The overall goal of this protocol is to provide a robust, reproducible and simple procedure for the isolation of primary myoblasts from adult and aged humans. This protocol proposes an optimized workflow for the isolation of primary human myoblasts as an in vitro model for the functional studies of processes associated with muscle homeostasis during aging. This protocol maintains the cells'natural behavior by avoiding a stressor such as centrifugation, and allows isolation and long time culture of primary myoblast to retain phenotype.
Begin this procedure by preparing a 60 milliliter petri dish or two wells of a six well plate. Next, pipette a minimum amount of laminin solution to completely cover the surface of the dish. Then, incubate the culture dish for at least 30 minutes in a humidified incubator at 37 degrees Celsius, before plating the cells.
Next, prepare the enzymatic solution in serum free DMEM. Mix well and filter the enzymatic solution through a 0.2 micrometer filter membrane for sterilization. Following sample collection, keep the muscle at four degrees Celsius in DPBS until digestion.
Prepare at least two milliliters of enzymatic solution for 18 to 20 milligrams of muscle tissue and warm it to 37 degrees Celsius before tissue dissociation. Next, immerse the muscle biopsy briefly in 70 percent ethanol. Wash it with fresh DPBS and then 70 percent ethanol and then place the tissue on a new petri dish in enough enzymatic solution to cover the sample.
Discard as much fibrotic and fat tissue as possible and tear the muscle gently into small but distinguishable pieces. Afterward, transfer the sample into a 50 milliliter tube with at least one milliliter of enzymatic solution. Incubate the tissue at 37 degrees Celsius for up to 30 or 40 minutes, and move the muscle tissue by gently agitating the tube every five to 10 minutes.
Next, coat the pipettes with media to avoid adhesion of the released cells to the plastic wall of the pipettes. Then, add four milliliters of sterile growth media to stop the digestion. Pipette up and down several times to help the cells release from the muscle fibers.
Subsequently, filter the muscle solution through a 70 micrometer cell strainer over a 50 milliliter conical tube. Wash the remaining cells with more media. And filter it through the strainer.
Then, centrifuge the sample for five minutes at room temperature to pellet the cells. After five minutes, discard the supernatant carefully. Then, dissolve the pellet into F12 growth medium.
In this step, collect the culture vessel previously coated with laminin from the incubator. Carefully remove the excess of laminin from the culture dish, and avoid touching the surface. Then, wash the culture dish gently with DPBS.
Next, plate the cells directly on the laminin coated vessel. And incubate them for 24 hours in a humidified incubator at 37 degrees Celsius. Visualize the cells under the bright field microscope on the following day.
Small round cells attach to the surface and the remaining debris should be seen in the culture media. After that, change the media to fresh F12 media. In this procedure, change the media every two to three days, and split the cells as soon as groups of cells are visible under the microscope, in order to avoid spontaneous differentiation.
For cell passaging, remove the media and wash the cells twice with DPBS. Add the least amount of 0.25 percent EDTA trypsin needed to cover the cells'surface. Rock gently to ensure all the cells are covered by the detachment solution, and then incubate for 10 seconds at room temperature before removing it.
After that, incubate the cells in a humidified incubator at 37 degrees Celsius for three to five minutes. Tap gently and check under the bright light microscope that the cells are rounded but not completely detached from the surface. Then, add five milliliters of growth medium to collect the cells, mix well, and transfer the cells with the new media into a T75 flask.
Subsequently, wash the remaining cells with another 5 milliliters of growth medium. To preplate at the first passage, incubate the cells at 37 degrees Celsius for 40 minutes to one hour. After that collect the supernatant containing the unattached cells and incubate them in a new T75 flask.
This should enrich the culture and myoblasts, as most of the fiberblasts should've attached in the first flask. Change the media every two to three days. And split the cells as soon as they achieve 70 percent of confluency.
For differentiation, once the cells reach 70 to 80 percent confluency, change the culture medium to differentiation medium. The cells should be differentiated within five to seven days, depending on the quality and the purity of the myoblast culture. At passage zero, and after five days of being plated, cells are still round and small, but visible under bright light microscope.
Myoblasts will then adopt an elongated shape as in passage. As shown here, MyoD is expressed in myoblasts, but not in fiberblasts. Human primary myoblasts can be characterized using staining for cell viability, Ki67 immunostaining for proliferation, MF20 immunostaining for differentiation, and senescence associated betagalactocidase staining for senescence assays.
Primary myoblasts can be used for functional assays. In this figure, MF20 immunostaining of the differentiated primary myotubes from adult humans show the effects of overexpression or inhibition of miR-378 on myotube size and number. QPCR analyses show that miR-378 mimic and anti miR transfections were successful, and led to changes in the relative expression of miR-378 and LGF 1R, a validated miR-378 target gene, following miR-378 overexpression or inhibition in human primary myoblasts.
Once mastered, this technique can be performed in less than two hours. This robust and reproducible protocol is particularly useful when other, more sophisticated and expensive matters, such as cell sorting, are not possible or practical. This simplified technique is useful for researchers in the field of muscle to explore the phenomenon of aging in human in vitro model systems.
In summary, this proposed method allows the isolation of myogenic progenitor cells from the muscle of not only adult but also aged humans for functional studies in vitro that can also be applied to animal models.
