The authors are grateful to Dr. Matthew Watt at the University of Melbourne and Dr. Anastasia Kralli at Johns Hopkins University for assistance adopting this protocol based on the work of Mokbel et al.6. We also thank Dr. Sabine Jordan for assistance developing and adopting this protocol in our laboratory. This work was funded by the National Institutes of Health R01s DK097164 and DK112927 to K.A.L.

This protocol follows the animal care guidelines of Scripps Research.
1. Collection and Processing of Muscle Tissue Explants
<ol>
	<li>The day prior to dissection, sterilize all dissection equipment (forceps, razor blades, and scissors) and prepare all required media: phosphate buffered saline (PBS), MB Plating media (12.5 mL of DMEM, 12.5 mL of HAMS F12, 20 mL of heat-inactivated fetal bovine serum (FBS), 5 mL of amniotic fluid medium supplement), and Coating Solution (24 mL of DMEM, 24 mL of HAMS F12, 1.7 mL of collagen, 1 mL of matrigel).</li>
	<li>The day of dissection, coat one 6-cm dish with Coating Solution for each muscle to be dissected. Add 2 mL of Coating Solution to the surface of each plate, shake gently to create an even coat on the surface, and incubate the plates with solution at 4 &#176;C for 1 h.</li>
	<li>Remove Coating Solution from the plates and return to the stock solution. 
	NOTE: Coating Solution can be reused for up to six months and should be stored at 4 &#176;C.</li>
	<li>Rinse the plates twice with 2 mL of PBS to remove unbound collagen and matrigel.</li>
	<li>Place the plates in a 37 &#176;C tissue culture incubator during dissections.</li>
	<li>Prepare a moist chamber by placing 2-3 sheets of thick absorbent paper into a plastic bag or a sterile 15 cm dish and use a pipette to wet the surface of the paper with sterile water.</li>
	<li>Place the chamber under UV light for 5 min to sterilize.</li>
	<li>Dissect desired muscles from a 4 to 8-week old mouse. To sterilize muscle tissue, rinse gently in PBS containing 40 &#956;g/mL gentamicin. 
	NOTE: For quadriceps and gastrocnemius muscles, plate one muscle per plate. For soleus, plantaris and EDL, combine muscles from both legs in one plate.</li>
	<li>Use sterile forceps to transfer the muscle to a sterile 10 cm non-coated Petri dish. Add 0.5-1.0 mL of Plating media over the muscle such that the tissue is moist but not floating (Figure 1A).</li>
	<li>Use a sterile scalpel or razor blade to gently slice the muscle into small fragments (approximately 1-3 mm3). 
	NOTE: It is important to minimize handling the muscle tissue for best results.</li>
	<li>Use forceps or a pipette to transfer the muscle fragments onto the surface of a pre-coated 6-cm plate. Very gently overlay an additional 0.8 mL of Plating Media over the tissue. There should be enough media to keep the tissue pieces hydrated but not floating (Figure 1B).</li>
	<li>Place the 6-cm dishes containing the muscle fragments inside the moist chamber and return to an incubator (37 &#176;C, 5% CO2) for 48 h (Figure 1C). 
	NOTE: It is vital that muscle fragments adhere to the surface of the plate to allow for myoblast outgrowth. Do not move the plates/chamber for at least 48 h.</li>
	<li>After 48 h, carefully check the plates to ensure that muscle fragments have adhered. Overlay with 2 mL of Plating Media, taking care not to dislodge the fragments. 
	NOTE: If there is visible contamination or debris on the plates, carefully wash the muscle pieces. Plate 2 mL of PBS/gentamicin solution at the edge of the plate and tip gently to wash over the muscle tissue. Remove the PBS and repeat the wash step. Following the second wash, overlay 2 mL of Plating Media.</li>
	<li>Keep the plates in the 37 &#176;C incubator for up to an additional 3 days (5 days from original dissection) before harvesting myoblasts. Check every other day for outgrowth of myoblasts. 
	NOTE: The appearance of cells emerging from muscle explants will be variable and heterogeneous. Primary myoblasts appear as small, round, and bright cells. However, it is neither necessary nor reliable to identify them by their appearance at this stage (Figure 2).</li>
</ol>
2. Harvesting Outgrowing Myoblasts
<ol>
	<li>Prepare and pre-warm Myoblast Media (17.5 mL of DMEM, 17.5 mL of HAMS F12, 10 mL of FBS, 5 mL of amniotic fluid medium supplement), trypsin, and PBS/gentamicin. Coat one T25 flask per muscle group being harvested with Coating Solution and as described in step 1.2 (see Table 1).</li>
	<li>Remove Plating Media and gently rinse the muscle explants with 2 mL of PBS/gentamicin. Quickly remove PBS (rinse one plate at a time and do not let the plate sit in PBS at this step).</li>
	<li>Gently add 1 mL of PBS/gentamicin and place the plate in the 37 &#176;C incubator for 1 min.</li>
	<li>Use a P1000 pipette to collect PBS/cells in a 15 mL centrifuge tube.</li>
	<li>Add 1 mL of trypsin to the plate and return to the 37 &#176;C incubator for 3 min. Gently tap the plates to dislodge myoblasts. Collect the trypsin/cells and combine with the PBS collection. Add 8 mL of Myoblast Media to the centrifuge tube and gently invert to mix.</li>
	<li>Gently overlay 2 mL of Plating Solution on the muscle plates and return to the 37 &#176;C incubator.</li>
	<li>Spin the centrifuge tubes containing cells in a centrifuge for 3 min at 200 x g.</li>
	<li>Aspirate the supernatant to ~1 mL, being careful to avoid cell pellet. Gently add Myoblast Media (see Table 1) and transfer the cells to the pre-coated flask and place in the 37 &#176;C incubator. This is the P0 harvest. If observed under a microscope, there may be very few cells (Figure 3).</li>
	<li>Repeat the harvest described above every other day up to three times. After the third harvest, discard the explants.</li>
</ol>
3. Expansion and Enrichment of Proliferating Myoblasts
NOTE: The P0 harvest will be heterogeneous (~60% myoblasts). The next 2 passages use PBS to selectively harvest myoblasts. Many of the more adherent cells will be left behind and the rapidly proliferating myoblasts will be &#8805;95% pure within 2 passages. Once myoblasts are established, they should be maintained at a low density to avoid spontaneous differentiation.
<ol>
	<li>For each T25 flask of ~40-50% confluent cells from section 2, coat one T75 flask with 5 mL of Coating Solution and place at 4 &#176;C for 1-4 h.</li>
	<li>Remove Coating Solution from the flasks and return to stock solution. Rinse the flasks twice with 2 mL of PBS/gentamicin and place in the 37 &#176;C incubator.</li>
	<li>Aspirate the media from P0 myoblast T25 flasks. Rinse the cells briefly with 2 mL of warm PBS/gentamicin. Aspirate PBS from the flask. 
	NOTE: The purpose of this step (3.3) is to reduce the possibility of bacterial contamination. If performed quickly and gently, it should not result in loss of myoblasts. It can be omitted to maximize myoblast preservation if desired.</li>
	<li>Pipette 2 mL of warm PBS (not trypsin) into each flask containing myoblasts. Place the flasks with PBS into the 37 &#176;C incubator for 3 min. 
	NOTE: Myoblasts should easily detach from flasks with PBS. Using PBS rather than trypsin at this step is critical for reducing contamination of the myoblast population with other cell types.</li>
	<li>Remove the cells from the 37 &#176;C incubator and firmly tap side of the flasks to dislodge the cells. Check under a light microscope for freely floating myoblasts.</li>
	<li>Place the flasks upright in a tissue culture hood and rinse the bottom of the flasks with 10 mL of Myoblast Media 2-3 times to ensure all cells are dislodged.</li>
	<li>Collect the cell/media mixture in a 15 mL centrifuge tube. Centrifuge for 3 min at 200 x g.</li>
	<li>Aspirate the media to around 1 mL, being careful to avoid the cell pellet. Gently add an appropriate volume of Myoblast Media to centrifuge tube and gently mix.</li>
	<li>Distribute the cell mixture to new T75 flasks. Add 10 mL of Myoblast Media to each new T75 flask. Gently shake the flasks horizontally to distribute the cells and place in the 37 &#176;C incubator overnight.</li>
	<li>Two days later, passage once more with PBS, splitting each T75 flask into three T75 flasks. 
	NOTE: Do not allow myoblasts to become more than 50%-60% confluent, as this would cause them to start differentiating and lose prolifaterive capacity.</li>
	<li>For additional passages, use trypsin. Passaging twice with PBS yields &#62;95% myoblasts; attempts to further improve the purity tends to result in poorer differentiation.</li>
</ol>
4. Differentiation of Primary Myoblasts to Myotubes
<ol>
	<li>Plate P2 (or later passage) myoblasts in Myoblast Media on coated plates (see Table 1 for suggested coating and plating volumes and cell numbers). Two or three days later, when cells are at 70-80% confluency, change the media to Differentiation Media (24 mL of DMEM, 24 mL of HAMS F12, 1.5 mL of heat inactivated horse serum, 0.5 mL of Insulin-Selenium-Transferrin).</li>
	<li>Change Differentiation Media every other day during differentiation of primary myoblasts into myotubes. Differentiation is typically complete by Day 4-5 and will be marked by elongated, fused cells that spontaneously twitch. Perform experiments on differentiated myotubes within 6 days. Typically cells are assayed five or six days after initiating differentiation.</li>
</ol>
5. Measuring Oxygen Consumption Rate in Myoblasts or Myotubes in 96-well Plates
<ol>
	<li>Coat 96-well plates with 25 &#956;L of coating solution per well. Centrifuge the plates at 58 x g for 1 min to remove any bubbles.</li>
	<li>Incubate the plates in coating solution for 1-4 h at 4 &#176;C. Use multi-channel pipette to remove the coating solution and wash three times in 25 &#956;L of cold PBS/gentamicin. Centrifuge at least one of these washes to ensure that no bubbles are trapped.</li>
	<li>Add 40 &#956;L of Differentiation Media to each well. Centrifuge the media on the coated plate with no cells for 1 min at 58 x g to avoid bubbles and remove surface tension to get a uniform layer of media.</li>
	<li>Add cells suspended in an additional 40 &#956;L of media to each well. Spin again at 58 x g. 
	NOTE: Consistency in plating is important for best results and the number of cells plated per well should be optimized for each experimental setup, and will likely be in the range of ~10,000-25,000 myoblasts plated in differentiation media per well of a 96-well plate.</li>
	<li>Gently change the media daily during differentiation. Do not aspirate the media but rather remove it with a pipette, leaving a small volume behind to avoid dislodging the cells or exposing them to the air. For example, replace 50 &#956;L at a time for three repeats rather than all 80 &#956;L at once.</li>
	<li>Use 8-15 wells per condition. It may be necessary to omit some wells if cells do not form a uniform layer of myotubes. 
	NOTE: Omit wells on the edges of the plate because they are highly susceptible to evaporation. Vary the plate setup for experimental replicates to avoid systematic errors.</li>
	<li>Perform desired assay on differentiated myotubes within 1-2 days of full differentiation (Day 4-6 from start of differentiation). Recommended instruments and reagents for measurement of oxygen consumption rates are listed in the Table of Materials.</li>
</ol>

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Skeletal muscle is vital for the establishment and maintenance of metabolic homeostasis11. The study of muscle physiology is complicated by interindividual variability, as well as difficulty in obtaining samples, particularly in the case of human studies. Cultured primary myotubes have been shown to recapitulate many features of muscle physiology, including calcium homeostasis12, regeneration of damaged muscle tissue5, metabolic alterations in response to exercise13, and alterations to metabolism resulting from diseases such as diabetes14. Primary culture of myoblasts and myotubes from mice enables investigation of muscle cells harboring well defined genetic manipulations, and provides a complement to studies of myotubes derived from human muscle biopsies12,15,16. Therefore, methods for isolation and culture of mouse primary myoblasts and myotubes are essential to enable reproducible, high-throughput investigation of muscle cell function ex vivo. The protocol described here allows for the establishment and study of primary mouse myoblasts and myotubes under a variety of experimental manipulations.
While previous protocols have described the isolation of muscle stem cells from explant cultures, this protocol provides a method for the successful isolation of myoblasts from multiple different types of muscle tissue. In addition, this method yields a significantly larger population of stem cells for further experimental manipulation. Further, this method has been validated as yielding differentiated myotubes that express markers of mature muscle cells17, and exhibit normal physiology, such as circadian rhythms17 and mitogen activated protein kinase (MAPK) signal transduction18.
The critical steps in the protocol are the dissection and processing of the muscle tissue explants, as well as the avoidance of contamination between harvests. Care should be taken to avoid over-processing of the tissues. While smaller pieces of muscle yield larger numbers of myoblasts, excessive cutting of the muscle could prevent stem cell outgrowth. While it is important not to dislodge the explants once they are plated, careful washing of the plates with PBS/gentamicin is critical for reducing contamination. Harvested myoblasts may be frozen as P2 cells in cryovials using a 10% DMSO/90% Myoblast Media mixture. While myoblasts do not need to be maintained at a high density to facilitate growth, it is advised that cells are frozen at 40-50% confluence. Typically, one T75 flask yields 4 cryovials of cells.

While often cited as an indication of overall metabolic health, multiple studies have shown that body mass index (BMI) in older adults is not consistently associated with higher risk of mortality. To date, the only factor shown to be consistent with reduced mortality in this population is increased muscle mass1. Muscle tissue represents one of the largest sources of insulin-sensitive cells in the body, and is therefore critical in the maintenance of overall metabolic homeostasis2. Activation of skeletal muscle tissue via exercise is associated with increases in both local insulin sensitivity and overall metabolic health3. While in vivo models are essential for studying muscle physiology and the impact of muscle function on integrated metabolism, primary cultures of myotubes provide a tractable system that reduces the complexity of animal studies.
Myoblasts derived from post-natal muscles can be used to study the impact of numerous treatment and growth conditions in a highly reproducible manner. This has long been recognized and several methods for myoblast isolation and culture have been described4,5,6,7,8,9. Some of these methods use neonatal muscles and yield relatively low numbers of myoblasts5,8, requiring several animals for larger scale studies. Also, most widely used methods for culturing myoblasts use &#34;pre-plating&#34; to enrich for myoblasts, which are less adherent than other cell types. We have found the alternative enrichment method described here to be much more efficient and reproducible for enriching a highly proliferative myoblast population. In summary, this protocol enables the isolation of highly proliferative myoblasts from young adult muscle explants, via outgrowth into culture media. Myoblasts can be harvested repeatedly, over several days, rapidly expanded, and induced to differentiate into myotubes. This protocol reproducibly generates a large number of healthy myoblast cells that robustly differentiate into spontaneously twitching myotubes. It has enabled us to study metabolism and circadian rhythms in primary myotubes of mice of a variety of genotypes. Finally, we include methods for preparing myotubes for the study of oxidative metabolism, using measurements of oxygen consumption rates in 96-well plates.

<table><tbody><tr><td>&lt;strong&gt;Coating Solution:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>10569010</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 24 mL</td></tr><tr><td>HAMS F12</td><td>Lonza</td><td>12-615F</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 24 mL</td></tr><tr><td>Collagen</td><td>Life Technologies</td><td>A1064401</td><td>1.7 mL</td></tr><tr><td>Matrigel</td><td>Fisher</td><td>CB40234A</td><td>1 mL</td></tr><tr><td>&lt;strong&gt;Plating Media:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>10569010</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 12.5 mL</td></tr><tr><td>HAMS F12</td><td>Lonza</td><td>12-615F</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 12.5 mL</td></tr><tr><td>Heat Inactivated FBS</td><td>Life Technologies</td><td>16000044</td><td>20 mL; can be purchased as regular FBS and heat-inactivated by placing in a 40 &amp;deg;C water bath for 20 minutes</td></tr><tr><td>Amniomax</td><td>Life Technologies</td><td>12556023</td><td>5 mL</td></tr><tr><td>&lt;strong&gt;Myoblast Media:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>10569010</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 17.5 mL</td></tr><tr><td>HAMS F12</td><td>Lonza</td><td>12-615F</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 17.5 mL</td></tr><tr><td>Heat Inactivated FBS</td><td>Life Technologies</td><td>16000044</td><td>10 mL; can be purchased as regular FBS and heat-inactivated by placing in a 40 &amp;deg;C water bath for 20 min</td></tr><tr><td>Amniomax</td><td>Life Technologies</td><td>12556023</td><td>5 mL</td></tr><tr><td>&lt;strong&gt;Differentiation Media:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>10569010</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 24 mL</td></tr><tr><td>HAMS F12</td><td>Lonza</td><td>12-615F</td><td>Always add gentamicin (1:1,000 by volume) prior to use; 24 mL</td></tr><tr><td>Heat Inactivated Horse Serum</td><td>Sigma</td><td>H1138</td><td>1.5 mL</td></tr><tr><td>Insulin-Selenium-Transferrin</td><td>Life Technologies</td><td>41400045</td><td>0.5 mL</td></tr><tr><td>&lt;strong&gt;Other Materials:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>PBS</td><td>Gibco</td><td>14040133</td><td></td></tr><tr><td>Gentamicin</td><td>Sigma</td><td>G1397</td><td></td></tr><tr><td>TrypLE</td><td>Gibco</td><td>12604013</td><td></td></tr><tr><td>DMSO</td><td>Sigma</td><td>472301</td><td>Prepare as 10% DMSO in Myoblast Media for freezing cells</td></tr><tr><td>Forceps</td><td>Any</td><td></td><td></td></tr><tr><td>Razor Blades</td><td>Any</td><td></td><td></td></tr><tr><td>Scissors</td><td>Any</td><td></td><td></td></tr><tr><td>Whatman paper</td><td>VWR</td><td>21427-648</td><td></td></tr><tr><td>60 mm plate</td><td>VWR</td><td>734-2318</td><td></td></tr><tr><td>10 cm plate</td><td>VWR</td><td>25382-428 (CS)</td><td></td></tr><tr><td>T25 Flasks</td><td>ThermoFisher</td><td>156367</td><td></td></tr><tr><td>T75 Flasks</td><td>ThermoFisher</td><td>156499</td><td></td></tr><tr><td>Centrifuge Tubes (15 mL)</td><td>BioPioneer</td><td>CNT-15</td><td></td></tr><tr><td>&lt;strong&gt;Oxygen Consumption Rates:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Seahorse XFe96 Analyzer</td><td>Agilent</td><td>Seahorse XFe96 Analyzer</td><td>Instrument used to measure oxygen consumption rates read out by acidification of the extracellular media</td></tr><tr><td>Seahorse XFe96 FluxPak</td><td>Agilent</td><td>102416-100</td><td>96-well plates for use in XFe96 Analyzer</td></tr><tr><td>Seahorse XF Cell Mito Stress Test Kit</td><td>Agilent</td><td>103015-100</td><td>components may be purchased from other suppliers once assay is established; some recommendations are listed below</td></tr><tr><td>Seahorse XF Palmitate-BSA FAO substrate</td><td>Agilent</td><td>102720-100</td><td>components may be purchased from other suppliers once assay is established; some recommendations are listed below</td></tr><tr><td>Palmitic acid</td><td>Sigma</td><td>P5585-10G</td><td>for measurement of fatty acid oxidation</td></tr><tr><td>Carnitine</td><td>Sigma</td><td>C0283-5G</td><td>for measurement of fatty acid oxidation</td></tr><tr><td>Etomoxir</td><td>Sigma</td><td>E1905</td><td>for measurement of fatty acid oxidation</td></tr><tr><td>BSA</td><td>Sigma</td><td>A7030</td><td>used as control or in conjugation with palmitic acid for use in measurement of fatty acid oxidation</td></tr></tbody></table>

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.

Following Section 1 of the provided protocol should yield primary cells emerging from the explants that will be visible under a standard light microscope (Figure 2). A heterogeneous cell population will be seen growing out of and surrounding each muscle tissue explant. Myoblasts will appear as small, round, bright spheres. Following Section 2 of the protocol will yield early harvests of myoblasts from tissue explants, which will contain few cells and will be heterogeneous (Figure 3). Section 3 of the protocol describes passaging early harvests with PBS (rather than trypsin), which will provide a relatively pure population of myoblasts for further culturing. Following Section 4 of the protocol will yield fully differentiated myotubes for further experimental manipulation. Differentiation of myoblasts typically takes 4-6 days, during which the morphology of the cells will change from single, round spheres to elongated, fused, long multinucleated fibers (Figure 4). Following section 5 of the protocol will produce differentiated myotubes in 96-well plates to enable a variety of metabolic characterizations based on the changes in oxygen consumption and extracellular acidification rates10 (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60310/60310fig1.jpg" /> 
Figure 1: Dissection and processing of quadriceps muscle. (A) Quadriceps muscle that has been freshly dissected and rinsed with PBS prior to transfer to a 10 cm dish. 1 mL of plating media has been overlaid for processing. (B) Quadriceps muscle tissue pieces after transfer to pre-coated 6 cm plate. (C) 6 cm plates inside moist chamber prior to placement in the 37 &#176;C incubator. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60310/60310fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60310/60310fig2.jpg" /> 
Figure 2: Outgrowth of myoblasts. Outgrowth of myoblasts from quadriceps muscle explants. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60310/60310fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60310/60310fig3.jpg" /> 
Figure 3: Early passage myoblasts. P0 myoblasts after transfer and attachment to T25 flask. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60310/60310fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60310/60310fig4.jpg" /> 
Figure 4: Plating and differentiation of primary myotubes. (A) Myoblasts one day after initiating exposure to Differentiation Media. (B,C) Differentiated myotubes five (B) or six (C) days after initiating differentiation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60310/60310fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60310/60310fig5.jpg" /> 
Figure 5: Myotubes ready for measurement of oxygen consumption rates. Fully differentiated myotubes five days after plating 20,000 myoblasts in Differentiation Media in each well of a 96-well cell culture microplate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60310/60310fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants at JoVE.com

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

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 ...

isolation-differentiation-primary-myoblasts-from-mouse-skeletal

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

17.0K Views.  Department of Molecular Medicine. 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.

Video: Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

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 ...

Preparation and Culture of Myogenic Precursor Cells/Primary Myoblasts from Skeletal Muscle of Adult and Aged Humans

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting occurs. Loss of muscle mass and function is associated with muscle fiber type atrophy, fiber type switching, defective muscle regeneration associated with dysfunction of satellite cells, muscle stem cells, and other pathophysiological processes. These changes are associated with changes in intracellular as well as local and systemic niches. In addition to most commonly used rodent models of muscle ageing, there is a need to study muscle homeostasis and wasting using human models, which due to ethical implications, consist predominantly of in vitro cultures. Despite the wide use of human Myogenic Progenitor Cells (MPCs) and primary myoblasts in myogenesis, there is limited data on using human primary myoblast and myotube cultures to study molecular mechanisms regulating different aspects of age-associated muscle wasting, aiding in the validation of mechanisms of ageing proposed in rodent muscle. The use of human MPCs, primary myoblasts and myotubes isolated from adult and aged people, provides a physiologically relevant model of molecular mechanisms of processes associated with muscle growth, atrophy and regeneration. Here we describe in detail a robust, inexpensive, reproducible and efficient protocol for the isolation and maintenance of human MPCs&#160;and their progeny&#160;&#8212; myoblasts and myotubes from human muscle samples using enzymatic digestion. Furthermore, we have determined the passage number at which primary myoblasts from adult and aged people undergo senescence in an in vitro culture. Finally, we show the ability to transfect these myoblasts and the ability to characterize their proliferative and differentiation capacity and propose their suitability for performing functional studies of molecular mechanisms of myogenesis and muscle wasting in vitro.

This protocol describes a robust, reproducible and simple method of isolation and culture of myoblast progenitor cells from the skeletal muscle of adult and aged people. The muscles used here include foot and leg muscles. This approach enables the isolation of an enriched population of primary myoblasts for functional studies.

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting ...

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 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.

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 ...

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a robust and reliable method for isolating relatively large numbers of proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. The authors have used micro-dissected explant cultures to analyse the growth characteristics of skeletal muscle cells in wild-type and dystrophic muscles. Each of the components of tissue growth, namely cell survival, proliferation, senescence and differentiation can be analysed separately using the methods described here. The net effect of all components of growth can be established by means of measuring explant outgrowth rates. The micro-explant method can be used to establish primary cultures from a wide range of different muscle types and ages and, as described here, has been adapted by the authors to enable the isolation of embryonic skeletal muscle precursors.
Uniquely, micro-explant cultures have been used to derive clonal (single cell origin) skeletal muscle stem cell (SMSc) lines which can be expanded and used for in vivo transplantation. In vivo transplanted SMSc behave as functional, tissue-specific, satellite cells which contribute to skeletal muscle fibre regeneration but which are also retained (in the satellite cell niche) as a small pool of undifferentiated stem cells which can be re-isolated into culture using the micro-explant method.

The micro-dissected explants technique is a robust and reliable method for isolating proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. Uniquely, these cells have been clonally derived to produce skeletal muscle stem cell lines used for in vivo transplantation.

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a ...

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 ...

Isolation, Culture, and Transplantation of Muscle Satellite Cells

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal nuclei in muscle fibers. In adult muscle, satellite cells are normally mitotically quiescent. Following injury, however, satellite cells initiate cellular proliferation to produce myoblasts, their progenies, to mediate the regeneration of muscle. Transplantation of satellite cell-derived myoblasts has been widely studied as a possible therapy for several regenerative diseases including muscular dystrophy, heart failure, and urological dysfunction. Myoblast transplantation into dystrophic skeletal muscle, infarcted heart, and dysfunctioning urinary ducts has shown that engrafted myoblasts can differentiate into muscle fibers in the host tissues and display partial functional improvement in these diseases. Therefore, the development of efficient purification methods of quiescent satellite cells from skeletal muscle, as well as the establishment of satellite cell-derived myoblast cultures and transplantation methods for myoblasts, are essential for understanding the molecular mechanisms behind satellite cell self-renewal, activation, and differentiation. Additionally, the development of cell-based therapies for muscular dystrophy and other regenerative diseases are also dependent upon these factors.

However, current prospective purification methods of quiescent satellite cells require the use of expensive fluorescence-activated cell sorting (FACS) machines. Here, we present a new method for the rapid, economical, and reliable purification of quiescent satellite cells from adult mouse skeletal muscle by enzymatic dissociation followed by magnetic-activated cell sorting (MACS). Following isolation of pure quiescent satellite cells, these cells can be cultured to obtain large numbers of myoblasts after several passages. These freshly isolated quiescent satellite cells or ex vivo expanded myoblasts can be transplanted into cardiotoxin (CTX)-induced regenerating mouse skeletal muscle to examine the contribution of donor-derived cells to regenerating muscle fibers, as well as to satellite cell compartments for the examination of self-renewal activities.

The isolation and culture of a pure population of quiescent satellite cells, a muscle stem cell population, is essential to the understanding of muscle stem cell biology and regeneration, as well as stem cell transplantation for therapies in muscular dystrophy and other degenerative diseases.&#160;

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal ...

Single Myofiber Isolation and Culture from a Murine Model of Emery-Dreifuss Muscular Dystrophy in Early Post-Natal Development

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate filament proteins that sustain the nuclear envelope and the components of the nucleoplasm. We recently reported that muscle wasting in EDMD can be ascribed to intrinsic epigenetic dysfunctions affecting muscle (satellite) stem cells regenerative capacity. Isolation and culture of single myofibers is one of the most physiological ex-vivo approaches to monitor satellite cells behavior within their niche, as they remain between the basal lamina surrounding the fiber and the sarcolemma. Therefore, it represents an invaluable experimental paradigm to study satellite cells from a variety of murine models. Here, we describe a re-adapted method to isolate intact and viable single myofibers from post-natal hindlimb muscles (Tibialis Anterior, Extensor Digitorum Longus, Gastrocnemius and Soleus). Following this protocol, we were able to study satellite cells from Lamin &#916;8-11 -/- mice, a severe EDMD murine model, at only 19 days after birth.
We detail the isolation procedure, as well as the culture conditions for obtaining a good amount of myofibers and their associated satellite-cells-derived progeny. When cultured in growth-factors rich medium, satellite cells derived from wild type mice activate, proliferate, and eventually differentiate or undergo self-renewal. In homozygous Lamin &#916;8-11 -/- mutant mice these capabilities are severely impaired.
This technique, if strictly followed, allows to study all processes linked to the myofiber-associated satellite cell even in early post-natal developmental stages and in fragile muscles.

Here, we propose a method to efficiently obtain single muscle fibers at early post-natal developmental stages from homozygous mutant Lamin &#916;8-11 mouse model, a very severe model for Emery-Dreifuss muscular dystrophy (EDMD).

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate ...

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 ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

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

Preparation and Culture of Myogenic Precursor Cells/Primary Myoblasts from Skeletal Muscle of Adult and Aged Humans

In Vitro Differentiation of Mature Myofibers for Live Imaging

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

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

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

Isolation, Culture, and Transplantation of Muscle Satellite Cells

Single Myofiber Isolation and Culture from a Murine Model of Emery-Dreifuss Muscular Dystrophy in Early Post-Natal Development

DNA Electroporation, Isolation and Imaging of Myofibers

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion