Isolation, Culture, and Differentiation of Primary Myoblasts Derived from Muscle Satellite Cells.

The skeletal muscle is key for body mobility and motor performance, but aging and diseases often lead to progressive loss of muscle mass due to wasting or degeneration of muscle cells. Muscle satellite cells (MuSCs) represent a population of tissue stem cells residing in the skeletal muscles and are responsible for homeostatic maintenance and regeneration of skeletal muscles. Growth, injury, and degenerative signals activate MuSCs, which then proliferate (proliferating MuSCs are called myoblasts), differentiate and fuse with existing multinuclear muscle cells (myofibers) to mediate muscle growth and repair. Here, we describe a protocol for isolating MuSCs from skeletal muscles of mice for in vitro analysis. In addition, we provide a detailed protocol on how to culture and differentiate primary myoblasts into myotubes and an immunofluorescent staining procedure to characterize the cells. These methods are essential for modeling regenerative myogenesis in vitro to understand the dynamics, function and molecular regulation of MuSCs.


PBS-T (see Recipe 13).
B. Muscle dissection 1. Sacrifice a mouse aged 4-6 weeks following the approved protocol in your laboratory. In this case, cervical dislocation is used.
Note: Dissection of a mouse does not need to be performed inside an A2 biosafety cabinet. Note: It is recommended to wash several times until hair is completely removed from muscle tissue.
2. Blot dry muscles on tissue paper.
3. Transfer muscle to 1.5 ml Eppendorf tube. 4. Mince the muscle using dissection scissors inside a 1.5 ml Eppendorf tube.
Note: Finely minced muscle will yield more myoblasts.
5. Transfer the minced muscle into a 15 ml Falcon tube. 6. Digest the muscle with digestion medium (5 ml per mouse) in a 37 °C water bath for 12 min, shaking the tube every 2 min. 7. Mix the digested muscle with a 10 ml serological pipette until the mixture can be smoothly pipetted.
8. Digest for another 12 min in the 37 °C water bath, shaking the tube every 2 min. 9. Stop digestion by adding 5 ml neutralization medium. 10. Place a 70 µm sterilized cell strainer on top of a 50 ml tube, and pre-wet the strainer with 3 ml neutralization medium.
Note: Pre-wetting is important for cells to pass smoothly through the cell strainer.
11. Collect the media containing cell mixture using a 10 ml serological pipette and filter through the 70 µm sterilized cell strainer.
Note: Collect cell filtrate underneath cell strainer using 1 ml pipette.
12. Spin the cell mixture at 2,000 x g for 5 min at room temperature.
13. Discard the supernatant using a vacuum pipette. Copyright  14. Resuspend the cell pellet with 5 ml growth medium. 15. Seed the cells from one mouse in a non-coated 100 mm culture plate and incubate at 37 °C for 4 days, and supplement 5 ml of growth medium on top of pre-existing medium each day for 3 days.

Notes:
a. This method only utilizes mechanical detachment, therefore trypsin is not necessary.

b. Confirm if most cells were detached from the plate by checking under a microscope.
3. Centrifuge the 50 ml tube at 2,000 x g for 5 min.

Note: Let centrifuge stop without applying brake. This usually takes a couple of minutes.
4. Aspirate the media from the 50 ml tube. 10. Steps D2-D9 (Pre-plating) can be repeated multiple times to obtain ≥ 95% purity of myoblasts.

Note: Pre-plating can be repeated twice to increase myoblast purity. Myoblasts should appear small and rounded, while fibroblasts will appear elongated and at times have bipolar processes.
E. Culturing and differentiation 1. Culture the purified primary myoblasts in the growth medium, changing the media every 2 days.
Maintain the cell density under 80% confluency to prevent the fusion of primary myoblasts.
Note: Collagen type I coated plate is not required after pre-plating step. Copyright  Note: More washing is recommended to minimize background staining. 16. Mount the coverslip with a drop of mounting medium and allow to dry.

Data analysis
This protocol includes visualization and evaluation of myoblasts in vitro before and after differentiation, after immunostaining with specific myogenic markers. Primary myoblasts cultured in growth medium were fixed by the addition of 4% PFA and were stained with Pax7 and MyoD, markers for undifferentiated myoblasts (Figure 1). After primary myoblasts were seeded on a Matrigel-coated plate, growth medium was changed to differentiation medium once the confluency of myoblasts reached 80%. After 3 days of differentiation, myoblasts were fixed and stained with MyoG and MF20, markers of early and late stages of myogenic differentiation, respectively ( Figure   2). For research purposes, myoblasts can be stained with other antibodies, but it is recommended to co-stain with one of the myogenic markers to confirm the differentiation status of the myoblasts.  4. Additional rounds of pre-plating can be performed if myoblast purity is less than 95%.
5. For staining, incubation of primary antibody at room temperature for 1 h is possible, but overnight incubation at 4 °C is strongly recommended. For 50 ml differentiation medium, DMEM will be supplemented with 1 ml horse serum and