Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

This is a protocol for the isolation of primary mouse muscle stem cells for the study of metabolism ex vivo. This protocol provides a much larger population of stem cells compared to other protocols that we've tried in the past. And importantly, once we differentiate these stem cells into myotubes, they exhibit normal physiology, so things like circadian rhythms.

When trying this procedure for the first time, take detailed notes and save several images because every tissue explant looks different and there will be variation in the outgrowth of myoblasts. Without visual demonstration, it is difficult to know if you're isolating the correct cells. We benefited from the generous help of others who showed us this method when we established it in our lab.

After preparing the dish for dissection according to the manuscript, prepare a moist chamber by placing two to three sheets of thick absorbent paper into a plastic bag and use a pipette to wet the surface of the paper with sterile water. Place the chamber under UV light for five minutes to sterilize. Dissect the desired muscle from a four to eight-week-old mouse and rinse gently in PBS containing 40 micrograms per milliliter gentamycin to sterilize.

Use sterile forceps to transfer the muscle to a sterile 10 centimeter non-coated Petri dish. Add 0.5 to one milliliter of plating media over the muscle such that the tissue is moist but not floating. Use a sterile scalpel or razor blade to gently slice the muscle into small fragments.

Use forceps or a pipette to transfer the muscle fragments onto the surface of a pre-coated six centimeter plate. Very gently overlay an additional 0.8 milliliters of plating media over the tissue. Place the six centimeter dish containing the muscle fragments inside the moist chamber and return it to an incubator at 37 degrees Celsius and 5%carbon dioxide for 48 hours.

Prepare and prewarm myoblast media, trypsin and PBS-gentamycin in a water bath at 37 degrees Celsius. To coat a T25 flask for each muscle group, add two milliliters of coating solution to the surface of the flask, shake gently to create an even coat on the surface, and incubate the flask at four degrees Celsius for one hour. Now with a pipette, remove the plating media from the muscle explants and gently rinse the muscle explants with two milliliters of PBS-gentamycin.

Quickly remove the PBS and do not let the plate sit in the PBS. Then gently add one milliliter of PBS-gentamycin and place the plate in the 37 degree Celsius incubator for one minute. Use a P1000 pipette to collect the PBS with cells into a 15 milliliter centrifuge tube.

Next, add one milliliter of trypsin to the plate and return it to the 37 degree Celsius incubator for three minutes. Gently tap the plates to dislodge the myoblasts. Collect the trypsin with cells and combine with the PBS collection.

Add eight milliliters of myoblast media to the centrifuge tube and gently invert to mix. Gently overlay two milliliters of plating solution on the muscle plates and return the plates to the 37 degree Celsius incubator. Spin the centrifuge tubes containing cells in a centrifuge for three minutes at 200 times g.

Aspirate the supernatant and keep approximately one milliliter in the tube. Gently add myoblast media, transfer the cells to the pre-coated flask and place the flask in the 37 degree Celsius incubator. Aspirate the media from P0 myoblast T25 flasks.

Rinse the cells briefly with two milliliters of warm PBS-gentamycin then aspirate the PBS from the flask. Pipette two milliliters of warm PBS into each flask containing myoblasts. Place the flasks with PBS into the 37 degree Celsius incubator for three minutes.

Then firmly tap the side of the flask to dislodge the cells. Check under a light microscope for freely floating myoblasts. Place the flasks upright in a tissue culture hood and rinse the bottom of the flasks with 10 milliliters of myoblast media two to three times to ensure all cells are dislodged.

Collect the cell media mixture in a 15 milliliter centrifuge tube. Centrifuge for three minutes at 200 times g. Aspirate the media to leave around one milliliter in the tube being careful to avoid the cell pellet.

Gently add an appropriate volume of myoblast media to the centrifuge tube and gently mix. Distribute the cell mixture to new T75 flasks six milliliters each. Add 10 milliliters of myoblast media to each new T75 flask.

Gently shake the flasks horizontally to distribute the cells and place them in the incubator at 37 degrees Celsius overnight. In this protocol, primary cells emerged from the explants were observed under a standard light microscope. Early harvests of myoblasts appeared as small round and bright spheres.

Differentiation of myoblasts took four to six days during which the morphology of the cells changed from single round spheres to elongated fused long multi-nucleated fibers. Fully differentiated myotubes were yielded which were ready for measurement of oxygen consumption rates. These cells can be used for a number of downstream applications.

For instance, we frequently use them to study substrate flux in Seahorse instruments.