoform expression from Nav1.4 toward Nav1.5 should result in a reduced TTX sensitivity. Figure 5, A–C, shows that the currents of cardiac-conditioned cells indeed exhibited a reduced TTX sensitivity. Two distinct populations of Na+ channels, a TTX-sensitive and a TTX-resistant fraction, could be clearly separated by fitting the data with a two-site binding function (e.g., Fig. 5C). As shown in Table 4, the IC50 values of Na+ channel block by TTX of control and cardiac-conditioned cells were similar. This was true for both the TTX-sensitive (IC50 ~ 20 nM) as well as the TTX-resistant (IC50 ~ 1–1.5 µM) channel fraction. These values compare well with the IC50 values of Nav1.4 and Nav1.5 channels found in the literature (e.g., Refs. 47, 50). In contrast to the IC50 values, the respective relative contributions of the two channel fractions were significantly different from each other. Thus the TTX-sensitive Na+ channel fraction in control cells amounted to 80% of the total channel fraction, whereas the corresponding value in cardiac-conditioned cells was reduced to 47% (Table 4). These results strongly suggest a shift in Na+ channel isoform expression from skeletal muscle Nav1.4 toward cardiac Nav1.5. To confirm these data, we performed semiquantitative RT-PCR experiments. Figure 5D shows that cardiac conditioning significantly decreased and increased the amounts of PCR product of Nav1.4 and Nav1.5, respectively. This was true for each individual experiment performed (n = 8) and suggests downregulation of Nav1.4 and upregulation of Nav1.5 expression, consistent with our TTX experiments.


Figure 5
View larger version (23K):
[in this window]
[in a new window]

  		Fig. 5. Effects of cardiac milieu on Na+ channel isoform expression. A: typical Na+ currents of a control (top traces) and cardiac-conditioned cell (bottom traces) elicited by 10-ms test pulses from –120 to –20 mV in the absence (large currents) or presence (small currents) of 1 µM tetrodotoxin (TTX). B: a summary of the TTX sensitivities of Na+ currents in control ({square}) and cardiac-conditioned C2C12 cells (bullet). The current peak amplitudes in the presence of TTX (1 nM–20 µM) were normalized to the respective amplitude in the absence of TTX and plotted against TTX concentration. C: plot of current inhibition by TTX in a typical control ({square}) and cardiac-conditioned cell (bullet). Solid lines show free fits of the data with a "two-site binding" function (see MATERIALS AND METHODS). The inset data give the respective fitting parameters (top, control; bottom, conditioned). D: semiquantitative RT-PCR of Na+ channel isoforms Nav1.4 and Nav1.5 in control and cardiac-conditioned cells. Top, representative agarose gel of RT-PCR products from Nav1.4, Nav1.5, and corresponding beta-actin mRNA amplification; bottom, summary of densitometric analyses executed on conditioned cell bands normalized to beta-actin expression. Data are presented as percentages of control cell bands (100%) (means ± SE; n = 8 for both Nav1.4 and Nav1.5). *P < 0.05, significant difference from the control (by 2-tailed Wilcoxon signed rank test performed with beta-actin-corrected raw data).

 

View this table:
[in this window]
[in a new window]

  		Table 4. TTX sensitivity of Na+ currents in control and cardiac-conditioned cells

 
Na+ current parameters in cardiac-conditioned cells after selective block of Nav1.4 channels. To further study the effect of Na+ channel isoform expression on Na+ current function in C2C12 cells, we superfused cardiac-conditioned cells with 1 µM TTX and detected their Na+ current parameters. TTX (1 µM) should completely eliminate Nav1.4 currents (IC50 ~ 20 nM, Table 4) but leave a considerable amount of Nav1.5 current (IC50 ~ 1–1.5 µM) and thus allow us to selectively study Nav1.5 Na+ current properties. Cardiac-conditioned rather than control cells were selected for these experiments, because only they exhibited sufficient Na+ current during TTX superfusion (see Fig. 5A). We found that TTX treatment significantly shifted Na+ current activation (Table 1) as well as fast and slow inactivation (Table 2) of conditioned cells to even more hyperpolarized voltages. In addition, the Na+ currents showed slowed fast inactivation kinetics (Table 1) and a higher resistance to undergo slow inactivation (Table 2). The values of all these Na+ current parameters of cardiac-conditioned cells lay in between those of control and conditioned cells treated with TTX. This is consistent with the idea that the Na+ currents of cardiac-conditioned cells were partly carried by Nav1.4 and Nav1.5 channels.

The coexistence of two Na+ channel populations in cardiac-conditioned cells is further confirmed by a more detailed comparison of the voltage dependencies of fast and slow inactivation in control, conditioned, and TTX-treated conditioned cells. Filatov and Rich (8) modeled the effects of various contributions of Nav1.4 and Nav1.5 channels on the voltage dependence of inactivation. They predicted that increasing the percentage of Nav1.5 gradually shifts the inactivation voltage dependence toward hyperpolarized voltages. A similar shift can be observed in our data (Table 2), consistent with an increased Nav1.5 channel fraction in cardiac-conditioned versus control cells and the highest Nav1.5 channel fraction in TTX-treated conditioned cells. Moreover, a mixed population of Nav1.4 and Nav1.5 channels should also reduce the slope of the voltage dependence of inactivation (8). Indeed, this was the case for both fast and slow inactivation in cardiac-conditioned compared with control cells (compare K values in Table 2). Together, these results strongly suggest that an increased fraction of Nav1.5 channels accounts for the Na+ current adaptations induced by cardiac conditioning.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In the present study, we have shown that residence of C2C12 skeletal muscle cells in a cardiac milieu shifts their functional Na+ current properties from skeletal muscle toward more cardiac-like properties. This is most likely achieved by an upregulation of expression of Nav1.5 versus Nav1.4 channels, both of which have previously been shown to be expressed in differentiated C2C12 cells (49), as well as in adult innervated human skeletal muscle (9). The experiments were performed on mouse C2C12 skeletal muscle cells, since these exhibit developmental regulation in the expression of voltage-gated ion channels much like that of skeletal muscle cells in vivo (4) and have been successfully used in mouse models of intracardiac myoblast transplantation (e.g., Refs. 16, 29, 32).

Electrophysiological adaptations of skeletal muscle cells in a cardiac cell environment. Leobon et al. (17) and Ott et al. (25) recently attempted to trace possible electrophysiological adaptations of skeletal myoblasts in a cardiac cell environment. In both studies, a myocardial infarction was artificially induced in rodent hearts, and genetically labeled myoblasts were transplanted into the infarcted myocardium, where they were allowed to reside for a certain time period before electrophysiological recordings were performed on these cells. Leobon et al. (17) reported that the grafted myoblasts differentiated into peculiar hyperexcitable myotubes very different from cardiomyocytes, whereas Ott et al. (25) found a decreased voltage-gated ion channel expression in the grafted myoblasts. In line with our results, these studies suggest that skeletal muscle cells do indeed change their electrophysiological properties in a cardiac cell environment. However, the electrophysiological changes reported by these authors are basically different from our findings. In contrast to Leobon et al. (17) and Ott et al. (25), we found more cardiac-like Na+ current properties and no significant change in Na+ current density after residence of skeletal muscle cells in a cardiac milieu, respectively. The reason for these differences is unknown and can only be speculated upon. First, we used a simple in vitro approach to simulate a cardiac cell environment for cultured skeletal muscle cells, whereas the two other studies were more complex in vivo investigations. Thus it is possible that unknown factors that were absent in our in vitro system cause different alterations in the electrophysiological properties of skeletal muscle cells in a cardiac milieu in vivo. This is supported by another in vitro study by Iijima et al. (14), who cocultured skeletal muscle cells with cardiomyocytes and, consistent with our data, found skeletal muscle-derived cells that showed cardiomyocyte-like APs. Second, electrophysiological adaptations of skeletal muscle cells may be basically different in a "healthy" cardiac milieu (present study; Ref. 14) and in the complex environment produced by an infarcted myocardium (17, 25). Alterations in Na+ channel kinetics and expression occur in surviving cardiomyocytes after a myocardial infarction (13), which creates a pathophysiological environment possibly also affecting skeletal muscle cell electrophysiology. For example, proinflammatory cytokines in the infarction area and/or hypoxia and related changes in cell metabolism or gene expression may produce effects. Future studies are required to clarify whether myoblasts transplanted into healthy myocardium undergo electrophysiological adaptations in vivo similar to those of our C2C12 cells in vitro in a simulated healthy cardiac milieu.

Cardiomyocytes alter the Na+ current properties of skeletal muscle cells via a paracrine mechanism. Our finding that medium preconditioned by primary cardiocytes increased the expression of Nav1.5 versus Nav1.4 in C2C12 cells may be caused by different mechanisms. First, during the conditioning procedure, cardiocytes may deplete medium nutrients, which could produce effects. Such a simple medium depletion effect seems highly unlikely, however, because our C2C12 cells were treated with a mixture of preconditioned and fresh medium. In addition, medium preconditioned by N1E neuroblastoma cells, which should also deplete the medium, did not produce any effects. Moreover, experiments with highly concentrated cardiac-conditioned medium, which allowed conditioning in the presence of a large quantity of fresh medium, produced effects similar to those of standard cardiac conditioning.

Second, medium preconditioned by cardiocytes may inhibit myoblast differentiation. It is well known that immature skeletal muscle cells express a considerable fraction of Nav1.5 channels (15), and in culture, this fraction is continuously decreased as cells mature during differentiation. In Fig. 1 we show that myoblast differentiation is not inhibited by the conditioning procedure. These data were recently confirmed by Pedrotty et al. (26), who used a similar approach and reported that cardiac milieu does not affect differentiation of skeletal myoblasts. From this, we conclude that the effects on Na+ channel expression reported in the present study were not simply caused by inhibited differentiation of skeletal myoblasts in a cardiac milieu.

Third, since neither medium depletion nor inhibited myoblast differentiation can account for the observed effects, we propose a paracrine mechanism: cardiocytes secrete a single factor or several factors that modulate Na+ channel isoform expression in skeletal muscle cells. Between cardiomyocytes and cardiac fibroblasts, we could identify the former, but not the latter, cell type to be effective. To our knowledge, this is the first experimental evidence suggesting that cardiomyocytes modulate an electrophysiological property of skeletal muscle cells via paracrine action. Importantly, a paracrine role for the heart recently was suggested, also in vivo (3), by which stem cell differentiation into cardiomyocytes is driven. Moreover, Abraham et al. (1) proposed a paracrine effect of skeletal myoblasts on cardiomyocytes. Medium preconditioned by skeletal myoblasts prolonged AP duration in cardiomyocytes.

Currently, the identity of the cardiac factor(s) affecting Na+ channel expression of skeletal muscle cells is unknown. As a first step to unravel its (their) nature, we boiled the medium preconditioned by cardiocytes before transfer onto C2C12 cells. This procedure should denature proteins and thus allow us to judge whether our effects are brought about by the action of proteins. The cooking procedure completely eliminated the effects of cardiac conditioning. This suggests that proteins are indeed responsible for the observed effects. Future studies are necessary to clarify their identity. Growth factors may be considered possible candidates. In C2C12 cells, ion channel expression was shown to be affected by the addition of a single mitogen (TGF-beta1) to the culture medium (4).

Future perspectives. The identification of factors that induce cardiac ion channel expression in skeletal muscle cells may open the possibility to improve both efficacy and safety of intracardiac myoblast transplantation therapy. Such factors, if coinjected in the course of myoblast transplantation and/or applied in the initial postoperative weeks, may result in cardiac ion channel expression in the skeletal muscle graft, a prerequisite for proper cardiac-like impulse conduction and contraction. Alternatively, it may be possible to boost the endogenous release of such factors by cardiocytes. To substantially improve cardiac contractility, the skeletal muscle graft should contract in synchrony with host cardiac tissue. This requires both electrical coupling between skeletal muscle cells and cardiomyocytes and cardiac ion channel function in skeletal muscle cells (31, 35). Electrical coupling does occur in vitro (10, 14, 28) but has seriously been questioned in vivo (16, 17, 34). This problem can possibly be overcome by genetic modification of skeletal muscle cells to overexpress the cardiac gap junction protein connexin43 (30, 40), which may permit electrical coupling to cardiomyocytes (30, 39). Notably, such an induced coupling may only be beneficial for the heart if the skeletal muscle graft indeed exhibits cardiac-like ion channel properties. Coupling to grafts with more skeletal muscle- or neuron-like electrophysiology (17), in contrast, may even be deleterious for the heart and generate arrhythmias. Future studies are needed to elucidate the potential of "antiarrhythmic engineering" (1) of skeletal muscle cells to improve the success of myoblast transplantation therapy.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Austrian Fonds zur Förderung der Wissenschaftlichen Forschung Grant P-15063 (to K. Hilber).


   ACKNOWLEDGMENTS
 
We thank Johannes Schmid and Lukas Weigl for helpful scientific comments.

Present address of W. Sandtner: Department of Anesthesiology, Division of Molecular Medicine, David Geffen School of Medicine, UCLA, BH-553 CHS, 650 Charles E. Young Drive, Los Angeles, CA 90095.


   FOOTNOTES
 

Address for reprint requests and other correspondence: K. Hilber, Center of Biomolecular Medicine and Pharmacology, Institute of Pharmacology, Medical Univ. of Vienna, Waehringerstrasse 13A, A-1090 Vienna, Austria (e-mail: karlheinz.hilber{at}meduniwien.ac.at)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Abraham MR, Henrikson CA, Tung L, Chang MG, Aon M, Xue T, Li RA, O' Rourke B, Marban E. Antiarrhythmic engineering of skeletal myoblasts for cardiac transplantation. Circ Res 97: 159–167, 2005.[Abstract/Free Full Text]
  2. Baharvand H, Azarnia M, Parivar K, Ashtiani SK. The effect of extracellular matrix on embryonic stem cell-derived cardiomyocytes. J Mol Cell Cardiol 38: 495–503, 2005.[CrossRef][Web of Science][Medline]
  3. Behfar A, Zingman LV, Hodgson DM, Rauzier JM, Kane GC, Terzic A, Puceat M. Stem cell differentiation requires a paracrine pathway in the heart. FASEB J 16: 1558–1566, 2002.[Abstract/Free Full Text]
  4. Caffrey JM, Brown AM, Schneider MD. Ca2+ and Na+ currents in developing skeletal myoblasts are expressed in a sequential program: reversible suppression by transforming growth factor beta-1, an inhibitor of the myogenic pathway. J Neurosci 9: 3443–3453, 1989.[Abstract]
  5. Camelliti P, Borg TK, Kohl P. Structural and functional characterisation of cardiac fibroblasts. Cardiovasc Res 65: 40–51, 2005.[Abstract/Free Full Text]
  6. Dib N, Michler RE, Pagani FD, Wright S, Kereiakes DJ, Lengerich R, Binkley P, Buchele D, Anand I, Swingen C, Di Carli MF, Thomas JD, Jaber WA, Opie SR, Campbell A, McCarthy P, Yeager M, Dilsizian V, Griffith BP, Korn R, Kreuger SK, Ghazoul M, MacLellan WR, Fonarow G, Eisen HJ, Dinsmore J, Diethrich E. Safety and feasibility of autologous myoblast transplantation in patients with ischemic cardiomyopathy: four-year follow-up. Circulation 112: 1748–1755, 2005.
  7. Fernandes S, Amirault JC, Lande G, Nguyen JM, Forest V, Bignolais O, Lamirault G, Heudes D, Orsonneau JL, Heymann MF, Charpentier F, Lemarchand P. Autologous myoblast transplantation after myocardial infarction increases the inducibility of ventricular arrhythmias. Cardiovasc Res 69: 348–358, 2006.[Abstract/Free Full Text]
  8. Filatov GN, Rich MM. Hyperpolarized shifts in the voltage dependence of fast inactivation of Nav1.4 and Nav1.5 in a rat model of critical illness myopathy. J Physiol 559: 813–820, 2004.[Abstract/Free Full Text]
  9. Fletcher JE, Wieland SJ, Karan SM, Beech J, Rosenberg H. Sodium channel in human malignant hyperthermia. Anesthesiology 86: 1023–1032, 1997.[Web of Science][Medline]
  10. Formigli L, Francini F, Tani A, Squecco R, Nosi D, Polidori L, Nistri S, Chiappini L, Cesati V, Pacini A, Perna AM, Orlandini GE, Zecchi OS, Bani D. Morphofunctional integration between skeletal myoblasts and adult cardiomyocytes in coculture is favored by direct cell-cell contacts and relaxin treatment. Am J Physiol Cell Physiol 288: C795–C804, 2005.[Abstract/Free Full Text]
  11. Hagege AA, Carrion C, Menasche P, Vilquin JT, Duboc D, Marolleau JP, Desnos M, Bruneval P. Viability and differentiation of autologous skeletal myoblast grafts in ischaemic cardiomyopathy. Lancet 361: 491–492, 2003.[CrossRef][Web of Science][Medline]
  12. Herreros J, Prosper F, Perez A, Gavira JJ, Garcia-Velloso MJ, Barba J, Sanchez PL, Canizo C, Rabago G, Marti-Climent JM, Hernandez M, Lopez-Holgado N, Gonzalez-Santos JM, Martin-Luengo C, Alegria E. Autologous intramyocardial injection of cultured skeletal muscle-derived stem cells in patients with non-acute myocardial infarction. Eur Heart J 24: 2012–2020, 2003.[Abstract/Free Full Text]
  13. Huang B, El Sherif T, Gidh-Jain M, Qin D, El Sherif N. Alterations of sodium channel kinetics and gene expression in the postinfarction remodeled myocardium. J Cardiovasc Electrophysiol 12: 218–225, 2001.[CrossRef][Web of Science][Medline]
  14. Iijima Y, Nagai T, Mizukami M, Matsuura K, Ogura T, Wada H, Toko H, Akazawa H, Takano H, Nakaya H, Komuro I. Beating is necessary for transdifferentiation of skeletal muscle-derived cells into cardiomyocytes. FASEB J 17: 1361–1363, 2003.[Abstract/Free Full Text]
  15. Kallen RG, Sheng ZH, Yang J, Chen LQ, Rogart RB, Barchi RL. Primary structure and expression of a sodium channel characteristic of denervated and immature rat skeletal muscle. Neuron 4: 233–242, 1990.[CrossRef][Web of Science][Medline]
  16. Koh GY, Klug MG, Soonpaa MH, Field LJ. Differentiation and long-term survival of C2C12 myoblast grafts in heart. J Clin Invest 92: 1548–1554, 1993.[Web of Science][Medline]
  17. Leobon B, Garcin I, Menasche P, Vilquin JT, Audinat E, Charpak S. Myoblasts transplanted into rat infarcted myocardium are functionally isolated from their host. Proc Natl Acad Sci USA 100: 7808–7811, 2003.[Abstract/Free Full Text]
  18. Lyles JM, Amin W, Weill CL. Matrigel enhances myotube development in a serum-free defined medium. Int J Dev Neurosci 10: 59–73, 1992.[CrossRef][Web of Science][Medline]
  19. Menasche P. Myoblast transplantation: feasibility, safety and efficacy. Ann Med 34: 314–315, 2002.[CrossRef][Web of Science][Medline]
  20. Menasche P. Skeletal muscle satellite cell transplantation. Cardiovasc Res 58: 351–357, 2003.[CrossRef][Web of Science][Medline]
  21. Menasche P. Skeletal myoblast for cell therapy. Coron Artery Dis 16: 105–110, 2005.[CrossRef][Web of Science][Medline]
  22. Menasche P, Hagege AA, Vilquin JT, Desnos M, Abergel E, Pouzet B, Bel A, Sarateanu S, Scorsin M, Schwartz K, Bruneval P, Benbunan M, Marolleau JP, Duboc D. Autologous skeletal myoblast transplantation for severe postinfarction left ventricular dysfunction. J Am Coll Cardiol 41: 1078–1083, 2003.[Abstract/Free Full Text]
  23. Murry CE, Wiseman RW, Schwartz SM, Hauschka SD. Skeletal myoblast transplantation for repair of myocardial necrosis. J Clin Invest 98: 2512–2523, 1996.[Web of Science][Medline]
  24. O'Reilly JP, Wang SY, Kallen RG, Wang GK. Comparison of slow inactivation in human heart and rat skeletal muscle Na+ channel chimaeras. J Physiol 515.1: 61–73, 1999.
  25. Ott HC, Berjukow S, Marksteiner R, Margreiter E, Bock G, Laufer G, Hering S. On the fate of skeletal myoblasts in a cardiac environment: down-regulation of voltage-gated ion channels. J Physiol 558: 793–805, 2004.[Abstract/Free Full Text]
  26. Pedrotty DM, Koh J, Davis BH, Taylor DA, Wolf P, Niklason LE. Engineering skeletal myoblasts: roles of three-dimensional culture and electrical stimulation. Am J Physiol Heart Circ Physiol 288: H1620–H1626, 2005.[Abstract/Free Full Text]
  27. Peters NS. Arrhythmias after cell transplantation for myocardial regeneration: natural history or result of the intervention? J Cardiovasc Electrophysiol 16: 1255–1257, 2005.[CrossRef][Web of Science][Medline]
  28. Reinecke H, MacDonald GH, Hauschka SD, Murry CE. Electromechanical coupling between skeletal and cardiac muscle. Implications for infarct repair. J Cell Biol 149: 731–740, 2000.[Abstract/Free Full Text]
  29. Reinecke H, Minami E, Poppa V, Murry CE. Evidence for fusion between cardiac and skeletal muscle cells. Circ Res 94: E56–E60, 2004.
  30. Reinecke H, Minami E, Virag JI, Murry CE. Gene transfer of connexin43 into skeletal muscle. Hum Gene Ther 15: 627–636, 2004.[CrossRef][Web of Science][Medline]
  31. Richmond JE, Featherstone DE, Hartmann HA, Ruben PC. Slow inactivation in human cardiac sodium channels. Biophys J 74: 2945–2952, 1998.
  32. Robinson SW, Cho PW, Levitsky HI, Olson JL, Hruban RH, Acker MA, Kessler PD. Arterial delivery of genetically labelled skeletal myoblasts to the murine heart: long-term survival and phenotypic modification of implanted myoblasts. Cell Transplant 5: 77–91, 1996.[Web of Science][Medline]
  33. Rohr S, Fluckiger-Labrada R, Kucera JP. Photolithographically defined deposition of attachment factors as a versatile method for patterning the growth of different cell types in culture. Pflügers Arch 446: 125–132, 2003.[Web of Science][Medline]
  34. Rubart M, Soonpaa MH, Nakajima H, Field LJ. Spontaneous and evoked intracellular calcium transients in donor-derived myocytes following intracardiac myoblast transplantation. J Clin Invest 114: 775–783, 2004.[CrossRef][Web of Science][Medline]
  35. Ruff RL. Cells use the singular properties of different channels to produce unique electrical songs. Biophys J 74: 2745–2746, 1998.
  36. Sheets MF, Hanck DA. Gating of skeletal and cardiac muscle sodium channels in mammalian cells. J Physiol 514: 425–436, 1999.[Abstract/Free Full Text]
  37. Siminiak T, Kalawski R, Fiszer D, Jerzykowska O, Rzezniczak J, Rozwadowska N, Kurpisz M. Autologous skeletal myoblast transplantation for the treatment of postinfarction myocardial injury: phase I clinical study with 12 months of follow-up. Am Heart J 148: 531–537, 2004.[CrossRef][Web of Science][Medline]
  38. Simpson P, Savion S. Differentiation of rat myocytes in single cell cultures with and without proliferating nonmyocardial cells. Cross-striations, ultrastructure, and chronotropic response to isoproterenol. Circ Res 50: 101–116, 1982.[Free Full Text]
  39. Stagg MA, Coppen SR, Suzuki K, Varela-Carver A, Lee J, Brand NJ, Fukushima S, Yacoub MH, Terracciano CM. Evaluation of frequency, type, and function of gap junctions between skeletal myoblasts overexpressing connexin43 and cardiomyocytes: relevance to cell transplantation. FASEB J 20: 744–746, 2006.[Abstract/Free Full Text]
  40. Suzuki K, Brand NJ, Allen S, Khan MA, Farrell AO, Murtuza B, Oakley RE, Yacoub MH. Overexpression of connexin43 in skeletal myoblasts: relevance to cell transplantation to the heart. J Thorac Cardiovasc Surg 122: 759–766, 2001.[Abstract/Free Full Text]
  41. Taylor DA. Cell-based myocardial repair: how should we proceed? Int J Cardiol 95, Suppl 1: S8–S12, 2004.
  42. Van den Bos EJ, Davis BH, Taylor DA. Transplantation of skeletal myoblasts for cardiac repair. J Heart Lung Transplant 23: 1217–1227, 2004.[CrossRef][Web of Science][Medline]
  43. Veldkamp MW, Viswanathan PC, Bezzina C, Baartscheer A, Wilde AA, Balser JR. Two distinct congenital arrhythmias evoked by a multidysfunctional Na+ channel. Circ Res 86: E91–E97, 2000.
  44. Vilin YY, Makita N, George AL Jr, Ruben PC. Structural determinants of slow inactivation in human cardiac and skeletal muscle sodium channels. Biophys J 77: 1384–1393, 1999.
  45. Wang DW, George AL Jr, Bennett PB. Comparison of heterologously expressed human cardiac and skeletal muscle sodium channels. Biophys J 70: 238–245, 1996.
  46. Wang DW, Makita N, Kitabatake A, Balser JR, George AL Jr. Enhanced Na+ channel intermediate inactivation in Brugada syndrome. Circ Res 87: E37–E43, 2000.
  47. Weiss RE, Horn R. Functional differences between two classes of sodium channels in developing rat skeletal muscle. Science 233: 361–364, 1986.[Abstract/Free Full Text]
  48. Zebedin E, Mille M, Modarressy-Yazdi K, Speiser M, Sandtner W, Szendroedi J, Zarrabi T, Todt H, Hilber K. On the electrophysiological fate of skeletal muscle cells in a cardiac cell environment (Abstract). J Muscle Res Cell Motil 25: P270, 2004.
  49. Zebedin E, Sandtner W, Galler S, Szendroedi J, Just H, Todt H, Hilber K. Fiber type conversion alters inactivation of voltage-dependent sodium currents in mouse C2C12 skeletal muscle cells. Am J Physiol Cell Physiol 287: C270–C280, 2004.[Abstract/Free Full Text]
  50. Zimmer T, Bollensdorff C, Haufe V, Birch-Hirschfeld E, Benndorf K. Mouse heart Na+ channels: primary structure and function of two isoforms and alternatively spliced variants. Am J Physiol Heart Circ Physiol 282: H1007–H1017, 2002.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 CirculationHome page
H. Q. Ly and S. Nattel
Stem Cells Are Not Proarrhythmic: Letting the Genie out of the Bottle
Circulation, April 7, 2009; 119(13): 1824 - 1831.
[Full Text] [PDF]

Home page
 Cardiovasc ResHome page
R. Martinez-Marmol, M. David, R. Sanches, M. Roura-Ferrer, N. Villalonga, E. Sorianello, S. M. Webb, A. Zorzano, A. Guma, C. Valenzuela, et al.
Voltage-dependent Na+ channel phenotype changes in myoblasts. Consequences for cardiac repair
Cardiovasc Res, December 1, 2007; 76(3): 430 - 441.
[Abstract] [Full Text] [PDF]

This Article
Free upon publication Free Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
292/1/H439    most recent
00119.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (4)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Zebedin, E.
Right arrow Articles by Hilber, K.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Zebedin, E.
Right arrow Articles by Hilber, K.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.