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Neuroscience |
1 Dipartimento Scienze del Farmaco
3 Centro Universitario Medicina dello Sport, Istituto Interuniversitario di Miologia, Ce.S.I. Centro di Scienze dell'Invecchiamento ‘University G. d'Annunzio’ Foundation, via dei Vestini, I-66013 Chieti, Italy
2 Dipartimento di Biologia Cellulare e Ambientale, Università di Perugia, via Pascoli 1, I-06123 Perugia, Italy
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(Received 22 November 2005;
accepted after revision 31 January 2006;
first published online 2 February 2006)
Corresponding author T. Pietrangelo: Ce.S.I. (Centro di Scienze dell'Invecchiamento), ‘Università G. d'Annunzio’ Foundation, Via Colle dell'Ara, I-66013 Chieti, Italy. Email: tiziana{at}unich.it
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Introduction |
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The physiological role of IKCa channels in myogenic cell lines is currently a matter of debate. In the 10T1/2-MFR-4 myogenic cell line, IKCa channels have been found to be up-regulated by the growth factor bFGF, and involved in the bFGF-induced proliferation (Peña et al. 2000). However, the same study shows that TGF-ß, which likewise up-regulates the IKCa channel, although to a smaller extent, does not stimulate cell proliferation, suggesting that the IKCa channel up-regulation does not necessarily induce proliferation. On the other hand, the IKCa channel up-regulation induced by both bFGF and TGF-ß appeared to inhibit myogenesis induced by these growth factors (Peña & Rane, 1997; Peña et al. 2000). We recently showed that IKCa channels in C2C12 myoblasts are down-regulated during myogenesis, but are not involved in the bFGF-induced proliferation (Fioretti et al. 2005).
Following the role of extracellular purines (mainly ATP-based) as neurotransmitters and modulators in the nervous system (see Burnstock, 2003), more recently a number of investigators have identified extracellular guanine-based purines (GTP, GDP, GMP and guanosine) as trophic and mitogenic factors in neuron and glial compartments (Neary et al. 1996; Ciccarelli et al. 1999; Rathbone et al. 1999; Guarnieri et al. 2004).
In muscular tissue, extracellular GTP affects different activities, such as the development of isometric twitch tension (Mancinelli et al. 1983) or contractility of muscles involved in the backward swimming of Paramecium tetraurelia (Clark et al. 1997; Mimikakis & Nelson, 1998). In vitro studies on C2C12, a skeletal muscle cell line widely used as a myogenic model, show that external GTP promotes a significant increase in intracellular calcium ([Ca2+]i) via two different mechanisms involving specific sites in the cell membrane (Pietrangelo et al. 2002). Indeed, two specific sites have been identified in C2C12 myoblasts: the high affinity GTP-binding site (Kd1= 15.4 ± 4.6 µM) and the low affinity GTP-binding site (Kd2= 170.0 ± 94.5 µM).
In this report we examine whether the GTP-induced [Ca2+]i increase promotes a significant IKCa channel activation, and whether this functional coupling has a role in the proliferation or differentiation of C2C12 myoblasts. We show that a GTP-dependent [Ca2+]i increase induces a significant membrane hyperpolarization through the activation of IKCa channels.
We additionally demonstrate that early GTP-induced transductive events are involved in the myogenesis process. GTP arrests the slow proliferation rate, dependent on the early phase of muscle differentiatiation, and promotes the accumulation of skeletal muscle heavy chain protein (MyHC), a specific and terminal marker of muscle differentiation. Moreover, ChTX, a known IKCa channel blocker, abolishes this effect at a concentration of 200 nM, suggesting a close relationship between GTP-dependent IKCa activation and skeletal muscle differentiation.
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Methods |
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C2C12 cells (American Type Culture Collection) were cultured as exponentially growing myoblasts in growth medium (GM), specifically, Dulbecco's modified Eagle's medium (DMEM) containing 20% fetal calf serum (FCS), 4 mML-glutamine and 100 IU ml–1–100 µg ml–1 penicillin–streptomycin. Cells were seeded in Petri dishes at a density of 1500–2000 cells cm–2 in GM, and subcultured by standard trypsinization every three days (Pietrangelo et al. 2002). After three days in GM, differentiation was induced using normal differentiation medium (DM) containing DMEM with 2% horse serum, L-glutamine and antibiotics, as previously described, or synthetic differentiation medium (SM) consisting of DMEM plus 1% (w/v) bovine serum albumin (BSA), L-glutamine and antibiotics, where specified.
Proliferation and cell cycle assays
Cellular proliferation was tested using a colorimetric assay of 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT). We seeded 1000 myoblasts per well in a final volume of 200 µl GM or DM in 96-well plates. Next, myoblasts were incubated with stimuli at different time points. Incubation was terminated by adding 20 µl of MTT solution (5 mg ml–1 in phosphate-buffered saline (PBS)) to each well, followed by incubation at 37°C for 3 h. The supernatant was removed, and 200 µl dimethylsulphoxide (DMSO) was added to each well. The plate was agitated for 5 min, and incubated for 30 min at 37°C. Finally, the plate was read at 540 nm on a Titertek Multiscan Microelisa Reader (Flow Laboratories, Urvine, UT, USA).
Cytofluorimetric determination was performed by incubating samples with a fluorochromic solution containing 250 mg sodium citrate, 5 mg ml–1 RNase, 750 µl NP40 Nonidet, and 16.5 mg propidium iodide in 200 ml ddH2O for 30 min at 37°C. Samples were prepared by standard trypsinization, and subsequently read in a cytofluorimeter (Beckman Epics XL Coulter) connected to a personal computer (PC).
Video-imaging
For [Ca2+]i and plasma membrane potential fluorescence measurements, 2000 cells cm–2 were plated in Petri dishes containing glass coverslips. Myoblasts were incubated for 45 min at 37°C in normal external solution (NES) supplemented with 10 mg ml–1 BSA and 5 µM Fura-2 AM (Molecular Probes, Eugene, OR, USA). The NES solution consisted of 10 mM glucose, 140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2 and 10 Hepes, pH 7.4. After Fura incubation for 35 min, 250 nM potentiometric dye, bis-(1,3-dibutyl-barbituric acid)trimethine oxol (DiBAC4, Molecular Probes), was added for a subsequent 10 min. Cells were rinsed with NES, and maintained for 15 min at 37°C to allow de-esterification hydrolysis of dyes. Coverslips were mounted in a chamber, as previously described (Pietrangelo et al. 2002), and living cells sequentially excited at 340 or 480 nm with a high-speed wavelength switcher Polychrome II (Till Photonics, Germany). Fluorescence images were collected using a 40x oil objective lens, acquired using an intensified CCD camera (Hamamatsu Photonics, Hamamatsu, Japan), stored on a PC, and analysed off-line. The acquisition time for each fluorescence emission was 0.5 s. Temporal plots were calculated as the mean value of the fluorescence signal (f) in a selected representative cellular area after the subtraction of background fluorescence (f0).
Electrophysiological recordings
Whole-cell (perforated) and cell-attached patch clamp configurations were used for electrophysiological recordings on C2C12 myoblasts. Electrode resistance was 3–5 M
for whole-cell, and 5–10 M for cell-attached experiments. Single-channel and whole-cell currents were amplified with a List EPC-7 amplifier (List Medical, Darmstadt, Germany), and digitized with a 12-bit A/D converter (TL-1, DMA interface; Axon Instruments, Inc., Union City, CA, USA). The pCLAMP software package (version 7.0; Axon Instruments) was used. For on-line data collection, macroscopic and single-channel currents were filtered at 5 and 0.5 kHz, and sampled at 20 and 200 µs/point, respectively. Experiments were carried out at room temperature (18–22°C), and the resulting data are normally presented as means ±S.E.M.
Macroscopic currents were recorded using the perforated-patch method. With this configuration, the bathing physiological salt solution (PSS) was (mM): NaCl 106.5, KCl 5, CaCl2 2, MgCl2 2, Mops 5, glucose 20, sodium gluconate 30, at pH 7.25; and the pipette solution was (mM): K2SO4 57.5, KCl 55, MgCl2 5, Mops 10, at pH 7.2. Electrical access to the cytoplasm was achieved by adding amphotericin B (200 µM) to the pipette solution. Access resistances in the range 10–20 M were achieved within 10 min following seal formation. In cell-attached recordings the bathing solution was as for perforated patch configuration experiments, and the pipette solution was (mM): KCl 150, CaCl2 2, MgCl2 2, Mops 5, glucose 10, pH 7.4. ChTX was purchased from Alomone Laboratories (Jerusalem, Israel). We routinely used clotrimazole (CTL) because of its greater selectivity towards IKCa channels, compared to other Ca2+-activated K+ channels. CTL, thapsigargin (TG) and amphotericin B were similarly dissolved in stock solutions using DMSO to concentrations of 20, 1 and 50 mM, respectively. The maximal DMSO concentration in the recording solution was less than 0.1%. GTP in Figs 2, 4, 5 and 6 was applied using Picospritzer II (General Valve Corporation, Fairfield, NJ, USA) via a glass pipette with tip diameter about twice as large as that used for whole-cell recording. The glass pipette was normally placed within 40 µm of the patched cell to accelerate the increase in drug concentration at the cell, and allow brief application times which would minimize desensitization of the GTP response, and rundown of IKCa channels. Since our perfusion systems may impart a shear stress to the membrane, we carried out control experiments in the absence of GTP to exclude possible effects due to mechanical stress (n= 10).
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Samples of differentiating cells (1500–2000 cells cm–2) in SM or DM for 20 and 42 h were washed twice with PBS, and fixed with 95% ethanol for 5 min at room temperature. Cells were treated with a 2% FCS solution for 30 min at 37°C. Next, cells were incubated with monoclonal mouse antimyosin heavy chain antibody (1 : 50 dilution) (MF20, Hybridoma Bank, Cornell University Medical College, NY, USA) in PBS for 90 min at 37°C, followed by three washes with PBS (5 min each at room temperature), and incubation with anti-mouse secondary antibody (1 : 200 dilution) (mouse immunoglobulins biotin, Dako code E0354 Dakocytomation, Glostrup, Denmark) in PBS for 40 min at room temperature. After three washes with PBS (5 min each at room temperature), we amplified the signal by incubating with the AB complex for 45 min at room temperature (StreptABComplex/HRP, Dako, code K0377). Following another two washes with PBS, staining with diaminobenzidine (DAB) and H2O2 was performed, as specified in the kit. Pictures of stained cells were acquired using a digital camera on an inverted microscope (Leica) at 200x magnification, and stored on a host computer. We calculated the percentage (mean ±S.E.M.) of MF20-positive cells from images obtained at different random fields on three distinct coverslips for each treatment. Experiments were repeated twice, both with cells differentiated in SM and DM.
Western blots
Cells were plated at a density of 1500–2000 cells cm–2 in GM for 3 days, and starved with SM for 2 days. Cellular pellets (2 days in SM with or without 500 µM GTP) were lysed in buffer containing 2.5% SDS. Proteins were quantified with the Bradford assay (Bio-Rad). Aliquots for electrophoresis (6 µg of proteins per sample) were separated with 10% acrylamide–bisacrylamide (30 : 0.8) (Amersham), and transferred to a nitrocellulose transfer membrane (Schleicher & Schuell). The membrane was incubated with primary antibody MF20 (1 : 1000 dilution) and secondary antibody goat anti-mouse (1 : 5000 dilution) (Pierce Biotechnology, Inc., Rockford, IL, USA), and developed using enhanced chemiluminescence ECL reagent (Amersham). The same blot was first probed with MF20, then stripped and reprobed with monoclonal antibodies against ß-cytoplasmatic actin (42 kDa), used as control (Sigma). The bands were digitized by a scanner and analysed by densitometry with the Imagemaster 1D software (Pharmacia Biotech).
Reagents for cellular culture were purchased from Gibco (Paisley, Scotland, UK). All chemicals used were of analytical grade. Dimethyl sulfoxide (DMSO), TEA, d-tubocurarine (d-TC), CTL, GTP, MTT, TG, Reactive Blue 2 (RB2) and salts were purchased from Sigma Chemical Co. (St Louis, MO, USA), unless otherwise specified.
Statistical analysis of data was performed using Student's paired t test, and automatically derived with Prism 2.0 software (GraphPad Software, Inc., San Diego, CA, USA).
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Results |
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Taking into account the strong link between [Ca2+]i and electrophysiological properties of the cell membrane in differentiating versus excitable cells, we investigated whether a GTP-induced [Ca2+]i increase modified the membrane polarization of single myoblasts, utilizing a single cell simultaneous fluorescence technique to record variations in both [Ca2+]i and plasma membrane potential.
Video-imaging experiments were performed by incubating cells with two fluorescent dyes: Fura-2, a useful calcium indicator, and DiBAC4, a well known and useful voltage-sensitive fluorescent dye (Brauner et al. 1984; González & Tsien, 1995) capable of monitoring variations in the C2C12 membrane resting potential stimulated by an increase in GTP-induced [Ca2+]i (membrane resting potential estimated as –26 ± 5 mV, see Fioretti et al. 2005).
As observed in Fig. 1, in response to 500 µM GTP, the myoblast displayed a marked increase in the 340 nm-dependent Fura-2 signal, indicative of a significant [Ca2+]i increase. The same cell also displayed a marked decrease in 480 nm-dependent DiBAC4 signal, implying hyperpolarization of the plasma membrane, with a slight delay with respect to the [Ca2+]i increase.
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Using electrophysiological methods we first assessed the level of hyperpolarization induced by GTP. We recorded the membrane potential of C2C12 myoblasts in perforated patch conditions following application of 500 µM GTP, and found that the membrane potential hyperpolarized from a mean value of –15 ± 6 mV to a mean value of –75 ± 11 mV (n= 6), with an average delay of 24 ± 5.8 s (n= 3, Fig. 2A). We then set out to determine how activation of GTP binding sites is translated into changes of the membrane potential, by recording macroscopic currents under voltage-clamp conditions. GTP (500 µM) stimulated transient outward currents (Fig. 2B) with a time course closely matching the voltage response (cf. Fig. 2A). The latency time of the current response was 19.4 ± 5.3 s (n= 8). This suggests a complex transduction pathway for GTP-induced current activation. The transient outward current induced by 500 µM GTP had a mean amplitude of 84.1 ± 21.7 pA at 0 mV of applied potential (n= 6). The GTP-induced current response (observed in 8 out of 9 cells) could be repeated several times on the same myoblast with little or no decrease in the current amplitude (cf. Fig. 4).
Application of 500 µM GTP for longer time periods evoked similar currents, although longer and higher in amplitude (Fig. 3A). To assess the nature of the GTP-evoked current we first estimated its reversal potential. By applying voltage ramps under control conditions and in the presence of GTP we obtained a mean value of –96 ± 11 mV (n= 9; cf. Fig. 2C), very close to the K+ equilibrium potential under our recording conditions (EK=–103 mV, as calculated from the Nernst relationship).
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The GTP-stimulated K+ current is an IKCa current
Along with published findings, data presented in this report highlight a KCa current as the end target of the GTP-receptor activation pathway. To verify this hypothesis, we assessed the pharmacological profile of the GTP-activated current to evaluate whether it matched any of the KCa currents. Application of either TEA (1 mM) or d-tubocurarine (d-TC; 100 µM) to the bath solution did not significantly affect the GTP-activated current (Fig. 4A, B and E). In contrast, application of 2 µM clotrimazole (CTL) or 100 nM charybdotoxin (ChTX) strongly inhibited the GTP-activated current (Fig. 4C, D and E). The pharmacological profile strongly suggests that GTP activates an IKCa current.
We investigated the biophysical properties of the GTP-activated IKCa current at a single channel level. The single channel conductance and rectification are congruent with the properties usually observed for cloned and native IKCa channels, including the IKCa channel in this tissue previously reported by our group (Fioretti et al. 2005). Figure 5 represents a typical single-channel recording from a cell-attached patch, following the application of 500 µM GTP to the bath. In these experiments, the pipette solution contained 140 mM KCl as the main salt, and the voltage applied was –40 mV. Under these conditions, an inward unitary current was activated (Fig. 5A) with a conductance of 42 pS (at negative voltages), and an inward rectifying I–V relationship (Fig. 5B and C). The average single-channel conductance measured on five cells (including the one shown) under similar conditions was 27 ± 9 pS.
Metabotropic cascade and Ca2+ source of GTP-activated IKCa currents
Suramin (250 µM) and RB2 (100 µM) are capable of abolishing the GTP-induced Ca2+ response (Pietrangelo et al. 2002). These findings suggest that IKCa channel activation by GTP is mediated by specific types of purinoceptors (metabotropic) present in this cell line. Additionally, in C2C12 cells, the metabotropic purinoceptor is functionally linked to PLC activation via the G-protein, resulting in IP3 and DAG production and Ca2+ release from intracellular stores (Henning et al. 1993). We confirmed the intracellular location of the Ca2+ source for GTP-induced IKCa current activation. In C2C12 myoblasts responsive to GTP, the application of 1 µM thapsigargin (TG) caused a transient increase in the IKCa current, probably reflecting the [Ca2+]i increase that followed the emptying of intracellular calcium stores. After this manoeuvre, GTP was no longer capable of inducing membrane current responses (n= 3; Fig. 6A), suggesting that Ca2+ release from TG-sensitive stores mediates the GTP-induced K+ current activation. Thereafter, no significant differences were observed in the GTP-induced current response in the presence of low (10 µM) external calcium solution (n= 3; Fig. 6B), suggesting that the GTP-induced response is not dependent on external Ca2+.
GTP modulation of myoblast proliferation
Several purines, including GTP, stimulate the proliferation of a wide range of cell types (Rathbone et al. 1992). Furthermore, the IK channel is a regulator of the proliferative state in different cell lines, including fibroblast, lymphocyte and a myogenic cellular model (Peña & Rane, 1999; Ghanshani et al. 2000).
We tested the GTP effect on growth rate of both cycling and differentiating cells. In fact, the onset of myogenesis commits myoblasts to proliferate, to escape the cell cycle and then to fuse into polynucleated myotubes (Andres & Walsh, 1996). The growth rates of cycling C2C12 myoblasts were tested in proliferation medium (GM) at different times (24–120 h) in the presence of 500 µM GTP (data not shown). Under these conditions, no statistically significant differences compared to the control group (without GTP) were observed for the tested times. A partial and apparently inconsistent effect on the proliferative rate of C2C12 cells in the presence of 50 and 500 µM GTP was observed when GM was replaced with differentiation medium from which serum was removed (DM). Under these conditions, the effect of GTP on proliferation was twofold: GTP stimulated proliferation in the first 24 h, whereas it inhibited proliferation later. In particular, during the first 24 h of incubation 500 µM GTP stimulated the proliferative rate slightly (about 15–20% more cells than control; Fig. 7A and B), whereas starting from the second day, GTP (50 and 500 µM) induced significant inhibition of cell division (Fig. 7A; here we show the effect of 500 µM GTP).
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The initial GTP-dependent proliferative boost was inhibited upon preincubation with 100 nM ChTX (Fig. 7B).
Cell cycle analysis confirmed this proliferation boost, since the transition from G0/G1 to G2/S was significantly increased in the presence of 500 µM GTP for 24 h (Fig. 7C). In particular, at zero time, we observed control cells, starting from 86 ± 5% G0/G1 and 14 ± 8 G2/S, while after 24 h of GTP incubation, approximately 13% of cells escaping G0/G1 phases accumulated in G2/S. In fact, 75 ± 2% of control cells were in G0/G1 and 25 ± 2% in G2/S, while 62.0 ± 0.2% of GTP-treated cells were in G0/G1, and 38 ± 0.7% in G2/S.
We wondered whether the GTP-induced proliferative boost recorded during the first 24 h of C2C12 differentiation was really mediated by GTP specific binding sites on plasma membrane. Considering that in our previous work we demonstrated that the non-specific inhibitor of purinergic receptors RB2 (100 µM) was capable of abolishing the GTP-induced Ca2+ response (Pietrangelo et al. 2002), we preincubated the cells with 100 µM RB2 and then we stimulated them with 500 µM GTP for 24 h. After this period of time, we measured the percentage of living cells. As can be seen in Fig. 8, the presence of RB2 prevented the 500 µM GTP-induced proliferative boost. The difference between cells in differentiation medium (107 ± 7%, bar not present in the graph) and cells differentiating in the presence of RB2 (100 ± 18%, white bar) was not significant.
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Notably, cells differentiating for 3 days in the presence of 500 µM GTP (in DM) were bigger and more hypertrophic than control cells. GTP-treated cells (randomly selected from three different pictures) were 63 ± 9% (n= 13) thicker than control cells (P
0.05). Moreover, GTP-treated myotubes (randomly selected from three different pictures) were 55 ± 8% (n= 13) longer than control myotubes (P 0.001). Figure 9 depicts a typical example. Experiments using either DM or SM were repeated twice, with similar results (data not shown).
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On the other hand, we performed experiments in which we differentiated the cells for 2 days and then we stimulated with 500 µM GTP for the subsequent 24 h. At the end of the third day we stopped the experiment and analysed the data. Under these conditions no significant biological effects related to the presence of GTP could be observed, neither the cell number increase nor the hypertrophic effect (data not shown).
Western blot and immunocytochemistry of myosin heavy chain proteins
During skeletal muscle differentiation, we observed a gradual increase in the expression and accumulation of different myofibrillar proteins essential for sarcomeric assembly. Some of these, in particular the different isoforms of myosin heavy chain (MyHC), are markers of myogenesis (Andres & Walsh, 1996; Berg et al. 2001).
Accordingly, we employed MyHC proteins to evaluate whether GTP positively modulates progression of the myogenic programme. Quantitative analysis of the GTP effect on MyHC was performed by Western blotting of C2C12 cells after 2 days of differentiation in SM in the presence of 500 µM GTP. In Fig. 10, lane 1 represents the MyHC protein quantity in the control, while lane 2 represents the MyHC content of cells incubated with 500 µM GTP over the same period. It is evident that after 2 days of serum starvation, treated cells expressed MyHC proteins more abundantly, compared to the control. The densitometric analysis of ß-cytoplasmic actin bands on C2C12 control (peak = 0.93; average = 0.89; trace as OD x mm = 1.76) and on GTP-treated C2C12 (peak = 1.02; average = 0.93; trace as OD x mm = 1.80) revealed no significant difference on protein loading.
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A comparison of MyHC protein expression in control and treated cells (50 or 500 µM GTP) after 20 and 42 h of incubation revealed significant differences between the experimental groups (see Table 1). In particular, at an incubation time period of 20 h, only mononucleated cells displayed a 50% increase (see Table 1) in MyHC positivity in the presence of GTP (50 and 500 µM).
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Discussion |
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This mechanism is ubiquitous in mammals, albeit with different expression and/or capacities, dependent on the different receptor typologies and mammalian species. For example, human astrocytoma cells expressing P2Y4 receptors are fully activated by UTP, GTP and ITP, while ATP and CTP are unable to increase [Ca2+] (Kennedy et al. 2000). On the other hand, in cells expressing rat P2Y4 receptors, all nucleotides tested increase [Ca2+]i with similar maximal effects (Kennedy et al. 2000). In PC12 cells, one specific GTP-binding site, associated with enhancing NGF-induced differentiation, increased [Ca2+]i through the CICR mechanism (Gysbers et al. 2000; Guarnieri et al. 2004). In the C2C12 cell line, two binding sites highly specific for GTP over ATP and UTP, and with different Kd values, were identified. Binding of GTP to these sites induced a significant [Ca2+]i increase, starting from a baseline of 87 ± 39 nM (Pietrangelo et al. 2002). The main sources of nucleotides into different tissue could be the cells themselves. In particular, the muscle cells could release GTP as a consequence of micro or more extensive lesions (as well as during physical exercise) and several species of nucleotide (mainly ATP) could be released during motoneuron firing (Burnstock, 1976). It is worth mentioning that the intracellular concentration of GTP is estimated ranging over micromolar concentrations (Traut et al. 1994) and, for this reason, cell damage or mechanical stress could be able to release into the extracellular medium up to hundreds of µM of guanosine nucleotides.
This study shows that IKCa channel activity is required for the enhancement of C2C12 differentiation promoted by GTP, with no other Ca2+-activated K+ channels being involved. This conclusion is based on the finding that ChTX was able to inhibit the GTP-induced differentiation, and on our previous observations that C2C12 cells cultured for 1–4 days in DM do not express ChTX-sensitive BK channels (Fioretti et al. 2005; and the authors' unpublished observations).
This result seems to contrast with a previous study on the 10T1/2-MRF4 myogenic cell line where IKCa channels were instead shown to be instrumental to the bFGF- and TGF-ß-dependent suppression of myogenesis (see Introduction; Peña et al. 2000). These apparently conflicting results could be due to the fact that while the 10T1/2-MRF-4 cell line permanently expresses the MFR-4 trascription factor (which controls muscle-specific genes, and is inhibited by IKCa channels), in C2C12 myoblasts this factor appears only later during myogenesis (Dedieu et al. 2002). It is possible that other Ca2+-dependent muscle-specific trascription factors, expressed earlier in the myogenesis of C2C12 cells, are activated by IKCa channels.
Electrical events are thought to be very important during myoblast differentiation, because of their control over [Ca2+]i that in turn regulates several Ca2+-dependent processes. For example, during differentiation of human muscle satellite cells the resting potential is hyperpolarized by as much as 55 mV, due to the sequential activation of ether-à-go-go and inward rectifier K+ channels. Such a hyperpolarization, which leads to an increased Ca2+ influx through T-type Ca2+ channels, is essential for cell differentiation, and fusion of competent myoblasts into multinucleated myotubes (Liu et al. 1998; Bernheim & Bader, 2002).
In this study we have shown that GTP is capable of hyperpolarizing the membrane by increasing [Ca2+]i and in turn activating the IKCa channel. Experiments with drugs capable of emptying internal Ca2+ stores (1 µM thapsigargin), or low extracellular Ca2+ (10 µM) show the pivotal role of the internal Ca2+ stores in the GTP-induced IKCa channel activation. It is thus hypothesized that GTP binding to its membrane sites stimulates PLC activity, which results in an increased cytoplasmic IP3 level that activates the IP3-operated Ca2+ channels, with consequent increase of the [Ca2+]i. This ultimately results in IKCa channel activation and membrane hyperpolarization. This cascade of events may have physiological significance. In fact, similar to events in human myoblast differentiation, the IKCa channel-mediated hyperpolarization may allow an increase in Ca2+ influx. In addition the IKCa channel activation may be necessary to hyperpolarize the membrane potential to levels negative enough to activate the inward-rectifier K channels, shown to be involved in the maintenance of a negative resting potential in differentiating human myoblasts (Bernheim & Bader, 2002), and also shown to be expressed in 40% of differentiating C2C12 myoblasts (Kubo, 1991).
The observation that it is sufficient to stimulate the cells with 500 µM GTP at the beginning of differentiation even if for only 2 h to improve this process and that GTP does not have an effect when applied late in differentiation (after 2 days), suggest that this nucleotide could lead to specific pathways of differentiation.
Moreover, the myoblasts stimulated with 500 µM GTP activated the IKCa current in 97% of the tested cells while only 33% of tested myoblasts were able to respond to 50 µM GTP in the experimental medium. Data previously reported by our group (Pietrangelo et al. 2002) show that 30% of the C2C12 undifferentiated population displays a particular [Ca2+]i kinetic derived from high-affinity GTP binding site activation, defined as ‘slow’. It is additionally possible that this slow [Ca2+]i increase is ineffective in IKCa current activation even if it could be effective in activating some other early signalling effector leading to the activation of a specific pathway implied by MyHC protein expression.
Several data from different laboratories establish that activation of the IKCa current by numerous growth factors plays an important role in stimulating the proliferation of different cell lines, including a particular myogenic cell line (Vandorpe et al. 1998; Ghanshani et al. 2000; Peña et al. 2000). We previously demonstrated the presence of IKCa channels in both C2C12 myoblasts and differentiating cells up to 2 days (about 20–30 pA pF–1). In fact, after 3–4 days of differentiation, the IKCa channel disappeared. Moreover, blocking of these channels with charybdotoxin did not modify the growth rate of myoblasts or differentiating cells (Fioretti et al. 2005). The data reported in this study show that activation of the IK current with 500 µM GTP does not modify progression through the cell cycle for long-term exposure, at least in the proliferating medium. Accordingly, we conclude that at least for the myogenic C2C12 cell line, the modification of IKCa conductance does not affect its replicative capacity.
The embryonic development of skeletal muscle is based on myogenic precursor cells regulated by growth signals derived from different sources in the surroundings. Initiation of the myogenic process requires the overexpression of myogenin, Myf5, MyoD and MFR4, transcriptional activators of the myogenic regulatory factor family (MRF) (Parker et al. 2003). In particular, during differentiation, myogenin-positive C2C12 cells are initially proliferating myoblasts, and then withdraw from the cell cycle, establishing a postmitotic state to become terminally differentiated myocytes expressing muscle-specific proteins, such as MyHC (Andres et al. 1996).
At effective concentrations, extracellular GTP commits the myoblasts present in C2C12 cultures to utilize this pathway, since it not only increases the number of MyHC-positive myocytes, but also accelerates some phases (e.g. the anticipation of proliferative boost and block) of the differentiative process, compared to the differentiation induced by growth factor removal from the medium. These effects disappeared in the presence of 100–200 nM ChTX in medium containing GTP. Specifically, only the binding of GTP to low-affinity binding sites favours the fusion process. In fact, after 42 h of incubation, samples treated with 500 µM GTP displayed a significant increase in multinucleated myotubes expressing MyHC proteins.
In our opinion, the two specific GTP binding sites could activate different signal transduction pathways both leading in a different manner and with a different strength to a common scenario of skeletal muscle differentiation. In fact the low efficiency of the high affinity GTP binding site in the recruitment of IK current could justify the lowest efficiency in the fusion process during skeletal muscle differentiation.
This consideration is further strengthened by our unpublished observation revealing that C2C12 myoblasts differentiated in the presence of 500 µM GTP for 24 h show a significant increase of the basal cytoplasmic intracellular [Ca2+] while 50 µM GTP incubated for the same period did not significantly increase the basal calcium into the differentiating cells.
In conclusion, we have shown for the first time that an extracellular GTP pool could act as a myogenic inducer able to sustain all phases of the differentiative process up to MyHC protein expression. Moreover, the GTP-mediated activation of IKCa channels and resulting hyperpolarization may be considered an early step needed for skeletal muscle differentiation of C2C12 cells.
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References |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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Berg
JS, Powell
BC
&
Cheney
RE (2001). A millennial myosin census. Mol Biol Cell
12, 780–794.
Bernheim
L
&
Bader
R (2002). Human myoblast differentiation: Ca2+ channels are activated by K+ channels. News Physiol Sci
17, 22–26.
Brauner T, Hulser DF & Strasser RJ (1984). Comparative measurements of membrane potentials with microelectrodes and voltage-sensitive dyes. Biochim Biophys Acta 771, 208–216.[Medline]
Burnstock G (1976). Do some nerve cells release more than one transmitter? Neuroscience 1, 239–248.[CrossRef][Medline]
Burnstock G (2003). Purinergic receptors in the nervous system. In Current Topics in Membranes, vol 54, Purinergic Receptors and Signalling, ed. Schwiebert EM, pp. 1–27. Academic Press, San Diego.
Burnstock G (2004). P2Y receptors. Current Topics Med Chem 4, 793–803.[CrossRef]
Castle NA (1999). Recent advances in the biology of small conductance calcium-activated potassium channels. Perspectives Drug Discovery Design 15, 131–154.[CrossRef]
Ciccarelli R, Di Iorio P, Giuliani P, D'Alimonte I, Ballerini P, Caciagli F & Rathbone MP (1999). Rat cultured astrocytes release guanine-based purines in basal conditions and after hypoxia/hypoglycemia. Glia 25, 93–98.[CrossRef][Medline]
Clark KD, Hennessey TM, Nelson DL & Preston RR (1997). Extracellular GTP causes membrane-potential oscillations through the parallel activation of Mg2+ and Na+ currents in Paramecium tetraurelia. J Membr Biol 157, 159–167.[CrossRef][Medline]
Dedieu S, Mazères G, Cottin P & Brustis JJ (2002). Involvement of myogenic regulator factors during fusion in cell line C2C12. Int J Dev Biol 46, 235–241.[Medline]
Fioretti
B, Pietrangelo
T, Catacuzzeno
L
&
Franciolini
F (2005). The intermediate-conductance Ca2+-activated K channel is expressed in C2C12 myoblasts and is downregulated during myogenesis. Am J Physiol Cell Physiol
289, C89–C96.
Ghanshani
S, Wulff
H, Miller
MJ, Rohm
H, Neben
A, Gutman
GA, Cahalan
MD
&
Chandy
G (2000). Up-regulation of the IKCa1 potassium channel during T cell activation. J Biol Chem
275, 37137–37149.
González
JE
&
Tsien
RY (1995). Voltage sensing by fluorescence resonance energy transfer in single cells. Biophys J
69, 1272–1280.
Guarnieri S, Fanò G, Rathbone MP & Mariggiò MA (2004). Cooperation in signal transduction of extracellular guanosine 5'-triphosphate and nerve growth factor in neuronal differentiation of PC12 cells. Neuroscience 128, 697–712.[CrossRef][Medline]
Gysbers JW, Guarnieri S, Mariggiò MA, Pietrangelo T, Fanò G & Rathbone MP (2000). Extracellular guanosine 5'-triphosphate enhances nerve growth factor-induced neurite outgrowth via increases in intracellular calcium. Neuroscience 96, 817–824.[CrossRef][Medline]
Henning RH, Duin M, den Hertog A & Nelemans A (1993). Activation of the phospholipase C pathway by ATP is mediated exclusively through nucleotide type P2-purinoceptors in C2C12 myotubes. Br J Pharmacol 110, 747–752.[Medline]
Kennedy C, Qi A, Herold CL, Harden TK & Nicholas RA (2000). ATP, an agonist at the rat P2Y4 receptor, is an antagonist at the human P2Y4 receptor. Mol Pharmacol 18, 926–931.
Kubo
Y (1991). Comparison of initial stages of muscle differentiation in rat and mouse myoblastic and mouse mesodermal stem cell lines. J Physiol
442, 743–759.
Latorre R, Oberhauser A, Labarca P & Alvarez O (1989). Varieties of calcium-activated potassium channels. Annu Rev Physiol 51, 385–399.[CrossRef][Medline]
Liu
JH, Bijlenga
P, Fischer-Lougheed
J, Occhiodoro
T, Kaelin
A, Bader
CR
&
Bernheim
L (1998). Role of an inward rectifier K+ current and of hyperpolarization in human myoblast fusion. J Physiol
510, 467–476.
Mancinelli L, Fanò G, Ferroni L, Secca T & Dolcini BM (1983). Evidence for an ionotropic positive action of cGMP during excitation-contraction coupling in frog sartorius muscle. Can J Physiol Pharmacol 61, 590–594.[Medline]
Mimikakis JL & Nelson DL (1998). Evidence for two separate purinergic responses in Paramecium tetraurelia: XTP inhibits only the oscillatory responses to GTP. J Membrane Biol 163, 19–23.[CrossRef][Medline]
Neary JT, Rathbone MP, Cattabeni F, Abbracchio MP & Burnstock G (1996). Trophic actions of extracellular nucleotides and nucleosides on glial and neuronal cells. Trends Neurosci 19, 13–18.[CrossRef][Medline]
Parker MH, Seale P & Rudnicki M (2003). Looking back to the embryo: defining transcriptional networks in adult myogenesis. Nat Rev Genet 4, 495–505.
Peña
TL, Chen
SH, Konieczny
SF
&
Rane
SG (2000). Ras/MEK/ERK up-regulation of the fibroblast KCa channel FIK is a common mechanism for basic fibroblast growth factor and transforming growth factor ß suppression of myogenesis. J Biol Chem
275, 13677–13682.
Peña
TL
&
Rane
SG (1997). The small conductance calcium activated potassium channel regulates ion channel expression in C3H10T1/2 cells ectopically expressing the muscle regulatory factor MRF4. J Biol Chem
272, 21909–21916.
Peña TL & Rane SG (1999). The fibroblast intermediate conductance KCa channel, FIK, as a prototype for the cell growth regulatory function of the IK channel family. J Membrane Biol 172, 249–257.[CrossRef][Medline]
Pietrangelo T, Mariggiò MA, Lorenzon P, Fulle S, Protasi F, Rathbone MP, Werstiuk E & Fanò G (2002). Characterization of specific GTP binding sites in C2C12 mouse skeletal muscle cells. J Muscle Res Cell Motil 23, 107–118.[CrossRef][Medline]
Rathbone MP, Middlemiss PJ, Gusbers JW, Andrew C, Herman MAR, Reed JK, Ciccarelli R, Di Iorio P & Caciagli F (1999). Trophic effects of purines in neurons and glial cells. Prog Neurobiol 59, 663–690.[CrossRef][Medline]
Rathbone MP, Middlemiss PJ, Gysbers JW, DeForge S, Costello P & Del Maestro RF (1992). Purine nucleosides and nucleotides stimulate proliferation of a wide range of cell types. In Vitro Cell Dev Biol 28A, 529–536.[CrossRef]
Stocker M (2004). Ca2+-activated K+ channels: molecular determinants and function of the SK family. Nat Rev Neurosci 5, 758–770.[CrossRef][Medline]
Traut TW (1994). Physiological concentrations of purines and pyrimidines. Mol Cell Biochem 140, 1–22.[CrossRef][Medline]
Vandorpe
DH, Shmukler
BE, Jiang
L, Lim
B, Maylie
J, Adelman
JP, de Franceschi
L, Cappellini
MD, Brugnara
C
&
Alper
SL (1998). cDNA cloning and functional characterization of the mouse Ca2+-gated K+ channel, mIK1. Roles in regulatory volume decrease and erythroid differentiation. J Biol Chem
273, 21542–21553.
Vergara C, Latorre R, Marrion NV & Adelman JP (1998). Calcium-activated potassium channels. Curr Opin Neurobiol 8, 321–329.[CrossRef][Medline]
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