Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo

Sacrificing of animals must be performed in accordance with the national regulations for animal experimentation. The protocol described here was performed in accordance with the guidelines of the Leibniz Institute on Aging – Fritz Lipmann Institute and the European Union (EU) directive 2010/63/EU (license number for organ harvesting: O_JvM_18-20). Essential steps of the protocol are summarized in Figure 1.

1. Preparation of culture plates, media, and Pasteur pipettes

NOTE: All materials and equipment necessary to isolate and culture single myofibers need to be as sterile as possible. Therefore, it is recommended to isolate single myofibers under a semi-sterile dissection hood.

1. Use sterile HS (horse serum) to coat tissue culture plates. Coating prevents the attachment of single myofibers to the plastic surface. For each mouse, 4 wells of a 12-well plate for the isolation and a dedicated number of wells of a 24-well plate for culturing are required. Incubate the wells of a 12-well plate with 1 mL and the wells of a 24-well plate with 0.5 mL of HS for 5 min at RT (room temperature), then remove the HS and let the plates dry for another 5 min.
   NOTE: HS can be collected and re-used for coating purposes multiple times, if kept sterile.
2. Prepare myofiber isolation medium by supplementing DMEM (Dulbecco's modified Eagle's medium with 4.5 g/L glucose and sodium pyruvate) with 20% FBS (fetal bovine serum), filter through 0.22 µm filter. Add medium to the pre-coated isolation plates (12-well plate, 2-4 mL isolation medium per well) approximately 30 min before the isolation and equilibrate in a humidified 37 °C incubator with 5% CO₂.
3. Prepare myofiber culture medium by supplementing DMEM (Dulbecco's modified Eagle's medium with 4.5 g/L glucose and sodium pyruvate) with 20% FBS (fetal bovine serum) and 1% chicken embryo extract, filter through 0.22 µm filter. Add 0.5 mL medium to each well of the pre-coated culture plates (24-well plate) approximately 30 min before the isolation and equilibrate in a humidified 37 °C incubator with 5% CO₂.
4. Prepare collagenase digestion solution by dissolving 0.2% (w/v) collagenase type 1 (from Clostridium histolyticum) in DMEM (Dulbecco's modified Eagle's medium with 4.5 g/L glucose and sodium pyruvate), filter through 0.22 µm filter. For two EDL (extensor digitorum longus) muscles use 2.5 mL collagenase solution in a sterile 15 mL reaction tube. Additionally, pre-heat the solution ~10 min in a circulating water bath at 37 °C before starting the isolation.
5. Prepare sterile Pasteur pipettes for trituration of the collagenase-digested muscles. Use a diamond pen to cut the Pasteur pipettes.
   1. For each mouse, use one large bore pipette with an opening of about 0.3 cm and a length of about 10-12 cm and a second glass pipette with a small opening of about 0.1 cm and a length of approximately 22 cm (Figure 2F).
   2. Smoothen the edges of both pipettes by heat polishing with the flame of a Bunsen burner. Hold the pipette tip for 5-10 s into the flame with gentle movement to allow equal heat distribution until the sharp glass edges smoothen.
   3. Immediately before use, coat both types of glass pipettes with sterile HS by filling the whole pipette with ~ 2 mL of HS for 5 min, thereafter, eject the HS and let the pipettes dry for 5 min at RT.

2. EDL muscle isolation and collagenase digestion

1. Spray all equipment with 70% ethanol to avoid contamination.
2. Sacrifice the mouse in accordance with the national regulations for animal experimentation.
3. Spray the hindlimbs of the mouse with 70% ethanol. Use hardened fine curved scissors (24 mm cutting edge) and fine forceps (Dumont 7, curved or straight) to remove the skin and to expose the underlying muscles. Avoid any contact of the underlying muscles with fur (increases the risk of contamination).
4. Remove the surrounding fascia with fine curved forceps without damaging the underlying muscles (Figure 2A). Close the forceps to avoid bending.
5. Use curved forceps to expose the distal tendons of the TA (tibialis anterior) and EDL muscle. To remove the TA, grab the distal TA tendon with forceps and cut with fine Vannas spring scissors (5 mm cutting edge, 0.35 mm tip diameter). While holding the TA at the tendon, pull it towards the knee and cut the muscle close to the knee (Figure 2B), the EDL muscle is now exposed.
   NOTE: Make sure that only the TA tendon is grabbed in this step, otherwise the underlying EDL might get damaged. When cutting off the TA muscle make sure that the tendons at the knee can be seen easily afterwards.
6. Lift the distal EDL tendon with fine curved forceps and cut with fine Vannas spring scissors (Figure 2C). Expose the proximal EDL tendon by carefully pulling the EDL towards the knee. Cut the proximal tendon with fine Vannas spring scissors. Transfer the EDL muscle to the 37 °C preheated 2.5 mL of collagenase digestion solution in the 15 mL reaction tube from step 1.4. (Figure 2D).
   NOTE: Make a small incision at the outer knee connective tissue to fully expose the proximal EDL tendon. Make sure to only grab the tendons and not to stretch the EDL too much.
7. Repeat steps 2.3. to 2.6. with the second EDL. Add both EDL muscles to the same 15 mL reaction tube filled with 2.5 mL collagenase digestion solution.
8. Incubate the EDL muscles in the reaction tube at 37 °C in a circulating water bath.
   NOTE: Incubation time depends on several factors, such as collagenase activity, age of the mouse and amount of fibrotic tissue. The typical incubation time for EDL muscles of adult mice (2-6 months of age) is 1-1.5 h and for aged mice (18 months of age) 1.5-2 h.
9. To avoid excessive digestion of the muscles, check the muscles during the digestion time. Stop the digestion when muscles loosen up and single myofibers are visible (Figure 2E). Transfer muscles carefully with the large bore pipette to the first well of the prewarmed 12-well plate containing myofiber isolation medium (Figure 2G).

3. Myofiber dissociation and culture

1. For the following steps use a stereo binocular microscope, preferentially equipped with a heating plate (37 °C). Use the large bore pipette to flush the muscles with warm isolation medium until single myofibers are released. Dissociate the muscles with the large bore pipette until the desired number of myofibers are floating freely in the solution.
   NOTE: Avoid excessive trituration to reduce the risk of damaged myofibers. Using a heating plate for myofiber dissociation is strongly advised since temperature drops during the isolation process, which will result in myofiber death.
2. Transfer non-contracted myofibers (Figure 2H) with the HS coated small bore glass pipette to the second well filled with isolation medium to wash away debris and contracted (damaged) myofibers (Figure 2I).
   NOTE: To avoid excessive movement of isolated myofibers, they can be transferred to the second well and then the trituration process can be continued.
3. Transfer non-contracted myofibers to the next (3^rd) well filled with isolation medium to wash again.
4. Use the small-bore glass pipette, coated with HS, to transfer approximately 50-100 non-contracted myofibers to the 24-well plate containing myofiber culture medium.
5. Incubate myofibers at 37 °C, 5% CO₂ for dedicated time (96 h maximum are recommended).
   NOTE: MuSCs divide once either planar or apical-basal after 42 h of culture. Additionally, after 72 h of culture MuSCs form multicellular clusters consisting of self-renewing, proliferating or committed (differentiated) MuSCs.

4. siRNA transfection

1. 4 h after myofiber isolation, transfect myofiber associated MuSCs (50-100 non-contracted myofibers in one well of the 24-well plate filled with 500 µL culture medium) with lipid based transfection reagent, e.g. RNAiMAX according to the manufacturer's protocol with a final concentration of 5 pmol siRNA. Therefore, add 25 µL of Opti-MEM with the respective volume of siRNA to 25 µL Opti-MEM containing 1.5 µL of the transfection reagent. Incubate the reaction mix for 5 min and add it then to the 500 µL of culture medium.
   NOTE: A second transfection after 24 h or 48 h is recommended for longer culture periods, e.g., more than 48 h. A change of medium after transfection is not necessary.

5. Fixation and IF staining

1. For immunostaining use a stereo binocular microscope. All volumes in the following steps are adjusted to one well of a 24-well plate. Perform all following steps using a HS-coated small-bore pipette.
2. Carefully discard the myofiber culture medium while leaving some solution in the well (approximately 100 µL per 24-well). Do this for all further steps unless stated otherwise to allow floating of the myofibers. Add 500 µL of 2% PFA to fix the myofibers with their adjacent MuSCs, incubate for 5 min at room temperature (RT).
3. Remove the supernatant carefully and wash the myofibers three times with PBS (pH 7.4, 500 µL for 5 min at RT each).
4. Add 500 µL permeabilization solution (0.1% Triton X-100, 0.1 M Glycine in PBS, pH 7.4), incubate for 10 min at RT.
5. Remove the permeabilization solution and add 500 µL blocking solution (5% HS in PBS, pH 7.4) for 1 h at RT.
   NOTE: Check the recommended blocking solution for primary antibodies to avoid unspecific binding.
6. Remove the blocking solution and add 300 µL of primary antibody dilution (e.g., anti-Pax7 (PAX7, DSHB, undiluted), anti-MyoD (clone G-1, Santa Cruz, 1:200)) per well. Incubate at 4 °C overnight.
7. Wash three times with 500 µL of PBS per well (5 min at RT).
8. Remove PBS and add 300 µL of secondary antibody dilution (e.g., anti-mouse-IgG1-546 and anti-mouse-IgG2b-488 1:1000) per well. Incubate 1 h at RT protected from light. For the following steps, light reduced conditions are recommended.
9. Wash two times with 500 µL of PBS per well (5 min at RT).
10. Perform DAPI staining by using 500 µL of the solution per well (final DAPI concentration 10 µg/mL) for 5 min at RT.
11. Wash two times with 500 µL of PBS per well (5 min at RT).
12. Use a hydrophobic pen to draw a circle on a microscopic glass slide to create a water repellant barrier that will prevent spilling of liquid containing myofibers. Transfer the myofibers in the smallest volume possible to the microscopic glass slide and disperse them on the glass slide.
    NOTE: Avoid physical pulling of the myofibers over the microscopic glass slide since this might cause friction resulting in detachment of clusters from myofibers.
13. Remove the residual liquid with the small-bore Pasteur pipette or a 200 µL pipette.
14. Use two drops of aqueous mounting medium and cover the myofibers with a coverslip. Let the slides dry for the time recommended by the manufacturer. Store the slides at 4 °C in the dark.