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Am J Physiol Cell Physiol 294: C66-C73, 2008. First published November 14, 2007; doi:10.1152/ajpcell.00248.2007
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MUSCLE CELL BIOLOGY AND CELL MOTILITY

Neural agrin controls maturation of the excitation-contraction coupling mechanism in human myotubes developing in vitro

Elena Bandi,1,4 Marko Jevsek,3 Tomaz Mars,3 Mihaela Jurdana,1 Elena Formaggio,2 Marina Sciancalepore,1 Guido Fumagalli,2 Zoran Grubic,3 Fabio Ruzzier,1 and Paola Lorenzon1

1Department of Physiology and Pathology and Centre for Neuroscience B.R.A.I.N., University of Trieste, Trieste; 2Department of Medicine and Public Health, University of Verona, Hospital Borgo Roma, Verona, Italy; 3Institute of Pathophysiology, School of Medicine, University of Ljubljana, Ljubljana, Slovenia; and 4Interdepartmental Centre of Molecular Medicine, University of Trieste, Trieste, Italy

Submitted 12 June 2007 ; accepted in final form 12 November 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The aim of this study was to elucidate the mechanisms responsible for the effects of innervation on the maturation of excitation-contraction coupling apparatus in human skeletal muscle. For this purpose, we compared the establishment of the excitation-contraction coupling mechanism in myotubes differentiated in four different experimental paradigms: 1) aneurally cultured, 2) cocultured with fetal rat spinal cord explants, 3) aneurally cultured in medium conditioned by cocultures, and 4) aneurally cultured in medium supplemented with purified recombinant chick neural agrin. Ca2+ imaging indicated that coculturing human muscle cells with rat spinal cord explants increased the fraction of cells showing a functional excitation-contraction coupling mechanism. The effect of spinal cord explants was mimicked by treatment with medium conditioned by cocultures or by addition of 1 nM of recombinant neural agrin to the medium. The treatment with neural agrin increased the number of human muscle cells in which functional ryanodine receptors (RyRs) and dihydropyridine-sensitive L-type Ca2+ channels were detectable. Our data are consistent with the hypothesis that agrin, released from neurons, controls the maturation of the excitation-contraction coupling mechanism and that this effect is due to modulation of both RyRs and L-type Ca2+ channels. Thus, a novel role for neural agrin in skeletal muscle maturation is proposed.

neurotrophic factor; calcium homeostasis; differentiation; skeletal muscle

DEVELOPING NEURITES OF MOTOR nerves control the differentiation of skeletal muscle cells by providing electrical activity and soluble factors (1, 2). Among nerve-derived chemical messengers, a pivotal role is played by the proteoglycan agrin (3). When released from the motor neuron, agrin binds to the basal lamina of the synaptic cleft and activates the muscle-specific tyrosine kinase MuSK (4; reviewed by Ref. 2). The agrin-mediated signaling controls the growth and stabilization of acetylcholine receptor (AChR) clusters (2) and the distribution and expression of other components of the postsynaptic apparatus (5), including receptors for neurotransmitters and trophic factors (6), cytoskeletal proteins (7), and enzymes (8, 9).

The effects of neural agrin are not spatially limited to the postsynaptic apparatus. For example, agrin causes remodeling of the entire muscle cell surface and induces extensive microprocess formation upon nerve arrival (10). Reorganization of cytoskeletal elements along the muscle fiber has also been shown to occur in denervated rat muscles exposed to exogenous agrin (11). These observations raise a new scenario on the mechanisms whereby the nerve controls the properties of the extrajunctional portions of the muscle fiber, which, up to now, have been considered to be strictly dependent on the nerve-induced muscle activity (11, 12, 13).

In human skeletal muscles, innervation modulates the maturation of the skeletal excitation-contraction (E-C) coupling mechanism (14). The new extrajunctional role recognized for agrin raises the question of whether the trophic factor plays a role in the nerve-induced maturation of the E-C coupling mechanism. The goal of the present work was to investigate this aspect of the human skeletal muscle cell physiology. Using videoimaging and electrophysiological techniques, we demonstrate that neural agrin modulates the maturation of the E-C coupling mechanism by increasing the number of cells in which RyRs and dihydropyridine-sensitive L-type voltage-dependent Ca2+ channels were present.

Some of these results have previously been published in abstract form (15, 16).


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Human muscle cell culture. Experiments reported here were approved by the Ethical Commission at the Ministry of Health of the Republic of Slovenia (permit number 63/01/99) in accordance with the Declaration of Helsinki. Cultures of human primary myotubes were prepared as described by Mars et al. (17). Briefly, myoblasts were derived from satellite cells isolated from pieces of hallucis longus muscle discarded during orthopaedic operations of equinovarus condition on 5-, 10-, and 12-year-old healthy donors. Myoblasts were grown in MEM (GIBCO, Grand Island, NY) supplemented with 15% FBS (GIBCO) and maintained at 37°C in saturated humidity and in CO2 (5%)-enriched air. To induce the differentiation into myotubes, myoblasts were plated onto glass coverslips (24 mm) coated with a 1:2 human plasma-gelatin mixture (1.5%; Sigma-Aldrich, St. Louis, MO) and grown in a differentiation medium consisting of F14 medium (GIBCO) supplemented with 10% FBS, 50 ng/ml FGF, 10 ng/ml EGF, and 10 µg/ml insulin (all obtained from Sigma-Aldrich). For the preparation of cocultures, timed pregnant Wistar rats were killed by CO2 exposure at 14 days of gestation, as specified by the Veterinary Administration of the Ministry of Agriculture, Forestry and Food of Slovenia (permit number 323-349/2003-3). All efforts were made to minimize the number of animals used and their suffering. Spinal cords were aseptically dissected from embryonic day 14 embryos and cut into 1-mm-thick cross-sections preserving the anatomical connections with dorsal root ganglia and the integrity of meninges. Up to five explants were placed on the monolayer of myotubes cultivated for 3 days in the differentiation medium. Cocultures were grown further in F14 medium supplemented with 5% FBS and 10 µg/ml insulin.

The experiments were performed on human myotubes differentiated in four different conditions: 1) aneurally cultured (control), 2) cocultured with fetal rat spinal cord explants, 3) aneurally cultured in medium conditioned from 48 h of coculture, and 4) aneurally cultured in the presence of 1 nM of recombinant chick neural agrin. For comparison, the results were obtained in different culture conditions: 1) the cell treatment with the medium alone, 2) the addition of spinal cord explants to the cell muscle cultures, 3) the treatment with conditioned medium, and 4) the treatment with neural agrin started at the same time. Myotubes were maintained in culture up to 14 days by renewing the medium every 3 days. In the cell cultures treated with neural agrin, the trophic factor was added every time the medium was renewed.

Human skeletal muscle cells, unlike avian and rodent counterparts, never contract in vitro unless innervated. Thus, in human skeletal muscle cells, contraction is an indication of innervation (17, 18, 19, 20, 21, 22). In human cocultured myotubes, the quantitative analysis of functional innervation was performed by evaluating the number of contracting units (i.e., group of myotubes contracting simultaneously; Ref. 17). The area of innervation was identified as the area in which outgrowing neurons and contracting units were present. Within the innervation area, functional neuromuscular transmission was ascertained by testing the sensitivity of contracting units to {alpha}-bungarotoxin (100 nM) or D-tubocurarine (1 mM).

Mouse muscle cell culture. Mouse myoblasts were established from satellite cells, and they were differentiated in vitro as described in more detail elsewhere (23). Animals were killed by cervical dislocation as approved by the local Animal Care Committee and in agreement with European legislation. Myoblasts were maintained in a medium consisting of Ham's F-10 (ICN Biomedicals, Costa Mesa, CA) containing 20% FCS (PAA Laboratories, Linz, Austria), 2 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin (all Sigma-Aldrich). To obtain cell fusion and differentiation, 1 day after plating, the medium was replaced with DMEM (ICN Biomedicals) supplemented with 2% horse serum (ICN Biomedicals), L-glutamine, penicillin, and streptomycin in the same concentrations as above. The cells were cultivated at 37°C in CO2 (5%)-enriched air. The cells were cultivated up to 14 days in differentiation medium, and the medium was renewed every 3 days. The experiments were performed on myotubes differentiated in the absence (control) and in the presence of neural agrin. In mouse muscle cells treated with neural agrin, the trophic factor was added every time the differentiation medium was renewed.

Western blots of conditioned media. Media conditioned from cocultures and aneurally cultured myotubes were collected every 2–4 days and analyzed by Western blotting for agrin. Part of the conditioned medium from cocultures was incubated with prewashed heparin-Sepharose beads (Amersham Pharmacia Biotech) at a dilution of 10:3 to remove agrin (24, 25, 26). After incubation for 1 h at 4°C with gentle mixing, the suspension was centrifuged at 3,000 rpm for 5 min and the surnatant was collected, filtered, and used for cell culturing.

The conditioned media were probed for the presence of agrin by Western blotting with the anti-murine agrin monoclonal antibody Agr33 (StressGen Biotechnologies, Victoria, BC, Canada), developed and characterized by Hoch et al. (27). Conditioned media were electrophoresed on 10% Bis-Tris gel using XCell SureLock Mini-Cell electrophoresis system (Invitrogen, Carlsbad, CA). Proteins were transferred onto polyvinylidene difluoride membranes (Immobilon, Millipore, MA). Blots were incubated with Agr33 diluted 1:500 in blocking solution consisting of PBS supplemented with 0.1% Tween 20 (Sigma-Aldrich) and 5% I-block (Tropix Applied Biosystems, Bedford, MA). After being washed with blocking solution (3 x 10 min), blots were incubated with alkaline phosphatase-conjugated secondary antibody (1:5,000; Sigma-Aldrich) and developed by nitro blue tetrazolium/5-bromo-4-chloro-3-indoylphosphate-p-toluidine salt (NBT-BCIP; Roche Applied Science, Mannheim, Germany) in a developing buffer solution (0.1 M Tris-HCl, 0.1 M NaCl, and 0.05 MMgCl2).

Purification of recombinant neural agrin. Recombinant full-length chick neural agrin (chick agrin cAgrin7A4B8; see Ref. 28) was purified from the conditioned media of transfected HEK 293 (gift to G. Fumagalli from M. A. Ruegg, University of Basel, Basel, Switzerland) using mono Q-Sepharose fast flow beads (Amersham Pharmacia Biotech, Piscataway, NJ) as described by Bezakova et al. (11). High molecular mass material ({approx}400 kDa) was eluted with 2 M NaCl, analyzed by SDS-PAGE, visualized by silver staining, and Western blotted with anti-agrin antibody (rabbit polyclonal provided by M. A. Ruegg). In the collected fraction, >95% of the material was agrin. The same procedure was applied to the conditioned medium of mock-transfected HEK 293 cells; no material was detected in the surnatants.

Quantitation of AChR aggregation. The formation of AChR clusters was detected by Texas red conjugated {alpha}-bungarotoxin staining (Molecular Probes, Eugene, OR). Cells were fixed in 3.7% paraformaldehyde (10 min), washed in PBS supplemented with 0.1% BSA, and incubated for 1 h at room temperature with Texas red {alpha}-bungarotoxin (1:1,000). After being washed twice for 5 min in Dulbeccos PBS supplemented with 0.1% BSA and once for 10 min in Dulbeccos PBS, coverslips were mounted on glass slides with PBS/glycerol supplemented with 1,4 diazabiciclo[2.2.2]octane (Sigma-Aldrich) as an antifading agent. Slides were observed using an LSM 510 confocal microscope (Zeiss, Oberkochen, Germany) with a laser excitation wavelength at 546 nm (red fluorescence) and a long-pass 560-nm emission filter set. The quantitation of AChR aggregates was performed by counting the number of clusters in randomly chosen microscope fields (x200; 460 µm per 460 µm). For each experiment, at least six fields were counted from not less than three different coverslips. Signals were examined for the length of their longitudinal axes and were considered as AChR clusters only if the corresponding signals were at least 2 µm in length.

Videoimaging. Videoimaging experiments were carried out in a normal external solution (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 5 mM MgCl2, 10 mM glucose, and 10 mM HEPES-NaOH, pH 7.4) or in a Ca2+-free solution (140 mM NaCl, 2.8 mM KCl, 2 mM EGTA, 5 mM MgCl2, 10 mM glucose, and 10 mM HEPES-NaOH, pH 7.4). Fura-2 pentoacetoxymethyl ester (fura-2 AM) was used as fluorescent Ca2+ dye. Cell loading and intracellular Ca2+ measurements were performed as previously described in more detail (29, 30). Image acquisition was done at the rate of four frames per second; the calculation of 340/380 ratio (pixel by pixel) and its temporal plot were performed off-line. For each cell, the fluorescence ratio at rest was assumed to be 1 and only variations corresponding to a peak ≥1.5 were considered as a cell response {i.e., a significant increase of intracellular Ca2+ concentration ([Ca2+]i)}. To estimate the percentage of responsive cells, at least 5 optical fields were observed; the minimum number of cells analyzed was 27 for each condition. During the experiments, the fura-2-loaded cells were maintained at a constant temperature of 37°C.

Electrophysiology. The electrophysiological experiments were performed using the patch-clamp technique in the whole cell configuration. Data were acquired at room temperature using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA) and digitized with a Digidata 1321A interface. Currents were acquired at a sampling time of 200 µs and low-pass filtered at 2 kHz. Leakage and capacity currents were digitally subtracted. The pipettes were made from borosilicate glass (Harvard Apparatus, Edenbridge, UK) and heat-polished to resistances of 4–6 M{Omega}. Pipette solutions contained the following: 130 mM CsCl, 0.005 mM CaCl2, 1 mM MgCl2, 5.6 mM glucose, 10 mM HEPES-NaOH, 1 mM EGTA, and 2 mM ATP, pH 7.3 adjusted using tetraethylammonium OH. The composition of the bath solution was 135 mM tetraethylammonium Cl, 2.5 mM CaCl2, 0.8 mM MgCl2, 5.6 mM glucose, and 10 HEPES-NaOH, pH 7.4 (all the above from Sigma-Aldrich). Membrane capacitance was assessed by integrating the area of membrane responses to a 5-mV hyperpolarizing command from a holding potential of –60 mV. For data acquisition and analysis, the pCLAMP softwares suite (v.8.0, Axon Instruments) and Origin 7 (Microcal Software, Northampton, MA) software were used.

Statistical analysis. The statistical analysis was performed using Prism 4.0 software (GraphPad Software, San Diego, CA). Data were analyzed by Student's unpaired t-test as appropriate or by the repeated measures ANOVA method (for data in Fig. 8C). All values are means ± SE, and differences were considered significant at P < 0.05.


Figure 8
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  		Fig. 8. Recombinant neural agrin increases the number of human myotubes responsive to caffeine (Caff). A: representative temporal plot of the [Ca2+]i transient elicited by 40 mM caffeine in a responsive human myotube. B: when myotubes were differentiated in the presence of neural agrin, the percentage of caffeine-responsive cells increased significantly only in the human cells. C: time course of caffeine-induced [Ca2+]i transients measured in agrin-treated and untreated human cells. Traces are averages of temporal plots recorded in different cells (control: n = 29 cells; agrin: n = 48). Points are means ± SE. The analysis of the curves does not reveal any statistical difference (F-test of between-subjects effects, {alpha} > 0.8). *P < 0.05 vs. control.

 

   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
In skeletal muscle, membrane depolarization leads to contraction due to a mechanical interaction between type 1 RyRs (RyR1s) and dihydropyridine-sensitive L-type voltage-dependent Ca2+ channels. In more detail, the depolarization-induced conformational change of L-type Ca2+ channels causes the opening of RyR1s and the release of Ca2+ from the sarcoplasmic reticulum, which occurs independently from the extracellular Ca2+ and Ca2+ influx (31, 32). In accordance with previous studies (see Ref. 31), in the following sections, we use the term E-C coupling mechanism to indicate the mechanical link between RyR1s and L-type Ca2+ channels.

The effects of coculturing on the establishment of the skeletal type E-C coupling mechanism. With the use of the videoimaging technique, the presence of the functional skeletal E-C coupling mechanism was evaluated by measuring, in each microscopic field, the fraction of myotubes in which a depolarizing solution (60 mM K+ concentration) caused an increase of the [Ca2+]i in the absence of extracellular Ca2+ (Ca2+-free solution supplemented with 2 mM EGTA; Fig. 1A).


Figure 1
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  		Fig. 1. Coculturing improves the maturation of the excitation-contraction (E-C) coupling mechanism. The presence of the skeletal type E-C coupling mechanism was assessed by depolarization with high K+ concentration in a Ca2+-free solution. A: trace represents the temporal plot of the intracellular Ca2+ concentration ([Ca2+]i) variation induced by depolarization in one of the responsive human myotubes. B: in the presence of spinal cord explants, the number of cells exhibiting the E-C coupling mechanism was higher than in aneurally differentiated human myotubes (control). Similar percentages of responsive cells were detected within and outside the innervation zone. *P < 0.05 vs. control.

 
Experiments were carried out on human myotubes differentiated for 12 days in two different culture conditions: 1) aneurally cultured (control), and 2) cocultured with fetal rat spinal cord explants (cocultures). As expected, only in cocultures, we observed groups of myotubes contracting rhythmically and synchronously (see MATERIALS AND METHODS for further details). On day 12 of coculturing, 3.6 ± 0.6 contracting units per explant were detected (number of explants = 41). Contractions were inhibited by {alpha}-bungarotoxin (100 nM) or D-tubocurarine (1 mM), confirming that they were dependent on innervation (n = 95). No contracting units were observed at distances >5 mm from the rim of the explants where the administration of {alpha}-bungarotoxin (100 nM) or D-tubocurarine (1 mM) in itself did not elicit any effect. Therefore, as previously reported (17, 18, 33), the growing neurites reached and innervated only myotubes located within 5 mm from the explants also in our cell cultures.

In control myotubes, the mean percentage of myotubes showing functional skeletal E-C coupling mechanism was 33.04 ± 7.13% (n = 20 fields; Fig. 1B). In the cocultures, the percentage was significantly higher in the innervation zone (62.19 ± 13.91, n = 5 fields). Surprisingly, this value was similarly high in the noninnervation zone as well (65.58 ± 10.17, n = 5 fields; Fig. 1B). These results confirmed that the nerve favored the maturation of the E-C coupling mechanism (14) and suggested that this effect was dependent on soluble factors released by the nerve endings.

The properties of the medium conditioned by cocultures. To confirm the role of the soluble factors on the maturation of the E-C coupling mechanism, we differentiated aneural myotubes in medium conditioned for 48 h from cocultures. After 12 days of differentiation, the results indicated that the conditioned medium induced effects that were comparable with those observed in coculturing (69.65 ± 12.28, n = 8 fields vs. 63.59 ± 7.44, n = 10 fields; Fig. 2).


Figure 2
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  		Fig. 2. Medium conditioned by cocultures improves the maturation of the E-C coupling mechanism. Treatment with conditioned medium increased the percentage of cells with the skeletal E-C coupling mechanism to values comparable with those observed in cocultures. The number of responsive cells was significantly lower when they were cultivated in conditioned medium treated with heparin-Sepharose beads. *P < 0.05 vs. both conditioned and cocultured medium.

 
To determine whether agrin was one of the diffusible factors involved in this effect, we first checked whether the conditioned medium induced AChR aggregates, the well-known biological activity of neural agrin (34). After 24 h incubation, we evaluated the presence of AChR aggregates by fluorescent {alpha}-bungarotoxin staining. The number of aggregates was significantly higher in the myotubes exposed to the conditioned medium than in the control cells (mean = 5.77 ± 0.57 vs. 2.88 ± 0.39, n = 22 and 26 fields, respectively; Fig. 3).


Figure 3
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  		Fig. 3. Medium conditioned by cocultures induces the aggregation of the acetylcholine receptors (AChRs). The AChR aggregation was evaluated by Texas red conjugated {alpha}-bungarotoxin staining. A: representative confocal images of the distribution of AChRs in human myotubes treated with standard medium or with medium conditioned by cocultures. Scale bar = 20 µm. B: the mean number of AChR clusters per optical field was significantly higher in the cells treated with the conditioned medium. *P < 0.05 vs. control.

 
Western blot analysis using murine-specific anti-agrin monoclonal antibodies (33) revealed two bands of approximate molecular masses of 200 and 110 kDa, confirming that the conditioned medium contained agrin (35; Fig. 4).


Figure 4
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  		Fig. 4. Western blot analysis of medium conditioned from cocultures. In the conditioned medium from cocultures, the monoclonal antibody recognized two bands of ~200 and ~110 kDa (lane 1). The 200-kDa protein band is reflective of the core agrin protein size predicted by the cDNA nucleic acid sequence. Low molecular mass species are probably proteolytic fragments of the agrin core protein (Refs. 28, 34, 35). Agrin was not detected in the medium collected by cocultures preincubated with heparin-Sepharose beads (lane 2) and in the culture medium conditioned by aneurally cultivated human myotubes (lane 3). Molecular masses of standard proteins are indicated in kDa.

 
When the conditioned medium was preincubated with heparin-Sepharose beads, a procedure used to remove agrin from solution (24, 25, 26), the monoclonal antibody did not stain any agrin-immunoreactive bands (Fig. 4). The treatment with heparin-Sepharose abolished the effects of the conditioned medium both on AChR aggregates (data not shown; Refs. 24, 25, and 26) and on the E-C coupling mechanism (see Fig. 2).

The effects of recombinant neural agrin on the establishment of the skeletal type E-C coupling mechanism. To further confirm the control of neural agrin on the maturation of the E-C coupling mechanism, we planned to differentiate human myotubes aneurally (control) and in the presence of recombinant chick neural agrin up to 12 days. We first checked whether the purified neural agrin retained its biological activity by measuring AChR-aggregating activity (34). To this aim, both human and in mouse myotubes were exposed to 1 nM purified agrin and the number of AChR clusters per optic field was determined by fluorescence microscopy 24 h later. The number of AChR clusters was significantly higher compared with control (human cells: 11.00 ± 0.71 vs. 2.00 ± 0.38, n = 15 and 19 fields, respectively; mouse cells: 17.33 ± 1.84 vs. 2.92 ± 0.42, n = 15 and 12 fields, respectively; Fig. 5), showing that the purified chick neural agrin was endowed with biological activity on both human and mouse myotubes.


Figure 5
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  		Fig. 5. Recombinant chick neural agrin induces AChR aggregation in human and mouse myotubes. In both human and mouse myotubes, the mean number of AChR clusters per optical field significantly increased after the incubation with chick neural agrin (1 nM). The supernatant of mock-transfected HEK 293 cells had no AChR-aggregating activity. *P < 0.05 vs. control.

 
The analysis of the maturation of the E-C coupling in control human myotubes revealed that the mean percentage of cells with E-C coupling increased significantly from day 3 (12.50 ± 8.40%; n = 8 fields) to day 6 (23.13 ± 7.02%; n = 7) of differentiation and then remained stable up to day 12 when the mean percentage of responsive cells was 28.58 ± 10.09% (n = 9; Fig. 6). In human myotubes cultured in the presence of 1 nM of recombinant neural agrin, the mean percentage of cells with E-C coupling was significantly higher at each differentiation time: 38.33 ± 1.66% (n = 6 fields), 61.67 ± 7.92% (n = 5), and 70.44 ± 5.52% (n = 7) on days 3, 6, and 12, respectively (Fig. 6). When similar experiments were performed on murine myotubes, the occurrence of E-C coupling was similar in aneurally cultured and agrin-treated mouse myotubes (Fig. 6).


Figure 6
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  		Fig. 6. Effects of recombinant neural agrin on the maturation of the E-C coupling mechanism. In aneurally cultivated human cells differentiated in the presence of recombinant neural agrin (1 nM), the number of cells exhibiting the E-C coupling mechanism was higher than in controls at all time points considered. Nevertheless, the establishment of the E-C coupling mechanism reached a plateau level at the same time in both conditions. Neural agrin had no effect on the E-C coupling in mouse myotubes. *P < 0.05 vs. control.

 
Interestingly, after 12 days of treatment with recombinant neural agrin, the percentage of human cells characterized by a skeletal E-C coupling mechanism was comparable with that of cells differentiated for 12 days in the presence of spinal cord explants (70.44 ± 5.52 vs. 63.59 ± 7.44%, n = 7 and 10 fields, respectively; Fig. 7). The addition of recombinant neural agrin to cocultured cells did not enhance the effect of coculturing per se (60.00 ± 9.25%, n = 5 fields; Fig. 7). On the other hand, when added to medium conditioned by cocultures and preincubated with heparin-Sepharose beads (i.e., depleted of endogenous agrin), recombinant neural agrin restored the capability of the medium to improve the maturation of the E-C coupling (54.29 ± 2.97%, n = 6 fields; Fig. 7).


Figure 7
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  		Fig. 7. Recombinant neural agrin mimics the effect of coculturing and restores the physiological activity of heparin-Sepharose-treated conditioned medium. The proportion of cells with the E-C coupling mechanism was comparable in agrin-treated and cocultured myotubes. Recombinant neural agrin did not improve the number of cells exhibiting the E-C coupling mechanism when added to cocultures, but it did revert the effect of the heparin-Sepharose treatment of conditioned medium (cf. Fig. 4).

 
In cells maintained in supernatant obtained from mock-transfected HEK 293 cells, the mean number of cells showing an E-C coupling mechanism (24.57 ± 7.80%, n = 6 fields) was comparable with control (aneurally cultured cells) excluding any aspecific effect of the supernatant (Fig. 7).

Effects of recombinant neural agrin on the number of caffeine-responsive cells. To determine whether the nerve explant and the agrin-induced maturation of the E-C coupling mechanism involved modulation of RyRs, we measured the changes in [Ca2+]i induced by 40 mM caffeine, an agonist of RyRs, using videoimaging (Fig. 8A). The caffeine responsiveness was evaluated from day 6 of differentiation, i.e., when the maturation of the E-C coupling mechanism reached the plateau both in aneurally cultured (control) and in agrin-treated myotubes (cf. Fig. 6).

In control human myotubes, even after 12 days of differentiation, the mean percentage of cells showing an increase in [Ca2+]i after caffeine application was 31.33 ± 5.56% (n = 20 fields). In cells treated for 12 days with 1 nM agrin the percentage of responsive cells increased significantly to 70.00 ± 6.88% (n = 17 fields). No changes, compared with control, were observed in mouse cells (Fig. 8B).

Interestingly, in human myotubes, the treatment with neural agrin did not affect the mean amplitude or the kinetics of the [Ca2+]i transients elicited by caffeine (Fig. 8C).

Effects of recombinant neural agrin on the number of cells with voltage-dependent L-type Ca2+ channels. We used an electrophysiological approach to determine if agrin affected the activity of L-type voltage-dependent Ca2+ channels. To this aim, patch-clamp experiments were performed in the whole cell configuration. Due to the size of differentiating myotubes, the patch-clamp experiments were carried out on myotubes differentiated for 7–10 days, when the maturation of the E-C coupling apparatus was completed (cf. Fig. 6) and the myotubes were still small enough to avoid space clamp problems. The Ca2+ currents were recorded under voltage-clamp mode with depolarizing voltage steps to +10 mV (500-ms long) starting from a holding potential of –30 mV (700-ms long). Under such experimental conditions, the low voltage activated T-type Ca2+ currents, present during development in skeletal muscle cells (36, 37, 38), were inactivated.

In aneurally differentiated human cultures (control), L-type Ca2+ currents (ICa,L) were detected in 5 out of 11 cells (~45%) and the mean ICa,L peak density was 0.25 ± 0.08 pA/pF (n = 10). Representative currents are shown in Fig. 9A. When the same currents were recorded in human muscle myotubes differentiated in the presence of 1 nM of recombinant neural agrin, the percentage of cells with ICa,L increased to 90% (10 out of 11; Fig. 9B) and the ICa,L peak density increased to 1.05 ± 0.11 pA/pF (n = 10; Fig. 9C).


Figure 9
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  		Fig. 9. Recombinant neural agrin controls L-type voltage-dependent Ca2+ currents in human myotubes. A: representative L-type Ca2+ currents (ICa,L) elicited by a voltage step from –30 mV to + 10 mV (500 ms) in control (black trace) and in agrin-treated (grey trace) human and mouse myotubes. B: percentage of human and mouse cells with ICa,L in control conditions and after agrin incubation. C: ICa,L current density measured as the ratio between current amplitude and membrane input capacitance in the same cells as in B. *P < 0.05 vs. control.

 
In aneurally differentiated mouse myotubes (control), the ICa,L was present in all the cells tested (n = 24) and the peak density was higher than in the human counterpart (Fig. 9). In the presence of agrin, both the mean number of mouse myotubes presenting ICa,L and the mean ICa,L peak density did not change significantly (from 3.24 ± 0.25 to 2.54 ± 0.26 pA/pF, n = 13 and 8, respectively; Fig. 9B and C).


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Agrin is the master controller of neuromuscular junction formation, growth, and maintenance. Here we propose that neural agrin also acts as a trophic factor on the maturation of a nonsynaptic property of human skeletal muscle cells, the E-C coupling apparatus. The effects appeared to be dependent on the modulation of RyRs and voltage-dependent Ca2+ channels. To our knowledge, this is the first evidence of the involvement of neural agrin in the maturation of the skeletal muscle E-C coupling.

Several muscle properties, including the mechanism governing [Ca2+]i, are nerve dependent. In some species, including mouse, the maturation of the E-C coupling mechanism proceeds even in the absence of the nerve (29, 39, 40). In contrast, in human muscle cells, the nerve plays a critical role in the subcellular distribution and expression of L-type Ca2+ channels and RyRs, in the formation of striated junctions, and in the establishment of the E-C coupling mechanism (14).

The nerve controls the development of muscle cells via the release of trophic factors and/or the induction of muscle activity. The long-term nerve-driven electrical activity and the related Ca2+ signaling contribute to the cross-striated organization of the contractile apparatus in human muscle cells (14) as well as to the expression of several other biological properties of the muscle fibers (41). The persistent electrical activity also appears to be relevant for the maintenance of the E-C coupling and contractile machinery in vivo (13).

Differently from nerve-induced muscle activity, nerve-released diffusible factors are considered relevant for the control of local properties of the muscle fibers, specifically for the establishment, maturation, and control of the neuromuscular junction. However, recent evidence indicates a role for soluble factors, and for neural agrin in particular, also in the control of some of the properties of the extrajunctional region of the muscle fiber (10, 11, 14). With the present study, we investigated the involvement of neural agrin in the maturation of the E-C coupling mechanism.

In agreement with Tanaka et al. (14), we first observed that coculturing human myotubes with rat fetal spinal cord explants increased the number of muscle cells exhibiting a mature E-C coupling mechanism. Then, by analyzing systematically human cells, we noted that the maturation of the E-C coupling mechanism had spread to the entire culture and was not limited to the area of innervation. This observation strongly suggests that the enhancement in the maturation of the E-C coupling mechanism was induced by soluble factors released by the explants.

Western blot analysis and the ability of the conditioned medium to aggregate the AChRs indicated that neural agrin was released by the explants and that its concentration was high enough to trigger and control physiological phenomena in the surrounding muscle cells. Agrin binds to heparin (24, 25, 26), and we used heparin-Sepharose beads to deplete the conditioned medium of this factor. After the treatment, the conditioned medium lacked agrin immunoreactivity and the ability to cluster AChRs as expected. More interestingly, the conditioned medium preincubated with heparin-Sepharose also lacked the ability to modulate the E-C coupling, suggesting a critical role for neural agrin in favoring the establishment of such mechanism.

Our data do not exclude that other trophic factors could be released by the nerve endings. On the other hand, full functional activity was restored when recombinant neural agrin was added to the heparin-Sepharose-treated conditioned medium. These data indicate that, among the possible trophic molecules, neural agrin plays a primary role. This conclusion was further confirmed by the absence of any additive effect of recombinant neural agrin when it was added to cocultures.

In mouse muscle cells, 1 nM of recombinant neural agrin aggregated AChRs but did not affect the E-C coupling mechanism even if used at saturating levels (34). This suggests that, in contrast to humans, this aspect of muscle cell development is not controlled by agrin and/or by the nerve in mice.

We then analyzed in more detail the effects of neural agrin on the establishment of the E-C coupling mechanism in human myotubes.

We first studied the effect of recombinant neural agrin on the rate of occurrence of the E-C coupling mechanism and showed that, in the presence of neural agrin, the number of human cells exhibiting the E-C coupling was higher at all the time points considered. We also demonstrated that the treatment with neural agrin did not affect the amplitude and the kinetics of the [Ca2+]i increases elicited by caffeine, thereby excluding any effect of the trophic factor on the internal releasable Ca2+ reservoirs.

We also analyzed if neural agrin controlled the occurrence of the ion channels subserving the E-C coupling apparatus.

The videoimaging experiments showed that neural agrin increased the number of human myotubes responsive to caffeine. During development, both RyR3 and RyR1 isoforms are expressed; our experiments do not provide information on the specific effect of neural agrin on any of the two isoforms. On the other hand, the expression level of RyR3 decreases during development (42, 43, 44) and the RyR1 is the only isoform known to mechanically couple to dihydropyridine-sensitive L-type Ca2+ channels (31, 32). We therefore hypothesize that neural agrin acts by increasing the number of cells with functional and coupled RyR1s.

The patch-clamp experiments revealed an increased number of cells showing ICa,L current as well as an increase in the ICa,L current density after agrin treatment. This suggests that neural agrin favors the establishment of the E-C coupling mechanism also by upregulating the number of L-type voltage-dependent Ca2+ channels on the cell surface.

The absence of an effect on RyRs and ICa,L occurrence in mouse myotubes confirms the peculiar action of neural agrin on the E-C coupling mechanism in humans.

Whether the modulation of RyR1 and L-type Ca2+ channels in humans was due to effects of agrin on expression levels and/or on posttranslational mechanisms was beyond the aims of the present experiments and remains to be investigated.

In conclusion, our findings provide new insights into the regulation of the E-C coupling mechanism during myogenesis. We propose that neural agrin favors the maturation of the E-C coupling machinery in developing human muscle cells by enhancing the occurrence of functional L-type voltage-dependent Ca2+ channels and possibly also of RyR1s. These effects of neural agrin could contribute to the comprehension of the molecular mechanism responsible for the peculiar nerve dependency reported for the maturation of the E-C coupling mechanism in human skeletal muscle cells. Our study also reveals new aspects of the functional role of the neural agrin. In addition to the well-known local synaptogenic effects, neural agrin is endowed with trophic functions on the extrasynaptic regions of the human muscle cells. We speculate that the long-range extrajunctional effects of neural agrin also occur in vivo and are relevant for the coordinated development and maturation of skeletal muscle fibers.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by grants from Fondazione Benefica Kathleen Foreman-Casali of Trieste to P. Lorenzon; from the Ministero dell'Istruzione, dell'Universita e della Ricerca-Italy to M. Sciancalepore, F. Ruzzier, and P. Lorenzon; and from Regione F.-V.G. to F. Ruzzier. This work was also supported by grants from the Agency of Science Republic of Slovenia and a Fogarty International Research Collaboration Award from the Fogarty International Center, National Institutes of Health to Z. Grubic.


   ACKNOWLEDGMENTS
 
We thank Anton Wernig (Institute of Physiology II, University of Bonn) for providing mouse muscle cells and A. Constanti (School of Pharmacy, University of London) for useful comments on the manuscript. We also thank E. Nurowska and G. Cellot for assistance in statistical data analysis and electrophysiological recordings.


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. Lorenzon, Dept. of Physiology and Pathology, Univ. of Trieste, Via A. Fleming 22, I-34127 Trieste, Italy (e-mail: pielle{at}dfp.units.it)

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.


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