Intracellular Ca2+-handling differs markedly between intact human muscle fibers and myotubes
© Olsson et al. 2015
Received: 27 May 2015
Accepted: 21 July 2015
Published: 20 August 2015
In skeletal muscle, intracellular Ca2+ is an important regulator of contraction as well as gene expression and metabolic processes. Because of the difficulties to obtain intact human muscle fibers, human myotubes have been extensively employed for studies of Ca2+-dependent processes in human adult muscle. Despite this, it is unknown whether the Ca2+-handling properties of myotubes adequately represent those of adult muscle fibers.
To enable a comparison of the Ca2+-handling properties of human muscle fibers and myotubes, we developed a model of dissected intact single muscle fibers obtained from human intercostal muscle biopsies. The intracellular Ca2+-handling of human muscle fibers was compared with that of myotubes generated by the differentiation of primary human myoblasts obtained from vastus lateralis muscle biopsies.
The intact single muscle fibers all demonstrated strictly regulated cytosolic free [Ca2+] ([Ca2+]i) transients and force production upon electrical stimulation. In contrast, despite a more mature Ca2+-handling in myotubes than in myoblasts, myotubes lacked fundamental aspects of adult Ca2+-handling and did not contract. These functional differences were explained by discrepancies in the quantity and localization of Ca2+-handling proteins, as well as ultrastructural differences between muscle fibers and myotubes.
Intact single muscle fibers that display strictly regulated [Ca2+]i transients and force production upon electrical stimulation can be obtained from human intercostal muscle biopsies. In contrast, human myotubes lack important aspects of adult Ca2+-handling and are thus an inappropriate model for human adult muscle when studying Ca2+-dependent processes, such as gene expression and metabolic processes.
KeywordsHuman skeletal muscle Ex vivo model Intact human muscle fibers Satellite cells Myoblasts Excitation-contraction coupling Force production Gene expression Metabolic processes
Satellite cells are resident stem cells of the skeletal muscle that can be isolated from human muscle biopsies, activated into proliferating myoblasts, and differentiated into multinuclear myotubes ex vivo [1, 2]. Myotubes have been credited with morphological, metabolic, and biochemical properties similar to those in adult skeletal muscle fibers [3, 4]. This in combination with the difficulty of obtaining intact viable muscle fibers from humans have rendered myotubes a widely used model to study aspects of cell signaling and metabolism in human skeletal muscle [3, 5–8].
In addition to their reported similarities to adult muscle fibers, myotubes also exhibit features that distinguish them from adult muscle. These include the expression of immature forms of muscle proteins , differences in abundance and distribution of glucose transporters [10, 11], and dominance of anaerobic glycolysis  as compared with adult muscle fibers. Furthermore, although human myotubes can obtain contractile ability under specific culture conditions, generated contractile forces are lower, fused tetani are attained at lower stimulation frequencies, and kinetic parameters are slower than those in adult muscle fibers [12–14]. This suggests an immature intracellular Ca2+-handling in myotubes, a notion supported by findings in previous studies investigating the Ca2+-handling properties of human myotubes [15–17]. To date, there is a lack of knowledge regarding the quantitative aspects of these differences since no direct comparison between human intact muscle fibers and myotubes has been previously performed.
Extensive evidence demonstrates the importance of intracellular Ca2+ for the regulation of a wide range of skeletal muscle processes, including muscle force generation, gene expression, and cellular metabolism [18–21]. Given the importance of cytosolic free [Ca2+] ([Ca2+]i) transient amplitude and duration  as well as frequency of [Ca2+]i transients  for the downstream effects, differences in Ca2+-handling properties will have important implications when translating knowledge gained in myotubes to adult muscle fibers. Thus, we considered that a thorough characterization of the intracellular Ca2+-handling in human muscle fibers and myotubes was warranted in order to quantify the extent to which human myotubes differ in their Ca2+-handling properties compared to adult muscle fibers.
In the present study, we hypothesized that human muscle fibers and myotubes would display major functional differences in intracellular Ca2+-handling. To study this hypothesis, we developed a model of dissected intact muscle fibers obtained from human intercostal muscle biopsies and compared the intracellular Ca2+-handling of myotubes with that in adult muscle fibers. In addition to functional Ca2+-handling properties, we studied differences in gene expression, protein quantity and localization of Ca2+-handling proteins, and ultrastructural differences between the adult muscle fibers and myotubes. We show that viable, intact single muscle fibers can be obtained from human intercostal muscle biopsies and that these fibers display strictly regulated [Ca2+]i transients and force production upon electrical stimulation ex vivo. In contrast, myotubes display only a rudimentary development of Ca2+-handling properties and do not contract. Consistent with this, we identify functionally important differences in the quantity and localization of Ca2+-handling proteins and ultrastructure between human muscle fibers and myotubes.
Six recreationally active individuals suffering from lung cancer, four males and two females (age 65–76 years), scheduled for lobectomy at the Karolinska University Hospital in Stockholm, Sweden, were recruited for experiments on intercostal muscles. For patient characteristics see Additional file 1: Table S1. For the experiments on myoblasts and myotubes from vastus lateralis muscle biopsies, eight healthy recreationally active individuals, four males and four females (age 21–27 years), were recruited. The planned experiments and procedures were explained before subjects gave their written informed consent to participate. The study was approved by the Regional Ethical Review Board in Stockholm for experiments on intercostal muscles (Dnr 2012/2181-31/2) and for the experiments on myoblasts and myotubes (Dnr 2012/173-31/3).
External and internal intercostal muscle biopsies were obtained during thoracotomies. Biopsies were sampled at the midaxillary line in the fourth intercostal space during thoracoscopy or in the fifth intercostal space during open surgical procedures. Biopsies were collected with intact periostium at both ends to ensure non-disrupted muscle fibers and preserved tendons at both ends. Biopsies were placed in Dulbecco’s modified eagle medium (DMEM) (Gibco® Invitrogen, Life Technologies, Carlsbad, CA, USA) containing 0.2 % fetal bovine serum (FBS) (Gibco® Invitrogen, Life Technologies) bubbled with a mixture of 95 % O2 and 5 % CO2 prior to collection and immediately transported to the laboratory. Vastus lateralis muscle biopsies were obtained by the Bergström percutaneous needle biopsy technique .
Muscle fiber dissection and mounting
Human internal and external intercostal muscle fibers are 0.5–1 cm long with diameters ranging from 30 to 100 μm. Intact single muscle fibers with intact tendons were dissected as previously described for mouse toe muscle fibers . The isolated intercostal muscle fiber was mounted in a stimulation chamber between an Akers 801 force transducer and an adjustable holder, and the length was adjusted to that giving maximal tetanic force. The fiber was superfused with Tyrode solution (mM): NaCl, 121; KCl, 5.0; CaCl2, 1.8; MgCl2, 0.5; NaH2PO4, 0.4; NaHCO3, 24.0; EDTA, 0.1; glucose, 5.5. FBS (0.2 %) was added to the solution to improve cell survival. The solution was bubbled with a mixture of 5 % CO2 and 95 % O2, which gives an extracellular pH of 7.4. Experiments were performed at room temperature (approximately 24 °C).
Muscle fiber force and [Ca2+]i measurements
Isolation of myoblasts and cell culture
Human myoblasts were extracted from fresh muscle as previously described , with some modifications. Immediately following the biopsy procedure, approximately 100 mg of muscle tissue was placed in sterile phosphate buffered saline (PBS) (Gibco® Invitrogen, Life Technologies) containing 1 % antibiotic-antimycotic (ABAM) (Gibco® Invitrogen, Life Technologies) and incubated at 4 °C overnight. The following day, the muscle biopsy was incubated in 5 ml TrypLE™ Express Enzyme (1X) (Gibco® Invitrogen, Life Technologies) at 37 °C, 5 % CO2 with gentle agitation for 20 min. Undigested tissue was allowed to settle for 5 min at room temperature, and the supernatant containing the satellite cells were collected in 5 ml DMEM-F12 Glutamax (Gibco® Invitrogen, Life Technologies), containing 20 % FBS and 1 % ABAM. Digestion of the slurry was repeated twice. Subsequently, the supernatant containing the satellite cell population was collected. Isolated human myoblasts were cultured in DMEM-F12 Glutamax containing 20 % FBS and 1 % ABAM at 37 °C, 5 % CO2. Culture dishes were coated with collagen I (Collagen I, Bovine 5 mg/mL, Gibco® Invitrogen, Life Technologies) diluted to a final concentration of 50 μg/mL in 0.02 M acetic acid according to the manufacturer’s manual. Myoblasts were taken through serial passages to increase cell numbers prior to experimentation. All myoblasts were used for experimentation at passage 3–4. For experimentation, myoblasts were harvested and transferred to collagen I coated 35 mm glass-bottomed petri dishes (P35G-0-14-C, MatTek, Ashland, MA, USA) at a density of 8 × 104 cells per dish. Myoblasts were allowed to settle for 24 h after which half of the dishes with plated myoblasts were used for intracellular Ca2+ measurements while the other half was differentiated into myotubes. Myotube differentiation was promoted by substituting the proliferation media with DMEM-F12 Glutamax containing 2 % FBS, 25 pM Insulin (I9278, Sigma-Aldrich, St. Louis, MO, USA) and 1 % ABAM.
Immunomagnetic cell sorting
Enrichment of the cell population for myogenic cells was accomplished by a combination of pre-plating with magnetic activated cell sorting (MACS) separation as has previously been reported to produce a high yield of myogenic cells . MACS separation of myogenic and non-myogenic cells was carried out as previously described for human muscle derived cells , with some modifications. Briefly, muscle derived cells plated in T75 flasks were incubated with primary antibody for CD56 (MY31, BD Biosciences, Franklin Lakes, NJ, USA) dissolved in DMEM-F12 Glutamax for 30 min at 37 °C, 5 % CO2. Cells were subsequently pelleted and re-suspended in PBS containing 0.1 % FBS and microbeads (Miltenyi Biotech, Lund, Sweden). Cell suspension was incubated in the dark at 4 °C for 15 min before being rinsed with PBS containing 0.1 % FBS and re-pelleted. Cells were magnetically separated using a midiMACS magnet and LS column (Miltenyi Biotech). The cells that were bound to the anti-CD56 microbeads complex were maintained in the column and constituted the positive (myogenic) fraction of cells. This fraction was subsequently plated and used for experimentation.
Validation of myogenic origin of human myoblasts and myotube differentiation
At the time of plating cells for experimentation, a fraction of myoblasts was collected for confirmation of myogenic origin. Cells were spun down onto a cover glass for subsequent immunofluorescent staining of the myogenic marker desmin (D33, DAKO, Glostrup, Denmark). The fraction of desmin positive cells in the cell population was analyzed by dividing with the total number of nuclei stained with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) (Invitrogen/Molecular Probes) within each field. In the current study, 94 % (±1.32) of sub-confluent myoblasts were positive for desmin. To confirm myotube differentiation, cells were visually inspected and the presence of multinucleated (>2 nuclei) elongated cells positive for desmin was verified (Additional file 2: Figure S1A). Differentiation was further confirmed by an increase (P = 0.007) in mRNA levels of myogenin, a myogenic transcription factor involved in the terminal differentiation of myotubes  (Additional file 2: Figure S1B).
Myoblast and myotube [Ca2+]i measurements
In human myoblasts and myotubes, [Ca2+]i was measured with the non-ratiometric fluorescent Ca2+ indicator fluo-3, which was loaded into cells in the acetoxymethyl ester form (Fluo-3 AM, Invitrogen, Life Technologies) and confocal microscopy using a modified Bio-Rad MRC 1024 unit attached to a Nikon Diaphot inverted microscope with a Nikon Plan Apo 20X objective (NA 1.3). Stored confocal images were analyzed with ImageJ (National Institutes of Health, http://rsb.info.nih.gov/ij). To enable comparisons between cells, changes in the fluo-3 fluorescence signal (ΔF) were divided by the fluorescence immediately before a stimulation pulse was given (F0).
Cell stimulation protocol
Electrical stimulation was achieved by supramaximal current pulses (duration 0.5 ms) delivered from a lab built electrical stimulator via a pair of platinum plate electrodes lying parallel to the muscle fibers or along the edge of the glass bottom of the petri dish facing each other for myoblasts/myotubes. For chemical stimulation, myoblasts and myotubes were superfused for 1 min with Tyrode solution containing 5 mM adenosine triphosphate (ATP) (Sigma-Aldrich) or 1 mM 4-chloro-m-cresol (4-CmC) (Sigma-Aldrich). ATP- and 4-CmC-elicited changes in ΔF/F0 in myoblasts and myotubes were evaluated by comparing with changes in ΔF/F0 during exposure to normal Tyrode solution for the same time period.
Quantification of the Ca2+ decay time constant
The Ca2+ decay time constant was analyzed by fitting a single exponential function plus a constant to the [Ca2+]i transients of human myotubes and intercostal fibers stimulated with a single 70 Hz, 350 ms train of current pulses. For intercostal muscle fibers, original fluorescent ratios were smoothed one to two times using a seven-point quadratic polynomial to reduce noise prior to calculating [Ca2+]i by Eq. 1. The peak Δ[Ca2+]i/[Ca2+]i0 for intercostal fibers or ΔF/F0 for myotubes, respectively, after the end of the stimulation period was set as the first point of the fit interval. The fit was continued until baseline (defined as <5 % of peak Δ[Ca2+]i/[Ca2+]i0 or ΔF/F0) was reached.
Gene expression analysis
Total RNA was prepared by the Trizol method (Invitrogen, Life Technologies) and quantified spectrophotometrically by absorbance at 260 nm. One μg of total RNA from each sample was used for reverse transcription into cDNA for a final volume of 20 μl (High Capacity Reverse Transcription Kit, Applied Biosystems, Life Technologies). Real-time PCR (ABI-PRISMA 7700 Sequence Detector, Perkin-Elmer, Applied Biosystems, Life Technologies) procedures were employed to determine mRNA expression. Probes and primers (TaqMan) for SERCA1 (Hs01092295_m1), SERCA2 (Hs00544877_m1), RyR1 (Hs00166991_m1), α1s-DHPR (Hs00163885_m1), myogenin (Hs01072232_m1), MYH7 (Hs01110632_m1), MYH2 (HsHs0040042_m1), MYH1 (Hs00428600_m1), UBB (Hs00430290_m1), UBC (Hs00824723_m1), RPS18 (Hs01375212_g1), GAPDH (4352934E), 18S (4310893E), CTSB (Hs00157194_m1), and HK2 (Hs00606086_m1) were purchased from Applied Biosystems, Life Technologies. Reaction and amplification mixes (10 μl) consisted of the diluted (1:50) cDNA (4.5 μl), TaqMan Fast Universal PCR Master Mix (5.0 μl), and specific primers (0.5 μl). Subsequent cycling protocols were 2 min at 50 °C and 10 min at 90 °C followed by 40 cycles at 95 °C for 15 s and 60 °C for 1 min. To identify the optimal reference gene for normalization, the stability of the following reference genes was analyzed: UBB, UBC, RPS18, GAPDH, 18S, CTSB, and HK2. The geometrical mean of RPS18 and UBB was identified as the most stable reference according to the NormFinder algorithm , and target gene expression was therefore reported as a ratio to the geometrical mean of these two by the 2−ΔCT formula.
Protein extraction and Western blot
Cells and muscle bundles from intercostal muscle biopsies were homogenized in lysis buffer of the following composition (mM): 20 HEPES (pH 7.6), 150 NaCl, 5 EDTA, 1 Na3VO4 and 25 KF, 5 % glycerol (v/v), 0.5 % Triton X-100 (v/v), and protease inhibitor cocktail (one tablet/50 mL; Roche Diagnostics GmbH, Mannheim, Germany). Protein concentrations were subsequently determined using the Bradford technique. 50–60 μg of protein for myoblasts and myotubes or 15 μg per sample for intercostal muscles were loaded on 4–20 % SDS precast gels (Bio-Rad, Hercules, CA, USA) and separated through electrophoresis together with a protein ladder. Gels were transferred to PVDF-membranes using the Trans-Blot Turbo Transfer System from Bio-Rad. Blocking was completed using fluorescent blocking buffer (Merck Millipore, Darmstadt, Germany) during 60 min at room temperature (RT). Membranes were incubated overnight at 4 °C with primary antibodies (1:1000) for SERCA1 (ab2819, Abcam, Cambridge, FL, USA), SERCA2 (ab2861, Abcam), RyR (ab2868, Abcam) and/or (1:2000) for α1s-DHPR (sc-8160, Santa Cruz Biotechnology, Dallas, TX, USA). After the overnight incubation, membranes were washed (3 × 10 min) in PBST (0.1 %) and incubated with IRDye secondary antibody (LI-COR Biosciences, Cambridge, UK) for 60 min at RT. A final series of washes were then performed before scanning the membranes (Odyssey SA Infrared Imaging System, LI-COR Bioscience). The blots were subsequently quantified using ImageJ. Protein quantity was expressed as a ratio to total GAPDH abundance (1:2000; ab9485, Abcam).
Muscle fiber bundles containing four to five fibers were dissected manually and mounted in 35-mm petri dishes (P35G-0-14-C, MatTek). Myotubes and muscle fibers were fixed in 4 % formaldehyde for 20 min and then permeabilized by 0.3 % Triton X-100 in PBS solution. After rinsing, fibers were pre-incubated for 30 min in 4 % bovine serum albumin (BSA) (Sigma-Aldrich). Myotubes and muscle fibers were incubated overnight at 4 °C with primary antibodies (1:50) for SERCA1 (ab2819, Abcam), SERCA2 (ab2861, Abcam), RyR (MA3-925 (34C), Thermo Scientific, Waltham, MA, USA), α1s-DHPR (sc-8160, Santa Cruz) (dilution in PBS containing 1 % BSA and 0.1 % Triton-X) at 4 °C overnight. After the overnight incubation, myotubes and muscle fibers were washed (3 × 10 min) in PBS and incubated with secondary antibody (1:500) anti-mouse Alexa Fluor 568 and/or anti-goat Alexa Fluor 488 (Invitrogen, Life Technologies). Images of longitudinal thin sections of stained cells were obtained with laser confocal microscopy using a modified Bio-Rad MRC 1024 unit attached to a Nikon Diaphot inverted microscope with a Nikon Plan Apo 60X oil objective (NA 0.75). Excitation was at 491 and 561 nm, and the emitted light was collected through 522- and 605-nm narrowband filters. To provide an estimate of the overlap between RyR and α1s-DHPR signals in myotubes and intercostal fibers, the intensity plot of RyR and α1s-DHPR labeling was measured in ImageJ.
Transmission electron microscopy
Human myotubes and intercostal muscle fibers were fixed in 2.5 % glutaraldehyde in 0.1 M phosphate buffer, pH 7.4. After fixation, cells were rinsed in 0.1 M phosphate buffer and centrifuged. The pellets were then post-fixed in 2 % osmium tetroxide in 0.1 M phosphate buffer, pH 7.4 at 4 °C for 2 h, dehydrated in ethanol followed by acetone and embedded in LX-112 (Ladd, Burlington, VT, USA). Ultrathin sections (approximately 50–60 nm) were cut by a Leica EM UC 6 (Leica, Vienna, Austria). Sections were contrasted with uranyl acetate followed by lead citrate and examined in a Tecnai 12 Spirit Bio TWIN transmission electron microscope (FEI Company, Eindhoven, Netherlands) 100 kV. Digital images were taken by using a Veleta camera (Olympus Soft Imaging Solutions, GmbH, Münster, Germany).
SigmaPlot version 13.0 (Systat Software, Inc., San Jose, CA, USA) was used for statistical analysis. All variables were examined for normal distribution and were log- or square root-transformed before analysis as needed to better approximate a normal distribution. For comparisons between two data groups, two-tailed Student’s t test or Mann-Whitney Rank Sum test were used as appropriate. Statistically significant differences when comparing more than two groups were evaluated by the use of a one-way ANOVA. Tukey’s post hoc test was used to locate differences in mean values. Differences were considered significant at P < 0.05. Data are represented as means (± SEM) unless stated otherwise.
Human muscle fibers and myotubes show marked differences in [Ca2+]i transient kinetics and ability to contract
In intact muscle fibers, electrically evoked action potentials activate the voltage sensitive Ca2+ channels (DHPR) in the t-tubules that in turn triggers the opening of the sarcoplasmic reticulum (SR) Ca2+ release channel (RyR1), resulting in an increase in [Ca2+]i and force production. In this way, the events following α-motoneuron activity in vivo can be recapitulated ex vivo . We tested whether electrically stimulated human myotubes and myoblasts would also respond with an increase in [Ca2+]i and compared the resultant [Ca2+]i transients to those in human muscle fibers.
As the [Ca2+]i transient duration has been demonstrated to be of importance for the downstream effects of Ca2+ [22, 21], the Ca2+-handling properties of human muscle fibers and myotubes were further investigated regarding the [Ca2+]i transient kinetics. Aligned [Ca2+]i transients demonstrated similar Ca2+ release kinetics between human muscle fibers and myotubes (Fig. 1a). However, the decay of the Ca2+ transients was markedly slower in human myotubes than in the muscle fibers (Fig. 1a). By fitting a single exponential function plus a constant to the [Ca2+]i transient decay phase, the mean Ca2+ decay time constants were calculated. The mean Ca2+ decay time constant was approximately six times greater (888 ms (±66) vs. 146 ms (±13), P = 0.002) in myotubes than in muscle fibers.
The abundance of Ca2+-handling proteins differs markedly between human muscle fibers, myoblasts, and myotubes
Differences in the relative abundance of Ca2+-handling proteins will affect the functional Ca2+-handling properties of muscle cells . Thus, we hypothesized that the differences in functional Ca2+-handling observed between human muscle fibers, myoblasts, and myotubes would be explained by differences in the quantity of Ca2+-handling proteins. The mRNA and protein levels of RyR, DHPR, and the SR Ca2+ ATPases SERCA1 and SERCA2 were therefore analyzed in these cells.
In muscle fibers, the protein quantity of RyR when normalized to GAPDH was approximately 17-fold higher (P < 0.001) and that of DHPR approximately fourfold higher (P = 0.039) than in myotubes (Fig. 3e). RyR protein was non-detectable in myoblasts, and DHPR only weakly detected in myoblasts upon deliberate overexposure of the membrane (typical example in Fig. 3c). While readily detected in muscle fibers, SERCA1 protein was non-detectable by Western blot in either myotubes or myoblasts even as the membrane was deliberately overexposed (typical example in Fig. 3d). The protein quantity of SERCA2 when normalized to GAPDH was approximately 28-fold higher in muscle fibers (P = 0.003) than in myotubes (Fig. 3f) and only weakly detected in myoblasts upon deliberate overexposure of the membrane (typical example in Fig. 3d).
Finally, we investigated the mRNA abundance of the slow type skeletal muscle/β-cardiac myosin heavy chain (MHC I), type IIa MHC (MHC IIa), and type IIx MHC (MHC IIx) in human muscle fibers, myotubes, and myoblasts. In muscle fibers, the mRNA quantity of MHC I (P = 0.004 and P < 0.001, respectively), MHC IIa (P < 0.001) and MHC IIx (P < 0.001) was higher than in both myotubes and myoblasts (Additional file 4: Figure S3A). Following differentiation of myoblasts into myotubes, there was an increase (P < 0.001) in the mRNA quantity of all MHC isoforms investigated from very low levels of expression in myoblasts (Additional file 4: Figure S3A). In both muscle fibers and myotubes, MHC IIa was the predominantly expressed MHC isoform at the mRNA level (Additional file 4: Figure S3B).
The localization of Ca2+-handling proteins and ultrastructure differ markedly between human muscle fibers and myotubes
Myoblast differentiation into myotubes resulted in increased levels of Ca2+-handling proteins (Fig. 3a–d). One further step in the maturation process is the alignment of proteins into the arrangement required for fully functional Ca2+-handling in adult muscle fibers. To elucidate potential differences between human muscle fibers and myotubes in this regard, we used immunofluorescence staining and confocal microscopy to compare the localization of Ca2+-handling proteins. Additionally, transmission electron microscopy (TEM) was used to identify ultrastructural differences between human muscle fibers and myotubes.
Human myoblast differentiation into myotubes is paralleled by functional changes in intracellular Ca2+-handling
Myoblast differentiation into myotubes is associated with changes towards attaining morphological, metabolic, and biochemical properties similar to those in adult skeletal muscle [3, 4]. In addition, human myotubes but not myoblasts, respond to electrical stimulation by an increase in [Ca2+]i (Fig. 1a), and myoblast differentiation into myotubes results in increased levels of Ca2+-handling proteins (Fig. 3a–d). As this indicates that differences in terms of intracellular Ca2+-handling exist between myoblasts and myotubes, we compared the functional Ca2+-handling properties of these cells.
In the current study, we demonstrate the first measurements of [Ca2+]i and force production in intact single human muscle fibers. This novel model was employed to compare the Ca2+-handling properties of human muscle fibers with that of an existing ex vivo model of human muscle constituted by human myotubes. The major novel findings of the current study are the following: (1) intact single muscle fibers that allow for continuous measurements of [Ca2+]i and force production can be obtained from human intercostal muscle biopsies, (2) marked differences in intracellular Ca2+-handling exist between human muscle fibers and myotubes, and (3) these result from differences in the quantity and localization of Ca2+-handling proteins.
The basal and tetanic [Ca2+]i together with specific forces reported here in human muscle fibers are similar to values reported previously in intact single rodent muscle fibers [24, 34]. However, the specific forces generated by the intact single human muscle fibers are somewhat larger than what generally has been reported for skinned human muscle fibers [35–38]. As skinned intercostal muscle fibers generate forces similar to those reported in skinned muscle fibers obtained from other muscle groups , disparities between muscle groups are unlikely to explain this difference. Instead, this is likely explained by other factors that distinguish the two preparations, such as the loss of soluble proteins or altered spacing of myosin and actin filaments associated with the skinning procedure . This discrepancy underlines the importance of studying basic properties of human muscle in intact viable muscle fibers in addition to previously established models. Dissection of single human muscle fibers is technically challenging and requires human muscle biopsies with tendons intact at both ends. The current model is therefore unlikely to replace existing methods of clinical screening for diseased Ca2+-handling, such as malignant hyperthermia, but will provide a powerful tool for studies of Ca2+-dependent processes in human adult muscle. Moreover, the current model illustrates that mechanistic studies of the sort previously limited to animal models can be implemented also for studies of human skeletal muscle.
In adult skeletal muscle fibers, there is strictly controlled action potential-induced activation of the DHPR that, in turn, opens up the RyR1 to release Ca2+ from the SR [40, 41]. In the present study, approximately half of the investigated myotubes responded to electrical stimulation with an increase in [Ca2+]i. This is consistent with previous studies in human myotubes reporting depolarization-induced increases in [Ca2+]i in approximately one third  to one half of myotubes examined . These findings, together with the low protein expression and poor co-localization of DHPR and RyR found in the current study and by others [15, 17], show that the Ca2+-handling is functionally immature in human myotubes. Furthermore, the electron microscope images reveal that the ultrastructure, including the association between t-tubules and SR, is completely different in human muscle fibers and myotubes.
Further evidence of an immature Ca2+-handling in myotubes is the markedly slower decay of [Ca2+]i transients in myotubes than in adult muscle fibers. This finding is in accordance with previous reports of a slow restoration of [Ca2+]i to resting levels in immortalized human myotubes following depolarization . These authors suggested that this was due to low levels of SERCA1 expression and our results confirm that there is little SERCA1 and SERCA2 at either the mRNA or protein level in myotubes. In addition, we demonstrate that the structured organization of SERCA1 and SERCA2 in muscle fibers is absent in myotubes. Collectively, the differences in protein quantity and localization of SERCA1 and SERCA2 between muscle fibers and myotubes provide an explanation to the observed difference in [Ca2+]i transient decay. In addition, these differences may help explain the lower stimulation frequencies necessary to attain a fused tetanus in contracting human myotubes in comparison with muscle fibers [12–14]. Importantly, as diverse durations of [Ca2+]i transients differentially affect the cellular signaling initiated by increased [Ca2+]i [21, 22, 42–45], distinct rates of [Ca2+]i transient decay in myotubes and muscle fibers, as demonstrated in the current study, will result in divergent effects on Ca2+ signaling pathways. Thus, when studying Ca2+-dependent processes such as gene expression and metabolism, myotubes are likely to provide different answers to those given by intact adult muscle fibers.
In the present myotubes, contractions were not observed when [Ca2+]i was increased either by current pulses or 4-CmC application. This was explained by a low expression of sarcomeric MHCs and a lack of sarcomeric structures in human myotubes when analyzed by electron microscopy. The inability of human myotubes to contract is in accordance with the majority of previous studies and indicates a difference to rodent myotubes, which may contract when grown in culture [15–17]. Alternative culture protocols have attempted to grow myotubes that contract, including co-cultures with fetal rat spinal cord explants or neural agrin [15, 16], implementation of continuous electrical pulse stimulations into the culture protocol [5, 46], or bioengineering techniques . These approaches have been partially successful in growing contracting human myotubes ex vivo [5, 12, 13, 46–48] and highlight the difficulty of growing human myotubes with characteristics similar to those of muscle fibers. Importantly, even though contracting human myotubes may be obtained in culture, they produce less contractile force, attain fused tetanus at lower stimulation frequencies, and exhibit kinetic parameters that are slower than those in muscle fibers [12–14].
While the Ca2+-handling is less mature in human myotubes than in muscle fibers, our results also demonstrate substantial differences regarding Ca2+-handling properties between human myoblasts and myotubes. Changes in responsiveness to ATP and 4-CmC as well as an induced expression of Ca2+-handling proteins were observed as myoblasts were differentiated into myotubes. The purinergic receptor agonist ATP has previously been demonstrated to increase [Ca2+]i in human myoblasts and in the early process of myotube differentiation, with a reduced responsiveness to ATP in later developmental stages of myotube differentiation . Conversely, 4-CmC has been demonstrated to be a specific activator of the SR Ca2+ release channel RyR1  and would be expected to increase [Ca2+]i in myogenic cells with a functional SR. The current findings of increased [Ca2+]i upon stimulation with 4-CmC in myotubes therefore indicates a maturation of intracellular Ca2+-handling towards that in adult muscle fibers. Moreover, the present results suggest that myoblasts and myotubes are functionally distinct regarding their Ca2+-handling properties and can reliably be distinguished by their divergent responses to ATP and 4-CmC. This is in line with previous reports of other properties acquired following differentiation of myoblasts into myotubes that are similar to those in adult muscle [3, 4].
Intact single muscle fibers that display strictly regulated [Ca2+]i transients and force production upon electrical stimulation can be obtained by careful dissection of human intercostal muscle biopsies. In contrast, despite a more mature Ca2+-handling in myotubes than in myoblasts, human myotubes lack important aspects of adult Ca2+-handling. We conclude that human myotubes are an inappropriate model for human adult muscle when studying Ca2+-dependent processes, such as gene expression and metabolic processes. These results provide insights to some important functional differences between adult muscle fibers and myotubes and suggest caution when translating results obtained in human myotubes to muscle fibers.
- [Ca2+]i :
cytosolic free [Ca2+]
18S ribosomal RNA
bovine serum albumin
Dulbecco’s modified eagle medium
fetal bovine serum
magnetic activated cell sorting
- MHC I:
slow type skeletal muscle/β-cardiac myosin heavy chain
- MHC IIa:
type IIa myosin heavy chain
- MHC IIx:
type IIx myosin heavy chain
myosin heavy chain
phosphate buffered saline
ribosomal protein S18
sarcoplasmic/endoplasmic reticulum Ca2+ ATPase
transmission electron microscopy
The authors would like to acknowledge Dr. Tommy Lundberg for valuable input on the manuscript. This work was supported by funding from the Swedish Research Council, the Swedish National Centre for Research in Sports, the Swedish Heart-Lung Foundation and the Marianne and Marcus Wallenberg Foundation. KO is supported by a CSTP fellowship from Karolinska Institutet.
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