The overall goal of this procedure is to generate mature myofibers in vitro to study the dynamic processes of myofiber differentiation using isolated neonatal mouse myoblasts. This method can help answer a key question in the medicine field about the transverse shear triad formation, sarcomere and myofibril assembly and organelle positioning that occur during myofiber differentiation. One advantage of this technique is that the anti and myofibril differentiation process can easily be imaged by time lapse microscopy in vitro.
Begin by sterilizing the skin of a six to eight day post natal mouse with 70%ethanol. Next, make an incision in the dorsal skin and pull the skin gently toward the hind limbs until the hind limb musculature is completely exposed. Remove the fat tissue without damaging the muscles.
Then, to remove the dorsal hind limb muscles, keep the limb stretched and expose the heel tendons. Starting from the tendons, use the scissors to gently cut upwards along the bone. Placing the muscles in ice cold PBS as they are harvested.
Then, pinch the quadriceps with fine tip forceps and cut around the muscle without damaging the femur or the knee joint. Pooling the quadriceps with the other tissue samples. In a sterile laminar flow cell culture hood, discard the PBS from the collection dish and use sterile curved scissors to mince the tissues.
When all the samples have been processed, transfer the tissue pieces into a 50 milliliter conical centrifuge tube containing five milliliters of digestion mix for a 90 minute incubation at 37 degrees Celsius with agitation. At the end of the incubation, stop the digestion with six milliliters of dissection medium and pellet the remaining tissue by centrifugation. The isolation of a clean cell suspension after digestion is crucial for the success of the protocol.
If necessary, aspiration of the debris or extra centrifugations can be performed to avoid contamination of the sample. Carefully collect the supernatant and centrifuge the cell suspension. Resuspending the second pellet in five milliliters of fresh dissection medium.
Next, filter the slurry through a 40 micron cell strainer and add 25 milliliters of dissection medium to the single cell suspension. Incubate for four hours in a cell culture incubator to allow fibroblast adhesion. At the beginning of the last hour of the incubation, coat two 35 millimeter dishes per mouse with 500 microliters of 1%basement membrane matrix in cold IMDM medium for one hour at room temperature.
At the end of the pre plating, collect the supernatant and centrifuge the floating cells. Resuspend myoblasts and growth medium, and after counting cells, adjust the volume to the appropriate cell concentration for plating. Then wash the basement membrane matrix coated plates with DPBS and immediately plate the cells for incubation in the cell culture incubator.
After three days, treat the cells with the appropriate transfection reagents according the the manufacturers instructions and incubate the cells with lipid complexes of interest for five hours at 37 degrees Celsius in 5%carbon dioxide. Then, perform two washes in differentiation medium and return the cells to the incubator overnight. The following day, replace the medium in each dish with 200 microliters of 50%basement membrane matrix and ice cold differentiation medium.
After another 30 minutes in the cell incubator, supplement the cultures with agrin and fresh differentiation medium, monitoring the differentiation and viability of the cells daily until they reach full maturation. For immunostaining, at the experimental time point of interest, wash the cells in DPBS followed by fixation in 4%paraformaldehyde at room temperature. After 10 minutes wash the cells two more times in DPBS, followed by permeabilization with 0.5%Triton X 100 for five minutes at room temperature.
After two more DPBS washes, block the nonspecific binding with blocking solution for 30 minutes at room temperature. Then, incubate the cells with primary antibody of interest diluted in blocking solution overnight at four degrees Celsius. The next morning, rinse the cells with three five minute washes in DPBS at room temperature and incubate the cultures with the appropriate secondary antibody in 0.2 micrograms per milliliter of DAPI for one hour.
After three more DPBS washes as just demonstrated, mount the samples in 200 microliters of mounting medium and image the cells. At proliferation day two, the myoblasts should have adhered to the plate and should display the typical fusiform shape, with extensive proliferation that leads to spontaneous myotube formation. The myotubes quickly elongate and display multiple centrally aligned nuclei with the number of mature myofibers increasing with time displaying higher cell thickness, striations, and peripheral nuclei.
Myofibers fixed at differentiation day eight present transversal triads as confirmed by imaging the components of the T tubules and the sarcoplasmic reticulum, which are expected to colocalize at the triads. From differentiation day three onwards, the cells display spontaneous twitching that can be observed coupled with the calcium peaks after calcium sensor transfection. Further, after imaging, a 3D reconstruction can be made to better understand the organization and dynamics of muscle complex structures.
While attempting this procedure, it's important to remember to be fast and to avoid keeping these sensitive cells for extended periods outside of an incubator. Following this procedure, other methods like electron microscopy, super resolution microscopy, laser micro dissection or laser oblation can be performed for further downstream analysis for these neonatal muscle cells. After watching this video you should have a good understanding of how to isolate myoblasts from newborn muscles and to cultivate them for the in vitro generation of highly differentiated myofibers under optimal conditions for time lapse imaging.
