Excitation–contraction coupling is the physiological mechanism occurring in muscle cells whereby an electrical signal sensed by the dihydropyridine receptor located on the transverse tubules is transformed into a chemical gradient (Ca2+ increase) by activation of the ryanodine receptor located on the sarcoplasmic reticulum membrane. In the present study, we characterized for the first time the excitation–contraction coupling machinery of an immortalized human skeletal muscle cell line. Intracellular Ca2+ measurements showed a normal response to pharmacological activation of the ryanodine receptor, whereas 3D-SIM (super-resolution structured illumination microscopy) revealed a low level of structural organization of ryanodine receptors and dihydropyridine receptors. Interestingly, the expression levels of several transcripts of proteins involved in Ca2+ homoeostasis and differentiation indicate that the cell line has a phenotype closer to that of slow-twitch than fast-twitch muscles. These results point to the potential application of such human muscle-derived cell lines to the study of neuromuscular disorders; in addition, they may serve as a platform for the development of therapeutic strategies aimed at correcting defects in Ca2+ homoeostasis due to mutations in genes involved in Ca2+ regulation.
Skeletal muscle is a highly differentiated tissue made up of myofibres, a syncytium of cells filled with myofibrils and containing sarcomeres that generate the force necessary for muscle contraction. In the last few years, a number of techniques have been developed to isolate single muscle fibres from small rodents allowing detailed investigations of the functional properties of the EC (excitation–contraction) coupling mechanism at the ultrastructural, biochemical and cellular levels in normal and pathological conditions [1–4]. Because of their high degree of differentiation and specialization, it is difficult to maintain differentiated muscle fibres in culture for more than a few days  and it is nearly impossible to obtain mature fibres starting from precursor satellite cells. Nevertheless, starting from newborn mice, one can obtain cultures of contracting and striated myotubes that can be used for a number of manipulations. As to human muscle cells, primary cultures can be obtained in vitro by culturing satellite cells from biopsies and differentiating them into myotubes [6–9], but there is a clear necessity to develop cell lines from control and diseased individuals which will develop into myotubes and which can be exploited as a platform for drug screening, and for biochemical, cellular and physiological characterization [10–13].
For the last two decades, our laboratories, as well as others, have established primary cultures from human biopsies and characterized their intracellular Ca2+ homoeostasis, with particular emphasis on EC coupling, how endogenous mutations in the RyR (ryanodine receptor) Ca2+ channel (RyR1) affect its functional properties and some downstream effects of Ca2+ dysregulation such as subcellular localization of the transcription factor NFAT (nuclear factor of activated T-cells), pro-inflammatory cytokine release and production of reactive nitrogen species [9,14,15]. However, the use of primary cultures has some inherent drawbacks mainly relating to the fact that they are generally slow-growing and will undergo a limited number of divisions. To overcome this problem, immortalization of human myogenic cells has been established both from a normal individual [16,17] and from patients with different neuromuscular diseases [17–20]. In the present study, we characterized a new cell line derived from a normal individual with no overt neuromuscular disorder. We show that the myotubes derived upon differentiation by serum withdrawal express the transcripts and protein components of the skeletal muscle EC coupling machinery. In addition, we established by 3D-SIM (super-resolution structured illumination microscopy) the subcellular distribution of RyR1 and of the DHPR (dihydropyridine receptor) and assessed Ca2+ release via RyR by using optical and electrophysiological techniques. This human skeletal muscle cell line HMCL-7304 is a tool of paramount importance to study on a larger scale the changes occurring in human muscles under a variety of pathological conditions.
The immortalized myoblast cell line was established from the intercostal skeletal muscle of a 19-year-old female donor with no neuromuscular disorder, by double transfection with recombinant retroviruses containing hTERT (human telomerase reverse transcriptase) cDNA and CDK4 (cyclin-dependent kinase 4) cDNA, as described previously [16,17]. The experiments were approved by the Author's Institutional Ethical Committee and were in accordance with the Declaration of Helsinki (2008) of the World Medical Association. The donor gave written informed consent to the work. Immortalized myoblasts were maintained in skeletal muscle cell growth medium (PromoCell) in a low-oxygen atmosphere (5% O2 and 5% CO2) at 37°C. In order to induce differentiation, once the density had reached approximately 80%, cells were rinsed once with PBS (pH 7.2) (Life Technologies) and incubated with skeletal muscle differentiation medium (PromoCell). The process of differentiation took 5 days on average, after which multinucleated myotubes were clearly visible under low magnification. Hereinafter, the human muscle derived-cell line will be referred to as HMCL-7304.
Ca2+ concentration measurements
HMCL-7304 cells were grown and differentiated on glass coverslips coated with 10 μg/ml laminin (Life Technologies). Once myotubes had formed, cells were loaded with the fluorescent Ca2+ indicator Fluo-4/AM (Fluo-4 acetoxymethyl ester) (Life Technologies) for 40 min at 37°C as described previously . Cells were rinsed once with Krebs–Ringer medium (pH 7.4) containing 2 mM CaCl2 and then coverslips were mounted on to a 37°C thermostatically controlled chamber which was continuously perfused with Krebs–Ringer medium (pH 7.4); individual cells were stimulated with the indicated agonists (KCl, 4-chloro-m-cresol, caffeine) made up in Krebs–Ringer medium (pH 7.4) containing no added Ca2+ plus 100 μM La3+ in order to monitor changes in the free cytosolic Ca2+ concentration ([Ca2+]i) due to release from intracellular stores. Individual cells were stimulated by means of an eight-way 100-mm-diameter quartz micromanifold computer-controlled microperfuser (ALA Scientific Instruments), as described previously . Online fluorescence images were acquired using an inverted Nikon TE2000 TIRF (total internal reflection fluorescence) microscope equipped with a dry Plan Apochromat ×20 objective [0.17 NA (numerical aperture)] and an electron multiplier Hamamatsu CCD (charge-coupled device) camera C9100-13. Changes in [Ca2+]i were analysed using the MetaMorph imaging system (Molecular Devices) and the average pixel value for each cell was measured as described previously [14,15].
Electrophysiological measurements and confocal Ca2+ imaging
Myoblasts were grown on laminin-coated glass-bottomed 35-mm-diameter dishes (MatTek). After differentiation into myotubes, cells were patch-clamped in the whole-cell configuration with low-resistance borosilicate glass micropipettes (1–3 MΩ) using an Axopatch 200B amplifier (Axon Instruments) controlled by a custom-written data-acquisition software developed by LantibodiesView (National Instruments). The external solution contained 150 mM triethylammonium methylsulfonate, 2 mM CaCl2, 1 mM MgCl2, 10 mM Hepes, 0.001 mM TTX (tetrodotoxin) and 1 mM 4-aminopyridine; the pH was adjusted to 7.4 with CsOH. The pipette solution contained 140 mM CsCH3SO3, 10 mM Hepes, 6 mM MgCl2, 11.5 mM CaCl2, 4 mM Na2ATP, 20 mM EGTA, 14 mM CrPO4, 0.1 mM leupeptin and 0.1 mM Fluo-3 potassium salt; the pH was adjusted to 7.2 with CsOH . The reference electrode was connected to the bath solution with an agar bridge (4% agar in 3 M KCl). All measurements were carried out at room temperature (25°C). Holding potential was kept at −80 mV. Stepwise depolarizations were applied to activate the voltage-dependent DHPR and to trigger Ca2+ release mediated by RyR1, the intracellular Ca2+-release channel located in the SR (sarcoplasmic reticulum) membrane. Detailed voltage-clamp protocols are indicated. In order to determine the current–voltage relationship, long depolarizations of 800 ms were applied. Currents were analysed in IgorPro (Wavemetrics). Peak current amplitude was calculated from the difference between membrane currents before and during application of 500 mM CdCl2 (ΔICaL). For the investigations of electromechanical coupling between the DHPR and RyR1, short depolarizations (50 ms) were applied. Changes in the [Ca2+]i were monitored using the fluorescent Ca2+ indicator Fluo-3 (Biotium), which was loaded into the cell via the patch pipette. Using a MicroRadiance laser-scanning confocal microscope (Bio-Rad Laboratories), Fluo-3 was excited at 488 nm with an argon ion laser, and emission light was collected above 500 nm. Linescan images were recorded at a rate of 50 lines/s, analysed in ImageSXM (free software based on NIH Image)  and processed further using IgorPro. Changes in [Ca2+]i are shown as relative changes in fluorescence (ΔF/F0).
qPCR (quantitative real-time PCR) experiments
Total RNA was extracted from differentiated HMCL-7304 myotubes and biopsies from healthy individuals, using TRIzol® (Life Technologies) as described previously . cDNA was synthesized with the High Capacity cDNA synthesis kit (Applied Biosystems) and the primers listed in Table 1. Transcript levels were quantified using SYBR® Green reagent on an Applied Biosystems platform (7500 fast real-time PCR system) and levels were normalized to desmin expression.
|Primer||Forward primer sequence||Reverse primer sequence|
|Primer||Forward primer sequence||Reverse primer sequence|
Mouse muscle fibre isolation
Mouse FDB (flexor digitorum brevis) fibres were enzymatically dissociated at 37°C for 60 min in a cell culture incubator in Tyrode's solution containing 0.20% collagenase I (Sigma Fine Chemicals) and placed on glass coverslips coated previously with 1.5 μl of laminin (1 mg/ml) (Life Technologies) as described previously .
For immunofluorescence, myotubes cultured on glass coverslips were fixed with 4% (w/v) paraformaldehyde at room temperature for 15 min, washed and permeabilized with 0.1% Triton X-100 in PBS for 1 h before blocking with 3% (v/v) goat serum for 30 min, followed by 1 h of incubation in mouse anti-(fast myosin) monoclonal antibody (1:100 dilution; Sigma Fine Chemicals, catalogue number M4276) and mouse anti-(slow myosin) monoclonal antibody (1:50 dilution; Novo NCL, Leica Biosystems, catalogue number NCL-MHC) at room temperature. Coverslips were washed in 0.1% Triton X-100 in PBS and incubated with goat anti-(mouse IgG) secondary antibody conjugated to Alexa Fluor® 594 (Life Technologies, catalogue number A-11005) for 1 h at room temperature, followed by thorough washing in PBS. Nuclei were stained with Hoechst 33258 (Invitrogen) for 10 min. Slides were mounted in aqueous mounting medium and viewed with a Leica DMR fluorescent microscope equipped with a ×20 HC-PL Fluotar objective (0.50 NA). The percentage of fast and slow myosin-positive HMCL-7304-derived myotubes was calculated by counting the number of myosin-positive cells divided by the total number of cells as visualized by brightfield microscopy in the same area. For super-resolution microscopy, mouse FDB fibres or human skeletal muscle myotubes were fixed with 3.7% (w/v) paraformaldehyde (in PBS) for 30 min at room temperature, rinsed twice with PBS and permeabilized with 1% Triton X-100 in PBS for 30 min. After rinsing and blocking non-specific sites with 1:100 blocking solution (Roche), cells and fibres were incubated with anti-RyR1 monoclonal antibody (Thermo Scientific, catalogue number MA3-925) or goat anti-Cav1.1 (Santa Cruz Biotechnology, catalogue number sc-8160) antibody for 3 h at room temperature (10 μg/ml final concentration diluted in PBS with 0.01% Tween 20). Slides were rinsed with PBS containing Tween 20 five times for 5 min each and incubated with the appropriate secondary conjugate (Alexa Fluor® 555 or Alexa Fluor® 647, diluted 1:500 in PBS containing Tween 20) overnight at 4°C. Slides were rinsed with PBS containing Tween 20 and mounted with 10% (v/v) glycerol in PBS. Staining was visualized with a Zeiss Elyra microscope equipped with ×63 oil Plan-Apochromat (1.4 NA) objective. Raw datasets consisted of images acquired with three different grid angles and five different grid phases. Super-resolution images were calculated from the raw data using the built-in algorithm. The baseline was just shifted (not truncated) to allow inspection for potential ghosts arising from sample imperfections.
Electrophoresis and immunoblotting
Microsomes were prepared from differentiated HMCL-7304-derived myotubes as described previously . Protein concentration was determined using Protein Assay Kit II (Bio-Rad Laboratories) using BSA as a standard. SDS/PAGE, protein transfer on to nitrocellulose membranes and immunostaining were performed as described previously . The following primary antibodies were used: mouse anti-RyR1 (Thermo Scientific, catalogue number MA3-925), rabbit anti-RyR1 (a gift from Professor Vincenzo Sorrentino, University of Siena, Siena, Italy), goat anti-Cav1.1 (Santa Cruz Biotechnology, catalogue number sc-8160), rabbit anti-CSQ1 (calsequestrin 1) (Sigma, catalogue number C0743), rabbit anti-CSQ2 (Epitomics, catalogue number 2962-1), goat anti-SERCA1 (sarcoplasmic/endoplasmic reticulum Ca2+-ATPase 1) (Santa Cruz Biotechnology, catalogue number sc-8093), goat anti-SERCA2 (Santa Cruz Biotechnology, catalogue number sc-8095) and mouse anti-MYH1 (myosin heavy chain 1) (Millipore, catalogue number 05-716). Secondary peroxidase conjugates were Protein G–peroxidase (Sigma, catalogue number P8170) and peroxidase-conjugated goat anti-(mouse IgG) (Sigma, catalogue number A2304). The immunopositive bands were visualized by chemiluminescence using the Super Signal West Dura kit (Thermo Scientific).
Statistical analysis and graphical software
Statistical analysis was performed using Student's t test; means were considered statistically significant when the P value was <0.05. When more than two groups were compared, analysis was performed by the ANOVA test followed by the Bonferroni post-hoc test using GraphPad Prism 4.0 software. Origin software was used to generate dose–response curves and obtain EC50 values; images were assembled using Adobe Photoshop CS (version 8.0).
Expression levels of transcripts and proteins involved in EC coupling in cell line-derived myotubes
In order to functionally characterize the HMCL-7304-derived myotubes, we chose several genes that are well known to play a crucial role in skeletal muscle EC coupling. Transcript levels were quantified and compared with those present in muscle biopsies obtained from five healthy individuals. Figure 1(A) shows the relative expression levels on a logarithmic scale of different transcripts. Interestingly, we found that the RYR1 transcript was significantly lower (approximately 300-fold) than in differentiated muscle (P<0.0001), and there was no up-regulation of RYR3 mRNA, which was also significantly reduced in HMCL-7304 myotubes. SERCA1 and CSQ1 transcript levels were significantly lower (~1000-fold) in the cell line (P<0.003), whereas there was a 10-fold increase in the expression of the CSQ2 transcript. SERCA2 showed similar mRNA levels in the biopsies and the cell line as did CAV1.1, suggesting that expression of the L-type Ca2+ channel may appear at an early stage of development. The transcript levels of MYH1 and MYH2, that are characteristically expressed in slow-twitch and fast-twitch muscles respectively were lower in HMCL-7304 compared with biopsies; however, immunofluorescence shows that the slow myosin isoform is present in a larger percentage of HMCL-7304 cells, compared with fast myosin, with approximately 51% slow to 13% fast (36% negative for both fast and slow isoforms) (Figure 1B). Taken together, these results suggest that the differentiated myotubes express proteins that are more abundant in slow-twitch muscles.
Expression of EC coupling-associated proteins in HMCL-7304-derived myotubes
We are aware that the presence of a transcript does not necessarily reflect protein expression, thus we tested microsomes prepared from HMCL-7304-derived myotubes by immunoblotting. Figure 1(C) confirms that MYH1, SERCA1, SERCA2, Cav1.1 and RyR1 were all expressed. The double-immunopositive band seen in the RyR1 Western blot does not represent RyR3, since both bands were present when a RyR1-specific polyclonal antibody was used, thus the lower band is probably a degradation product. No bands were visualized when blots were probed with anti-CSQ1 antibodies (not shown), but a band migrating with an approximate molecular mass of 55 kDa (asterisk) was present when myotube microsomes were probed with anti-CSQ2 antibodies.
Cellular localization of EC coupling proteins
In mature skeletal muscle, EC coupling occurs and is fully dependent on the architecture of highly structured intracellular Ca2+ release units, whereby the voltage-sensing L-type Ca2+ channel is present in the transverse tubules and faces the terminal cisternae of the SR containing the RyR1 Ca2+ release channels. The SR contains the Ca2+-binding protein CSQ, whereas the SERCAs are located on the longitudinal SR. In order to define the cellular localization of these proteins in mature myotubes, we used a 3D-SIM microscope, because, compared with conventional confocal fluorescence microscopy and STED (stimulated emission depletion) microscopy, it offers improved axial resolution (approximately 300 nm) . Figure 2 shows the immunostaining of Cav1.1 and RyR1 both in myotubes (panels A–F) and in enzymatically dissociated mouse FDB fibres (panels G–I). As expected in FDB fibres, the staining with anti-Cav1.1 shows a double row of fluorescent particles (Figure 2G) that mostly overlap with particles which are stained with anti-RyR1 antibodies (Figure 2H). Human myotubes also show fluorescent particles whose distribution does not follow the highly regular pattern observed in mature FDB fibres. However, when observed at high magnification, it becomes apparent that human myotubes display areas containing multiple parallel longitudinal rows of fluorescence particles which are stained with anti-Cav1.1 antibodies (Figure 2D, double arrows). A large fraction of particles stained with anti-Cav1.1 antibodies co-localize with particles stained with anti-RyR1 antibodies (Figure 2F, arrowheads) and may represent Ca2+-release units involved in EC coupling. In the next set of experiments, we assessed the EC coupling characteristics of HMCL-7304-derived myotubes, by studying RyR-mediated Ca2+ release and Cav1.1-mediated Ca2+ currents either by optical methods or by using the patch-clamp technique in the whole-cell configuration.
Cellular localization of RyR1 and Cav1.1 in differentiated myotubes compared with mouse FDB fibres by 3D-SIM microscopy
Functional properties and pharmacological activation of Ca2+ release
Figure 3 shows the results obtained in cells loaded with the fluorescent Ca2+ indicator Fluo-4; stimulation of cells with 60 mM KCl caused an immediate increase in the resting [Ca2+]i which decayed back to resting values within approximately 5 s (Figure 3B); the peak increase in [Ca2+]i was similar irrespective of whether cells were stimulated via activation of the DHPR L-type Ca2+ channel by KCl-induced depolarization, or by direct activation of the RyR1 with 4-chloro-m-cresol and caffeine (Figure 3C). Figures 3(D), 3(E) and 3(F) show concentration- dependent peak Ca2+-release curves elicited by caffeine, 4-chloro-m-cresol and KCl respectively, as well as the calculated EC50 values. These results are similar to those obtained in primary human muscle myotubes explanted from biopsies from control individuals [14,15].
Characterization of RyR1-mediated Ca2+ release in HMCL-7304-derived myotubes
We then investigated the voltage-dependence of membrane currents (ICaL) from the DHPR in voltage-clamped HMCL-7304 myotubes. Starting from a holding potential of −80 mV and an initial pre-step to −40 mV, cells were stimulated by repetitive depolarizing steps of 800 ms in 10 mV increments to increasing membrane potentials. Figure 4(A) shows five steps of the stimulation protocol (upper trace) and the corresponding membrane currents (below) during control conditions (trace a, continuous line) and during inhibition of ICaL with Cd2+ (trace b, dotted line). The calculated difference current (ΔICaL, trace c) is indicated by the dashed line and peak current amplitudes at the end of the depolarizing steps are used for further analysis. Figure 4(B) summarizes the voltage-dependence of current activation and reveals a half-maximal activation (V1/2) at −9 mV.
Skeletal L-type Ca2+ currents in HMCL-7304
A parallel investigation of electromechanical coupling was achieved using a combination of voltage-clamp and confocal Ca2+ imaging. Cells were imaged in the linescan mode at close proximity to the position of the patch pipette. Membrane depolarization triggered significant Ca2+ release, indicating functional coupling between the DHPR and RyR1. Depolarization-induced Ca2+ release was monitored and recorded in linescan images. Figure 5(A) shows a voltage-clamped HMCL-7304 myotube loaded with Fluo-3; the position of the linescan is indicated in the illustrated myotube. Short depolarization (50 ms) starting from the holding potential of −80 mV to +10 mV activated the DHPR and triggered Ca2+ release from the SR. The resulting Ca2+ transient is displayed in the linescan image and in the corresponding line profile. Of note is the slow return of the [Ca2+]i to resting levels, which might be consistent with a low level of SERCA1 expression in immature myotubes (see Figure 1A). As expected, increasing membrane depolarization results in greater Ca2+ release, which saturates at a membrane potential of approximately 0 mV. Figure 5(B) shows the Ca2+ transient amplitude in response to increasing depolarization in a single protocol, and Figure 5(C) summarizes the voltage-dependence of Ca2+ release. In order to minimize side effects from long periods of scanning and accumulation of cytosolic Ca2+ due to slow Ca2+ extrusion processes, data points for peak Ca2+ transients in Figure 5(C) have been collected from repeated short linescan recordings (6 s). Ca2+ transients elicited under voltage clamp revealed a half-maximal release activation (V1/2) at −32 mV.
Depolarization-induced Ca2+ release in HMCL-7304 myotubes
In the present paper, we describe the biochemical, cellular and physiological characteristics of a novel cell line generated from human skeletal muscle. The necessity to establish cell lines of human origin from ‘normal’ individuals or patients affected by a number of neuromuscular conditions has been apparent for the last few decades, but early attempts to immortalize human myoblasts capable of differentiating into mature myotubes were unsuccessful (for example, see [26,27]). In the present paper, we report the successful myotube differentiation from myoblasts of the newly generated immortal human muscle cell line HMCL-7304 derived from a healthy individual and characterize its features at the biochemical, structural and functional levels, showing that its phenotype is similar to myotubes from primary cultures.
In the last few years, a number of groups have exploited vectors expressing hTERT and CDK4 and have reported successful immortalization of myoblasts from ‘normal’ individuals and individuals affected by various forms of muscular dystrophies and dysferlinopathies [16–20,28]. The possibility of generating such immortalized cell lines able to differentiate into myotubes constitutes an important tool to investigate human neuromuscular diseases much as the murine skeletal muscle C2C12 cell line has been exploited in the last few decades by laboratories worldwide to investigate key aspects of skeletal muscle physiology and plasticity. Therefore there is high demand for a reliable human skeletal cell line to fill the gap between understanding the pathophysiological mechanisms and the development of therapeutic strategies for human neuromuscular disorders. Nevertheless, although the use of an immortalized cell line has many advantages over primary cultures, the process of immortalization is known to modify many physiological parameters ranging from differentiation and secretion, to the expression of specific protein isoforms. For this reason, we characterized the EC coupling machinery and Ca2+ homoeostasis of HMCL-7304-derived myotubes. We chose to verify the levels of expression of the transcripts encoding the main components of skeletal muscle EC coupling, as well as their main isoforms and compared them with the levels found in biopsies from human skeletal muscle fibres from normal individuals. Transcripts encoding RyR1, CSQ1 and SERCA1 were significantly down-regulated as was RyR3, an isoform that is thought to be more abundantly expressed in developing muscles. On the other hand, Cav1.1 was expressed to similar levels in both mature fibres and myotubes and produced large Ca2+ currents, as shown in patch–clamp measurements. These results are in contrast with what occurs during skeletal muscle development, where components of the SR appear at an earlier stage of development compared with the transverse tubules containing the DHPR L-type Ca2+ channel [29–31]. Interestingly, the relative mRNA expression of HMCL-7304-derived myotubes was either similar to that of mature biopsies (SERCA2) or up-regulated (CSQ2). Thus it seems that the HMCL-7304 cell line has more of the characteristics of slow-twitch than that of fast-twitch muscle. This is confirmed further by the predominant expression of slow myosin compared with fast myosin in differentiated HMCL-7304 myotubes. There are several possible reasons for this observation: (i) either satellite stem cells are, by default, slow-twitch-like and it is the influence of innervation or electrical activity that enables them to become fast or slow , (ii) the immortalization procedure ‘selects’ satellite stem cells that have a ‘slow muscle’ phenotype, and/or (iii) as reported in hindlimb muscles of adult rats, the satellite cells contained within fast or slow twitch fibres are intrinsically different subpopulations  so that the HMCL-7304 cell line that originated from dorsal muscles resembles more a slow-twitch muscle from which it originated.
Two noticeable differences in the physiological characteristics of human cultured myotubes and mature fibres are that human (as opposed to rodent) -derived primary myotubes do not contract , and the timescale of the depolarization-induced Ca2+ transient occurs in hundreds of milliseconds in myotubes, whereas it lasts only a few milliseconds in mature fibres. Although the lack of contracture of myotubes derived from the immortalized human muscle cell line probably reflects the composition of the actomyosin contractile machinery, the slow Ca2+ transients observed in myotubes probably reflects the lower level of expression of SERCA1 and the absence of a highly organized micro-architecture. After Ca2+ release, cytosolic Ca2+ is rapidly removed and pumped back into the SR lumen by the SERCA, which is the main protein of non-junctional SR in skeletal muscle fibres . Reduced SERCA1 expression levels as shown in the HMCL-7304 cell line have a significant impact on the Ca2+-removal kinetics, leading to deceleration of Ca2+ re-uptake compared with normal muscle fibres. Thus lack of mature subcellular architectural organization and reduction in SERCA1 expression may explain the slow Ca2+ removal after release in the HMCL-7304 cell line.
In mature fibres, four Cav1.1 subunits on the T-tubular face in square formation, referred to as tetrads, are directly opposite the four subunits of one RyR1 tetramer on the SR junctional membrane, to form the Ca2+-release units . In intact FDB fibres, the Ca2+ release units form double rows on each side of the Z line. The highly organized arrangement is characteristic of mature skeletal muscle and is probably one of the features allowing the extremely rapid Ca2+ release kinetics upon T-tubule membrane depolarization. As is evident by the 3D-SIM images, HMCL-7304 myotubes show distinct rows of Cav1.1 that overlap, at least in part, with RyR1 and we believe that these structures may represent the Ca2+-release units of the HMCL-7304 myotubes. This conclusion is consistent with whole patch-clamp measurements showing that voltage-induced Ca2+ release is highly functional in these myotubes, indicating direct coupling between sarcolemmal DHPRs and RyR1 despite structural immaturity. HMCL-7304 myotubes exhibit Cd2+-sensitive Ca2+ currents, having half-maximal current activation at −9 mV, a value that is comparable with obtained by others [36,37]. Parallel imaging revealed that the voltage-dependence of Ca2+ release (V1/2) was −32 mV in the present study and −29.4 mV in non-immortalized primary human myotubes from control individuals , which is similar to the values (−26 mV) obtained on primary myotubes from wild-type mice . Furthermore, the process of immortalization does not affect the pharmacological characteristics of RyR1 activation since the caffeine and 4-chloro-m-cresol dose–response curves of the HMCL-7304 cell line are similar to those of non-immortalized human myotubes [14,40] or RyR1 isolated from mature rodent muscles .
In conclusion, in the present study, we characterized a human muscle cell line derived from a ‘normal’ individual and show that it retains a pattern of expression of proteins involved in EC coupling similar to that of slow-twitch muscles. The ability to perform pharmacological and electrophysiological studies illustrates the potential use of such a biological tool to a variety of approaches from studying the effect of mutations and gene silencing to testing drugs and pharmacological agents aimed at correcting Ca2+ dysregulation as occurs in a variety of neuromuscular disorders including core myopathies.
free cytosolic Ca2+ concentration
cyclin-dependent kinase 4
super-resolution structured illumination microscopy
flexor digitorum brevis
human telomerase reverse transcriptase
myosin heavy chain
quantitative real-time PCR
sarcoplasmic/endoplasmic reticulum Ca2+-ATPase
Ori Rokach performed experiments and analysed data; Nina Ullrich performed electrophysiological experiments and analysed the data; Martin Rausch performed the 3D-SIM experiments; Vincent Mouly established the immortalized cell line; Haiyan Zhou designed experiments and analysed data; Francesco Muntoni designed experiments and analysed data; Francesco Zorzato performed the 3D-SIM experiments, took care of the conception and design of the experiments, collection and analysis of data, and drafting of the paper; Susan Treves took care of the conception and design of the experiments, collection and analysis of data, and drafting of the paper.
We thank Professor Vincenzo Sorrentino for kindly providing the polyclonal rabbit anti-RyR1 antibody.
The support of the Medical Research Council Neuromuscular Centre and the Great Ormond Street Hospital Biomedical Research Centre to the Biobank is gratefully acknowledged. This work was supported by the Swiss National Science Foundation [grant number 3100 030_129785 (to S.T.)], Swiss National Science Foundation-Ambizione [grant number PZ00P3_131987 (to N.D.U.)], Muscular Dystrophy Association [grant number 174047 (to F.M.)] and the Botnar Stiftung (to O.R.). The support of the Muscular Dystrophy Association [grant number MDA68762 (to F.M.)] and the platform for immortalization of human cells from the Myologie Institute in Paris are also gratefully acknowledged. F.M. is supported by the Great Ormond Street Hospital Children's Charity.
These authors contributed equally to this work.