This work was supported by the Swiss National Science Foundation (grant number 310030-166313), The "Fondation Suisse pour la recherche sur les maladies musculaires," the "Foundation Marcel Levaillant," and the "Fondation pour la recherche ost&#233;o-articulaire."

Human muscle samples (tissues obtained from semitendinous muscles) were collected during orthopedic surgery on healthy patients as surgical waste. All methods relating to the human study were performed in accordance with the guidelines and regulations of the Swiss Regulatory Health Authorities and approved by the Commission Cantonale d&#39;Ethique de la Recherche from the Geneva Cantonal Authorities, Switzerland (protocol CER n&#176; 12-259). Informed and written consents were obtained from all adult subjects involved in the study.
1. Isolation of Myoblasts from Human Skeletal Muscle
<ol>
	<li>During all procedures, maintain sterile conditions by working under a level II tissue culture hood and using sterile medium and sterile buffers. 
	Note: This procedure can be applied to many human skeletal muscles.</li>
</ol>
2. Dissociation Protocol
<ol>
	<li>Transfer muscle samples (2 - 3 g) into a sterile 6 cm plate.</li>
	<li>Wash the muscle samples with 5 - 10 mL of phosphate buffered saline (PBS). Repeat the wash.</li>
	<li>Remove connective and adipose tissues using curved scissors and a clamp.</li>
	<li>Weigh the muscle sample (see step 2.13) and transfer it to a new sterile 6 cm plate.</li>
	<li>Add 5 mL of 0.05% Trypsin-EDTA.</li>
	<li>Using scissors, cut and mince the muscle tissue into small pieces (less than 2 mm).</li>
	<li>Using a 10 mL serological pipette, transfer the minced muscle to a 200 mL dissociation bottle containing a magnetic bar. Repeat this step until 90 mL of 0.05%Trypsin-EDTA has been transferred with all minced muscle to the dissociation box. Close the dissociation bottle and incubate at 37 &#176;C for 60 min under mild agitation.</li>
	<li>Remove the dissociation bottle from the heating bath and stop the enzymatic reaction by adding 10 mL of fetal calf serum (FCS; 10% final volume) to the dissociation bottle. Homogenize the mixture with a pipette.</li>
	<li>Prepare two 50 mL tubes with a 70 &#181;m cell strainer on each. Pass half of the cell suspension through the cell strainer on each tube by pipetting up and down on the filter until the suspension passes through.</li>
	<li>Centrifuge the tubes at 200 x g and 20 &#176;C for 5 min; aspirate and discard the supernatant.</li>
	<li>Resuspend the cells with 10 mL of growth medium (GM; Table 1) and pass the cells through a 40-&#181;m cell strainer placed on one 50 mL tube. Centrifuge the tube at 200 x g and 20 &#176;C for 5 min; aspirate and discard the supernatant.</li>
	<li>Resuspend the cells with 10 mL of GM and centrifuge the tube at 200 x g and 20 &#176;C for 5 min; aspirate and discard the supernatant.</li>
	<li>Resuspend the cells with 1 mL of GM and count the cells using a Neubauer chamber. 
	Note: Usually, a yield of 3 x 105 cells per g of muscle should be harvested. The yield is calculated by dividing the number of cells by the weight of the muscle biopsy (step 2.4).</li>
	<li>Put 2 x 105 cells in a sterile 6-cm plate containing 4 mL of GM and incubate at 37 &#176;C and 7% CO2. Prepare as many plates as necessary.</li>
	<li>Change the GM after 3 days. 
	Note: Freshly dissociated cells need time to adhere to the culture plate. Moreover, the first myogenic cell doubling time is around 50 - 60 h. After the first division, the doubling time is around 20 h.</li>
	<li>After 5 - 7 days, when cells reach 70 - 80% confluence (around 1.5 x 106 cells/per 6 cm plate), start antibody staining and cell sorting (step 3).</li>
</ol>
3. Antibody Staining and Cell Sorting
<ol>
	<li>After rinsing once with PBS, incubate the adherent cells (in the 6-cm plate) with 1 mL of 0.05% Trypsin-EDTA at 37 &#176;C and 7% CO2 for 3 min (or incubate at 5% CO2). Stop the reaction by adding 2 mL of GM and collect the cells in 15 mL tube. After centrifugation (200 x g for 5 min), resuspend the cells in 1 mL of GM.</li>
	<li>Count the cells using a Neubauer chamber.</li>
	<li>Perform the following procedures on ice. 
	Note: The procedure for setting the correct fluorescence compensation on the cytometer is essential and should be similar on any cytometer. Tubes 1 and 2 are used as negative controls. Tubes 3 - 7 are used to set the compensation on the cytometer during the first experiment (Table 2). The final concentration used for each antibody and its corresponding isotype should be the same (around 1 &#181;g/106 cells).</li>
	<li>Dispatch 1 x 105 cells per tube (5-mL tube) for tubes 1 - 7 (Table 2). Place the remaining cell suspension in tube 8 for cell sorting.</li>
	<li>Wash the cells with 1 mL of cold FACS buffer (PBS-5% FCS). Close and centrifuge the tubes at 200 x g and 4 &#176;C for 5 min; aspirate and discard the supernatant.</li>
	<li>Add 200 &#181;L of FACS buffer per tube. Add antibodies to the tube according to Table 2. Incubate on ice for 30 min.</li>
	<li>Wash the cells: add 1 mL of the FACS buffer to each tube. Close and centrifuge the tubes at 200 x g and 4 &#176;C for 5 min. Aspirate and discard the supernatant. Repeat this step one more time.</li>
	<li>Resuspend the cells in 500 &#181;L of FACS buffer. Close the tubes and keep them on ice until cell sorting. Prepare 4 tubes of 1.5 mL, each containing 500 &#181;L of GM, which is necessary to collect the cells during cell sorting. Keep them on ice.</li>
	<li>Sort human myoblasts by flow cytometry (Figure 1). After excluding CD45-positive hematopoietic cells, separate CD45-negative cells based on CD56 expression. Then, gate the CD56-positive population for CD34, CD144, and CD146 (human myoblasts are defined as CD56+/CD146+/CD45-/CD34-/CD144- cells). Reanalyze a small fraction of the sorted cell population to check the purity.</li>
	<li>Pool the tubes containing human myoblasts in one 15-mL tube and wash one time by adding 5 mL of GM. Centrifuge the tube at 200 x g and 20 &#176;C for 5 min; aspirate and discard the supernatant.</li>
	<li>Resuspend the cells in 1 mL of GM and plate the cells on an uncoated 6-cm plate containing 4 mL of GM (2 x 105 cells per plate). Incubate at 37 &#176;C and 7% CO2.</li>
</ol>
4. Cell Maintenance
<ol>
	<li>Change half of the culture medium every 2 days. Remove 2 mL of old GM from the culture dish and add 2 mL of GM. 
	Note: Myoblasts produce growth factors that are beneficial for culture conditions.</li>
	<li>Trypsinize the cells when they reach 70 - 80% confluence to avoid pre-differentiation. After rinsing once with PBS, incubate the adherent cells (in the 6-cm plate) with 1 mL of 0.05% Trypsin-EDTA at 37 &#176;C and 7% CO2 for 3 min. Stop the reaction by adding 2 mL of GM and collect the cells in a 15 mL tube. After centrifugation (200 x g for 5 min), resuspend the cells in 1 mL of GM.</li>
	<li>Count the cells and prepare 6 cm plates with 2 x 105 cells per plate in 4 mL of GM.</li>
	<li>Keep human myoblasts up to 6 passages or freeze them in GM containing 10% DMSO, 1 x 106 cells per mL. Freeze the cells in a progressive freezing box (1 &#176;C/min); for long-term storage, place the cells in liquid nitrogen.</li>
</ol>
5. Transfection of Human Myoblasts with siRNA
Note: Myoblasts are transfected using commercial transfection reagent two days before the induction of the differentiation. The effect of specific siRNA against a gene is always compared to the effect of a control siRNA. siRNAs are double-stranded RNAs that must be kept on ice and manipulated with gloves.
<ol>
	<li>Obtain a 70 - 80% confluent monolayer of human myoblasts in a 6-cm plate (around 1.5 x 106 cells). 
	Note: 5 x 105 cells per transfection are needed to obtain a confluent monolayer of transfected myoblasts after 2 days in GM.</li>
	<li>Prepare glass coverslips (described for the calcium imaging in step 7).</li>
	<li>Warm GM, 0.05% Trypsin-EDTA, and PBS to 37 &#176;C.</li>
	<li>Perform all the following steps in a level II tissue culture hood to maintain sterility.</li>
	<li>In a 1.5-mL tube, dispatch 500 &#181;L of reduced serum medium, 3 &#181;L of transfection reagent, and 1 &#181;L of siRNA at 20 &#181;M. Mix gently and keep at room temperature (RT) for 15 - 20 min.</li>
	<li>During this incubation time, prepare the myoblasts for transfection. After rinsing once with PBS, incubate the adherent cells (in the 6-cm plate) with 1 mL of 0.05% Trypsin-EDTA at 37 &#176;C and 7% CO2 for 3 min. Stop the reaction by adding 2 mL of GM and collect the cells in a 15 mL tube. After centrifugation (200 x g for 5 min), resuspend the cells in 1 mL of GM.</li>
	<li>Count the cells using a Neubauer chamber.</li>
	<li>Resuspend 5 x 105 cells in 200 &#181;L of GM for each transfection (i.e., for 2 transfections, resuspend in 400 &#181;L).</li>
	<li>Add 200 &#181;L of the cell suspension to each 500 &#181;L of siRNA-transfectant mixture. Pipette up and down twice and incubate for 5 min at RT.</li>
	<li>Add the 700 &#181;L (cells + siRNA-transfectant) to a 3.5 cm culture dish containing a glass coverslip. Add GM to get a final volume of 2 mL.</li>
	<li>After 24 h at 37 &#176;C and 7% CO2, completely replace the medium with 2 mL of fresh GM. After another 24 h, a confluent monolayer of transfected cells is usually present; if so, induce myoblast differentiation by replacing the GM with 2 mL of differentiation medium (DM; Table 1).</li>
	<li>Perform calcium imaging experiments 24 - 48 h after the induction of differentiation or on proliferating myoblasts. 
	Note: Immunofluorescence is usually performed 48 h after the induction of differentiation.</li>
</ol>
6. Immunofluorescence Staining of Differentiation Markers
Note: Immunofluorescence staining is performed after 48 h in DM, but it can also be performed at other differentiation times. Myogenin and MEF2 are nuclear proteins only expressed in differentiated cells (before fusion into myotubes), and their staining allows for the assessment of the differentiation process. Myosin heavy chain (MyHC) is only expressed in myotubes and is used to estimate the myotube size and fusion index (number of nuclei in myotubes/total number of nuclei).
<ol>
	<li>After 48 h in DM, wash the confluent monolayer of differentiated cells twice with 1 mL of PBS at RT. Do this carefully to avoid detaching the myotubes.</li>
	<li>Fix the cells with 1 mL of 4% paraformaldehyde (PFA) for 15 min at RT. Wash the cells with PBS twice, for 5 min each.</li>
	<li>Permeabilize the cells with 1 mL of PBS containing 0.25% Triton X-100 for 45 min and wash 3 times with 1 mL of PBS.</li>
	<li>Block unspecific binding sites by incubating the cells in PBS supplemented with 0.2% Tween-10 and 2% goat serum (block solution) for 30 min at RT.</li>
	<li>Prepare primary antibodies diluted in block solution (600 &#181;L per 3.5-cm plate). 
	Note: The following primary antibodies are used: mouse anti-myogenin (1:600), rabbit anti-MEF2 (1:300), and mouse anti-MyHC antibody (MF20, 1:1,000). It is possible to mix in the same blocking solution, mouse anti-myosin heavy chain or mouse anti-myogenin, with rabbit anti-MEF2.</li>
	<li>Remove the block solution and add 600 &#181;L of primary antibody solutions per 3.5-cm plate and incubate overnight at 4 &#176;C in a humidified chamber.</li>
	<li>Wash the cells 3 times with 1 mL of PBS. Incubate for 1 h at RT in a dark chamber with 600 &#181;L of the secondary antibodies prepared in the block solution, as follows: goat anti-mouse Alexa 488 (1:1,000), goat anti-rabbit Alexa 546 (1:1,000), and DAPI (1:50, stock solution at 15 &#181;M). 
	CAUTION: DAPI is a toxic compound that should be handle with gloves.</li>
	<li>Aspirate and wash the cells 3 times with 1 mL of PBS.</li>
	<li>Mount glass coverslips using polyvinyl alcohol mounting medium with antifading solution. Keep them at 4 &#176;C and protected from light until imaging the cells.</li>
</ol>
7. Calcium Imaging
<ol>
	<li>Prior culture, prepare glass coverslips 25 mm in diameter. Put the coverslips in 70% ethanol for 48 h to remove any grease and dirt from the surface, which reduces cell adherence to the coverslip.</li>
	<li>After rinsing the coverslipsseveral times with water, put them back in water and sterilize them (autoclaved).</li>
	<li>Put one coverslip per 3.5-cm culture plate and let them dry. Store the Petri dishes for further experiments or use them immediately.</li>
	<li>Add the transfected myoblasts (5 - 6 x 105 cells) to the glass coverslip. 
	Note: Experiments are done 24 - 48 h after the induction of differentiation, or on proliferating myoblasts.</li>
	<li>To perform Ca2+ imaging, wash the cells 2 times with PBS or directly with the medium used for the experiments (calcium-containing medium; Table 3). Load the cells with 2 &#181;M Fura-2/AM plus 1 &#181;M pluronic acid for about 30 min in the dark at RT. 
	Note: Pluronic acid helps to solubilize Fura-2. Fura-2/AM and pluronic acid are dissolved in the calcium-containing medium.</li>
	<li>Wash the cells twice in the same medium but without Fura-2/AM and pluronic acid and leave them in the dark for an additional 10 - 15 min to allow for the de-esterification of Fura-2/AM.</li>
	<li>If the loading efficiency of Fura-2 is too low, add more Fura-2 (e.g., 4 &#181;M) and/or incubate the cells for a longer time (e.g., 40 min).</li>
	<li>Remove the coverslip from the culture plate and install it in the experimental chamber. Add ~1 mL of calcium-containing medium (depending upon the volume of the experimental chamber). Install it on the microscope stage and start the recording. 
	Note: The fluorescent Ca2+-sensitive probe Fura-2 is excited alternately at 340 nm and 380 nm, and the emission light is collected at 510 nm. As the fluctuations of the Ca2+ concentration are relatively slow, acquire 1 ratio every 2 s.</li>
	<li>To activate SOCE, the following classical thapsigargin/Ca2+ re-addition protocol is used: leave the cells for 2-3 min in calcium-containing medium. Replace the medium with calcium-free medium for 1-2 min.
	<ol>
		<li>Add 1 &#181;M thapsigargin to deplete the internal calcium stores. 
		Note: Thapsigargin is an irreversible blocker of sarco-endoplasmic reticulum Ca2+ ATPase (SERCA) pump activity.After 8 - 10 min, rapidly replace the medium with the calcium-containing medium. Ca2+ will enter the cells through the SOCE channels. Wait around 5 min before terminating the experiments. 
		CAUTION: Thapsigargin is a toxic compound that should be handled with gloves.</li>
	</ol>
	</li>
</ol>
8. Ca2+ Measurement Analysis
Note: Perform the analysis with the acquisition software.
<ol>
	<li>Open the experiment. Draw region 1 in an area where there are no cells (background region), and draw the other regions in the cytoplasm of the cells to be analyzed (the regions can be drawn on any image: the 340-nm, the 380-nm, or the ratio image). Go to &#34;Run experiments&#34; &#62; &#34;reference images.&#34; Under the 340 and 380 channels, select &#34;region 1&#34; and then click on &#34;Subtract background.&#34;</li>
	<li>Run the entire experiment (F4: forward) to ensure that nothing interferes with the background region, such as a fluorescent dust spot that travels through the background region, and that the cells are not moving during the time of the experiments. 
	Note: The regions of interest that are selected should not include dark areas; otherwise, the ratio value will appear noisy, as part of the background will be included in the measurement.</li>
	<li>Click on &#34;Log data&#34; and play the entire experiment again. 
	Note: This will open a spreadsheet in which the time and the ratio are logged. It is also possible to save the individual 340 nm and 380 nm wavelengths.</li>
	<li>To get information about SOCE, extract the two important parameters from the experiment: the amplitude of the Ca2+ re-addition and the slope of the Ca2+ entry phase (Figure 3).</li>
	<li>Obtain the amplitude by subtracting the ratio value just before Ca2+ re-addition from the maximal amplitude of the Ca2+ elevation. Obtain the maximal slope by fitting a linear regression of the first seconds after Ca2+ re-addition. 
	Note: The area under the curve of the thapsigargin response is also an informative parameter to extract, as it reflects the amount of free calcium stored in the SR.</li>
</ol>

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The authors have no competing financial interests.

The isolation and culture of human myoblasts from adult skeletal muscle offers an in vitro model to study muscle differentiation and muscle regeneration. In this paper, we provide a protocol that allows for the purification of high yields of human myoblasts in a simple and cost-limited way. In addition, this technique provides reliable and reproducible results in terms of percentage of myoblasts isolated and their myogenic differentiation efficiency. Indeed, we obtained around 60...

Human skeletal muscles are composed of groups of contractile, multinucleated muscle fibers resulting from the fusion of myogenic precursor cells. Skeletal muscles have the capacity to regenerate after injury thanks to the presence of SCs, the skeletal muscle stem cells located between the plasma membrane of myofibers (sarcolemma) and the basal lamina. In uninjured muscle, SCs are mostly present in a quiescent state. In response to mechanical stress or injury, SCs become activated (myoblasts), proliferate, and undergo either differentiation to form new myofibers or self-renewal to replenish the SC pool1,2. Over the years, several techniques were developed to isolate SCs and their progeny, the myoblasts, from skeletal muscle biopsies. With the greater understanding of the cell surface markers expressed on these cells and the methodology of fluorescence-activated cell sorting (FACS)3,4,5, it is now possible to isolate pure populations of human myoblasts from muscle samples.
Calcium signaling regulates skeletal muscle development, homeostasis, and regeneration. In particular, the Ca2+ entry that is activated following intracellular store depletion, called SOCE, is of great importance6 for these processes. Upon cell stimulation, the Ca2+ level in the endo/sarcoplasmic reticulum (ER/SR) decreases, which in turn activates plasma membrane Ca2+ channels that permit Ca2+ entry to refill the ER/SR7. The two main proteins involved in SOCE are the ER/SR Ca2+-sensing stromal interaction molecule 1 (STIM1) protein and the plasma membrane Ca2+ channel Orai1. Skeletal muscle abundantly expresses these two proteins, as well as other proteins of the same families (STIM2 and Orai2-Orai3)8,9 and several Ca2+-permeable channels of the transient receptor potential canonical (TRPC) family10,11,12,13. Ca2+ entry through the SOCE pathway is of great importance for muscle formation/regeneration14. Mutations of STIM1 or Orai1 proteins have deleterious effects on muscle function, leading mainly to muscle hypotonia15. Animal models with SOCE impairment also display reduced muscle mass and force, together with enhanced fatigability6,16,17. As mentioned, other STIM and Orai proteins, as well as many TRPC channels, are expressed in skeletal muscle, and their respective roles have not been determined so far. By knocking down their expression level, it is thus possible to investigate their implications in SOCE and their roles during human skeletal muscle differentiation.
SOCE measurement can be performed using two different approaches: electrophysiological current recordings and Ca2+ measurements. Electrophysiology is certainly a more direct method, as it measures the current of interest and its associated electrophysiological signature. However, this technique is very difficult to apply to muscle cells, mainly because of the large size of the muscle cells and the small size of the endogenous SOCE current. Cytosolic Ca2+ imaging is technically very accessible, but the measure is more indirect, as the Ca2+ level measured in the cytosol reflects both the entry of Ca2+ and the re-pumping out of the cell or in the internal stores.
A methodology that includes isolating myoblasts from small pieces of human skeletal muscle after enzymatic digestion, amplification, and FACS is explained in this paper. The process of muscle differentiation and the immunofluorescence protocol to follow the expression of differentiation markers over time are described18. Finally, the measurement of SOCE and the role of different ion channels in Ca2+ signaling and skeletal muscle differentiation are explained.

<table><tbody><tr><td>Ham&amp;rsquo;s F10</td><td>ThermoFisher Scientific</td><td>31550-023</td><td></td></tr><tr><td>FCS</td><td>ThermoFisher Scientific</td><td>10270-106</td><td>Lot 41G5532K</td></tr><tr><td>BSA</td><td>Sigma</td><td>O5482</td><td></td></tr><tr><td>fetuin</td><td>Sigma</td><td>F3385</td><td></td></tr><tr><td>EGF</td><td>Corning</td><td>354001</td><td></td></tr><tr><td>Dexamethansone</td><td>Sigma</td><td>D1756</td><td></td></tr><tr><td>Insulin</td><td>Sigma</td><td>I9278</td><td></td></tr><tr><td>Creatine</td><td>Sigma</td><td>27900</td><td></td></tr><tr><td>Pyruvate</td><td>Sigma</td><td>P4562</td><td></td></tr><tr><td>Uridine</td><td>Sigma</td><td>U3003</td><td></td></tr><tr><td>Gentamycin</td><td>ThermoFisher Scientific</td><td>15710-049</td><td></td></tr><tr><td>Trypsin-EDTA 0.05%</td><td>ThermoFisher Scientific</td><td>25300-062</td><td></td></tr><tr><td>70 &amp;mu;m cell strainer</td><td>Falcon</td><td>352350</td><td></td></tr><tr><td>40 &amp;mu;m cell strainer</td><td>Falcon</td><td>352340</td><td></td></tr><tr><td>Falcon 50&amp;nbsp;mL tube</td><td>Falcon</td><td>352070</td><td></td></tr><tr><td>Falcon 15&amp;nbsp;mL tube</td><td>Falcon</td><td>352096</td><td></td></tr><tr><td>Falcon 5&amp;nbsp;mL tube (FACS)</td><td>Falcon</td><td>352058</td><td></td></tr><tr><td>Culture medium: OPTI-MEM</td><td>ThermoFisher Scientific</td><td>31985-047</td><td></td></tr><tr><td>Transfectant reagnt: RNAiMax</td><td>ThermoFisher Scientific</td><td>1756064</td><td></td></tr><tr><td>PBS</td><td>ThermoFisher Scientific</td><td>14190-094</td><td></td></tr><tr><td>Mounting medium</td><td>Fluka</td><td>10981</td><td></td></tr><tr><td>Mouse anti-MyHC *</td><td>DSHB</td><td>MF20&amp;nbsp;</td><td>concentrate</td></tr><tr><td>Mouse anti-myogenin</td><td>BD Pharmigen</td><td>556358</td><td></td></tr><tr><td>Rabbit anti-MEF2</td><td>Santa Cruz Biotech</td><td>C-21</td><td></td></tr><tr><td>Goat anti-mouse Alexa488</td><td>ThermoFisher Scientific</td><td>A11029</td><td></td></tr><tr><td>Goat anti-rabbit Alexa 546</td><td>ThermoFisher Scientific</td><td>A11035</td><td></td></tr><tr><td>Mouse anti human CD56-Alexa488</td><td>BD Pharmingen</td><td>557699</td><td></td></tr><tr><td>Mouse anti human CD146-PECy7</td><td>BD Pharmingen</td><td>562135</td><td></td></tr><tr><td>Mouse anti human CD45-PE-CF594</td><td>BD Pharmingen</td><td>562279</td><td></td></tr><tr><td>Mouse anti human CD34-APC</td><td>BD Pharmingen</td><td>345804</td><td></td></tr><tr><td>Mouse anti human CD144-PE</td><td>BD Pharmingen</td><td>560410</td><td></td></tr><tr><td>Alexa Fluor 488 mouse IgG1k</td><td>BD Pharmingen</td><td>557702</td><td></td></tr><tr><td>PECy7 mouse IgG1k</td><td>BD Pharmingen</td><td>557872</td><td></td></tr><tr><td>PE-CF594 mouse IgG1k</td><td>BD Pharmingen</td><td>562292</td><td></td></tr><tr><td>APC mouse IgG1k</td><td>BD Pharmingen</td><td>550854</td><td></td></tr><tr><td>PE mouse IgG1k</td><td>BD Pharmingen</td><td>556650</td><td></td></tr><tr><td>DAPI</td><td>Sigma</td><td>D9542</td><td>CAUTION:&amp;nbsp; toxic compound</td></tr><tr><td>Paraformaldehyde</td><td>Fluka</td><td>762400</td><td></td></tr><tr><td>Fura-2 AM</td><td>ThermoFisher Scientific</td><td>F1201</td><td></td></tr><tr><td>Pluronic F-127</td><td>ThermoFisher Scientific</td><td>P3000MP</td><td></td></tr><tr><td>Thapsigargin</td><td>Sigma</td><td>T9033</td><td>CAUTION&amp;nbsp;: toxic compound&amp;nbsp;</td></tr><tr><td>Ratio imaging acquisition software: Metafluor 6.3</td><td>Molecular Device</td><td></td><td></td></tr><tr><td>*&amp;nbsp;: MF 20 was deposited to the DSHB by Fischman, D.A.&amp;nbsp;</td><td></td><td></td><td></td></tr></tbody></table>

isolation of human myoblasts, assessment of myogenic differentiation, and store-operated calcium entry measurement

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for muscle regeneration. Upon injury, which occurs frequently in skeletal muscles, SCs are activated. They proliferate as myoblasts and differentiate to repair muscle lesions. Among many events that take place during muscle differentiation, cytosolic Ca2+ signals are of great importance. These Ca2+ signals arise from Ca2+ release from internal Ca2+ stores, as well as from Ca2+ entry from the extracellular space, particularly the store-operated Ca2+ entry (SOCE). This paper describes a methodology used to obtain a pure population of human myoblasts from muscle samples collected after orthopedic surgery. The tissue is mechanically and enzymatically digested, and the cells are amplified and then sorted by flow cytometry according to the presence of specific membrane markers. Once obtained, human myoblasts are expanded and committed to differentiate by removing growth factors from the culture medium. The expression levels of specific transcription factors and in vitro immunofluorescence are used to assess the myogenic differentiation process in control conditions and after silencing proteins involved in Ca2+ signaling. Finally, we detail the use of Fura-2 as a ratiometric Ca2+ probe that provides reliable and reproducible measurements of SOCE.

After the enzymatic dissociation of a human muscle sample, cells were amplified in GM. Human myoblasts, defined as CD56+/CD146+/CD45-/CD34-/CD144- cells, were obtained after FACS. Myoblasts represented more than 60% of the analyzed population (Figure 1). Primary human myoblasts were replated, grown to confluence, and cultured in DM for 48 h. After immunostaining for the expression of the transcription factor MEF2 and the muscle-specific protein myosin heavy chain (MyHC), we observed that a m...

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Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for ...

isolation-human-myoblasts-assessment-myogenic-differentiation-store

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

10.0K Views.  Geneva University Hospitals & Faculty of Medicine. Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Video: Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated from human muscle biopsy samples using enzymatic digestion and their myogenic properties studied in culture. Quantitatively, the two main adherent cell types obtained from enzymatic digestion are: (i) the satellite cells (termed myogenic cells or muscle precursor cells), identified initially as CD56+ and later as CD56+/desmin+ cells and (ii) muscle-derived fibroblasts, identified as CD56&#8211; and TE-7+. Fibroblasts proliferate very efficiently in culture and in mixed cell populations these cells may overrun myogenic cells to dominate the culture. The isolation and purification of different cell types from human muscle is thus an important methodological consideration when trying to investigate the innate behavior of either cell type in culture. Here we describe a system of sorting based on the gentle enzymatic digestion of cells using collagenase and dispase followed by magnetic activated cell sorting (MACS) which gives both a high purity (&#62;95% myogenic cells) and good yield (~2.8 x 106 &#177; 8.87 x 105 cells/g tissue after 7 days in vitro) for experiments in culture. This approach is based on incubating the mixed muscle-derived cell population with magnetic microbeads beads conjugated to an antibody against CD56 and then passing cells though a magnetic field. CD56+ cells bound to microbeads are retained by the field whereas CD56&#8211; cells pass unimpeded through the column. Cell suspensions from any stage of the sorting process can be plated and cultured. Following a given intervention, cell morphology, and the expression and localization of proteins including nuclear transcription factors can be quantified using immunofluorescent labeling with specific antibodies and an image processing and analysis package.

The main adherent cell types derived from human muscle are myogenic cells and fibroblasts. Here, cell populations are enriched using magnetic-activated cell sorting based on the CD56 antigen. Subsequent immunolabelling with specific antibodies and use of image analysis techniques allows quantification of cytoplasmic and nuclear characteristics in individual cells.

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated ...

In Vitro Differentiation of Mature Myofibers for Live Imaging

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily conserved structures that compose it are assembled remains under investigation. Their size and physiological features often constrain manipulation and imaging applications. The culture of immortalized cell lines is widely used, but it can only replicate the early steps of differentiation.
Here, we describe a protocol that enables easy genetic manipulation of myofibers originating from primary mouse myoblasts. After one week of differentiation, the myofibers display contractility, aligned sarcomeres and triads, as well as peripheral nuclei. The entire differentiation process can be followed by live imaging or immunofluorescence. This system combines the advantages of the existing ex vivo and in vitro protocols. The possibility of easy and efficient transfection as well as the ease of access to all differentiation stages broadens the potential applications. Myofibers can subsequently be used not only to address relevant developmental and cell biology questions, but also to reproduce muscle disease phenotypes for clinical applications.

In Vitro Differentiation of Mature Myofibers for Live Imaging

Muscle cells are among the most complex eukaryotic cells. We present a protocol for the in vitro differentiation of highly mature myofibers that allows for genetic manipulation and clear imaging during all developmental stages.

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily ...

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to differentiate into later stages of muscle development. The protocol provided here describes a robust method for the isolation and culture of a highly proliferative population of myoblast cells from young adult mouse skeletal muscle explants. These cells are useful for the study of the metabolic properties of skeletal muscle of different mouse models, as well as in other downstream applications such as transfection with exogenous DNA or transduction with viral expression vectors. The level of differentiation and metabolic profile of these cells depends on the length of exposure, and composition of the media used to induce myoblast differentiation. These methods provide a robust system for the study of mouse muscle cell metabolism ex vivo. Importantly, unlike in vivo models, the methods described here provide a cell population that can be expanded and studied with high levels of reproducibility.

Myoblasts are proliferating precursor cells that differentiate to form polynucleated myotubes and eventually skeletal muscle myofibers. Here, we present a protocol for efficient isolation and culture of primary myoblasts from young adult mouse skeletal muscles. The method enables molecular, genetic, and metabolic studies of muscle cells in culture.

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Biology

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Functional Characterization of Endogenously Expressed Human RYR1 Variants

More than 700 variants in the RYR1 gene have been identified in patients with different neuromuscular disorders including malignant hyperthermia susceptibility, core myopathies and centronuclear myopathy. Because of the diverse phenotypes linked to RYR1 mutations it is fundamental to characterize their functional effects to classify variants carried by patients for future therapeutic interventions and identify non-pathogenic variants. Many laboratories have been interested in developing methods to functionally characterize RYR1 mutations expressed in patients&#39; cells. This approach has numerous advantages, including: mutations are endogenously expressed, RyR1 is not over-expressed, use of heterologous RyR1 expressing cells is avoided. However, since patients may present mutations in different genes aside RYR1, it is important to compare results from biological material from individuals harboring the same mutation, with different genetic backgrounds. The present manuscript describes methods developed to study the functional effects of endogenously expressed RYR1 variants in: (a) Epstein Barr virus immortalized human B-lymphocytes and (b) satellite cells derived from muscle biopsies and differentiated into myotubes. Changes in the intracellular calcium concentration triggered by the addition of a pharmacological RyR1 activators are then monitored. The selected cell type is loaded with a ratiometric fluorescent calcium indicator and intracellular [Ca2+] changes are monitored either at the single cell level by fluorescence microscopy or in cell populations using a spectrofluorometer. The resting [Ca2+], agonist dose response curves are then compared between cells from healthy controls and patients harboring RYR1 variants leading to insight into the functional effect of a given variant.

Here methods used to study the functional effect of RYR1 mutations endogenously expressed in Epstein Barr Virus immortalized human B-lymphocytes and muscle biopsy derived satellite cells differentiated into myotubes are described.

More than 700 variants in the RYR1 gene have been identified in patients with different neuromuscular disorders including malignant hyperthermia ...

Medicine

DNA Electroporation, Isolation and Imaging of Myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of serially linked sarcomeres, the smallest contractile units in muscle. Sarcomeric dysfunction contributes to muscle weakness in patients with mutations in genes encoding for sarcomeric proteins. The study of myofibril mechanics allows for the assessment of actin-myosin interactions without potential confounding effects of damaged, adjacent myofibrils when measuring the contractility of single muscle fibers. Ultrastructural damage and misalignment of myofibrils might contribute to impaired contractility. If structural damage is present in the myofibrils, they likely break during the isolation procedure or during the experiment. Furthermore, studies in myofibrils provide the assessment of actin-myosin interactions in the presence of the geometrical constraints of the sarcomeres. For instance, measurements in myofibrils can elucidate whether myofibrillar dysfunction is the primary effect of a mutation in a sarcomeric protein. In addition, perfusion with calcium solutions or compounds is almost instant due to the small diameter of the myofibril. This makes myofibrils eminently suitable to measure the rates of activation and relaxation during force production. The protocol described in this paper employs an optical force probe based on the principle of a Fabry-P&#233;rot interferometer capable of measuring forces in the nano-Newton range, coupled to a piezo length motor and a fast-step perfusion system. This setup enables the study of myofibril mechanics with high resolution force measurements.

Presented here is a protocol to assess the contractile properties of striated muscle myofibrils with nano-Newton resolution. The protocol employs a setup with an interferometry-based, optical force probe. This setup generates data with a high signal-to-noise ratio and enables the assessment of the contractile kinetics of myofibrils.

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of ...

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo

Adult skeletal muscle tissue harbors a stem cell population that is indispensable for its ability to regenerate. Upon muscle damage, muscle stem cells leave their quiescent state and activate the myogenic program ultimately leading to the repair of damaged tissue concomitant with the replenishment of the muscle stem cell pool. Various factors influence muscle stem cell activity, among them intrinsic stimuli but also signals from the direct muscle stem cell environment, the stem cell niche. The isolation and culture of single myofibers with their associated muscle stem cells preserves most of the interaction of the stem cell with its niche and is, therefore, the closest possibility to study muscle stem cell functionality ex vivo. Here, a protocol for the isolation, culture, siRNA transfection and immunostaining of muscle stem cells on their respective myofibers from mouse EDL (extensor digitorum longus) muscles is provided. The experimental conditions outlined here allow the study and manipulation of muscle stem cells ex vivo including investigation of myogenic activity without the inherent need for in vivo animal experiments.

In this protocol an in vitro culturing and functional analysis method for muscle stem cells is described, which preserves most of their interactions with their endogenous niche.

Adult skeletal muscle tissue harbors a stem cell population that is indispensable for its ability to regenerate. Upon muscle damage, muscle stem cells ...

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Summary

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Chapters in this video

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

In Vitro Differentiation of Mature Myofibers for Live Imaging

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Functional Characterization of Endogenously Expressed Human RYR1 Variants

DNA Electroporation, Isolation and Imaging of Myofibers

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca²⁺ Transients and L-Type Calcium Current

Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo