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Method Article

Isolation of Quiescent Stem Cell Populations from Individual Skeletal Muscles

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DOI:

10.3791/64557

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December 9th, 2022

 ,  ,  ,  , 
1Department of Biomedicine, Aarhus University

* These authors contributed equally

In This Article

Summary

This protocol describes the isolation of muscle stem cells and fibro-adipogenic progenitors from individual skeletal muscles in mice. The protocol involves single muscle dissection, stem cell isolation by fluorescence activated cell sorting, purity assessment by immunofluorescence staining, and quantitative measurement of S-phase entry by 5-ethynyl-2´-deoxyuridine incorporation assay.

Abstract

Skeletal muscle harbors distinct populations of adult stem cells that contribute to the homeostasis and repair of the tissue. Skeletal muscle stem cells (MuSCs) have the ability to make new muscle, whereas fibro-adipogenic progenitors (FAPs) contribute to stromal supporting tissues and have the ability to make fibroblasts and adipocytes. Both MuSCs and FAPs reside in a state of prolonged reversible cell cycle exit, called quiescence. The quiescent state is key to their function. Quiescent stem cells are commonly purified from multiple muscle tissues pooled together in a single sample. However, recent studies have revealed distinct differences in the molecular profiles and quiescence depth of MuSCs isolated from different muscles. The present protocol describes the isolation and study of MuSCs and FAPs from individual skeletal muscles and presents strategies to perform molecular analysis of stem cell activation. It details how to isolate and digest muscles of different developmental origin, thicknesses, and functions, such as the diaphragm, triceps, gracilis, tibialis anterior (TA), gastrocnemius (GA), soleus, extensor digitorum longus (EDL), and the masseter muscles. MuSCs and FAPs are purified by fluorescence-activated cell sorting (FACS) and analyzed by immunofluorescence staining and 5-ethynyl-2´-deoxyuridine (EdU) incorporation assay.

Introduction

Skeletal muscle has a high capacity for regeneration due to the presence of muscle stem cells (MuSCs). MuSCs are located on the myofibers, underneath the basal lamina, and reside in a quiescent state of prolonged, reversible cell cycle exit1,2,3,4. Upon injury, MuSCs activate and enter the cell cycle to give rise to amplifying progenitors that can differentiate and fuse to form new myofibers2,5. Previous work has showed that MuSCs are absolutely essential for muscle regeneration6,7,8. Moreover, a single MuSC can engraft and generate both new stem cells and new myofibers9. Skeletal muscle also harbors a population of mesenchymal stromal cells called fibro-adipogenic progenitors (FAPs), which play a crucial role in supporting MuSC function during muscle regeneration6,10,11,12.

Due to their potential to coordinate muscle regeneration, there has been tremendous interest in understanding how MuSCs and FAPs work. Quiescent MuSCs are marked by expression of the transcription factors Pax7 and Sprouty1, and the cell surface protein calcitonin receptor, whereas quiescent FAPs are marked by the cell surface protein platelet-derived growth factor receptor alpha (PDGFRa)10,12,13,14,15. Previous studies have showed that MuSCs and FAPs could be purified from skeletal muscles using cell surface markers and fluorescence-activated cell sorting (FACS)9,15,16,17,18,19,20,21. While these protocols have greatly advanced the ability to study MuSCs and FAPs, one drawback is that most of these protocols require the isolation of MuSCs from a pool of different muscle tissues. Recent work from us and others have revealed differences in cell phenotype and gene expression levels between MuSCs isolated from different tissues22,23. MuSCs from the diaphragm, triceps, and gracilis show faster activation than MuSCs from lower hindlimb muscles22, while MuSCs from extraocular muscle show faster differentiation than MuSCs from the diaphragm and lower hindlimb muscles23.

This protocol describes the isolation of MuSCs and FAPs from individual skeletal muscles (Figure 1). This includes the dissection of the diaphragm, triceps, gracilis, tibialis anterior (TA), soleus, extensor digitorum longus (EDL), gastrocnemius (GA), and masseter muscles. Dissected muscles are subsequently dissociated by enzymatic digestion using collagenase II (a protease that specifically targets the Pro-X-Gly-Pro amino sequence in collagen, enabling the degradation of connective tissue and tissue dissociation24) and dispase (a protease that cleaves fibronectin and collagen IV, enabling further cell dissociation25). MuSCs and FAPs are isolated from single-cell suspensions by FACS. As examples of downstream assays for cell analysis, stem cell activation is determined by assaying 5-ethynyl-2´-deoxyuridine (EdU) incorporation, while cell purity is determined by immunofluorescence staining for the cell-type specific markers Pax7 and PDGFRa.

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Protocol

The present protocol was performed in accordance with animal care guidelines at Aarhus University and local ethics regulations.

NOTE: Make sure to comply with the regulations of the local ethical committee for animal experimentation and handling of post-mortem rodent samples. Mice are a potential source of allergens; if available, turn on exhaust ventilation and place it over the workspace to avoid excessive exposure to allergens. Alternatively, wear a face mask if the experiment is carried out regularly. This protocol involves working with sharps, and researchers are recommended to familiarize themselves with the procedures and logistics for applying first aid in the case of a cut.

1. Preparation (1-2 h; the day before dissection)

NOTE: The solutions, plates, and media are prepared under sterile conditions and filtered (0.45 µm) prior to use unless otherwise noted. Prepare stock solutions of dispase (11 U/mL in PBS) and collagenase II (1.000 U/mL in PBS) and store them at -20 °C (Table 1). The stocks are thawed and used for secondary digestion in step 4.2.6.

  1. Collagen coating of a 96-well half-area plate
    1. Prepare acidic water. Add 5.15 mL of glacial acetic acid to 895 mL of autoclaved water in a 2 L beaker.
    2. Filter the solution using a 0.45 µm, 500 mL filter unit. Transfer 800 mL of the filtered acidic water to a 1 L bottle.
    3. Add 40 mL of sterile collagen stock solution to the bottle. Gently mix the solution by swirling and store at 4 °C, protected from light, until use.
    4. Coat one or more 96-well half-area plates with collagen. Add 50 µL of collagen coating solution to each well and coat the plate overnight (ON) at 4 °C.
    5. Aspirate the collagen solution the following day using a vacuum-based aspirator and wash the plate 2x by adding 100 µL of autoclaved water and aspirating.
    6. Tilt the lid of the plate and let the plate dry in the hood for 20-30 min.
    7. When the plate is fully dry, wrap it in aluminum foil, and store at 4 °C until use. Coated plates can be stored for up to 4 weeks.
      ​NOTE: Collagen coating enables cell adhesion. It is important to wash away free collagen as it might interfere with cell function and signaling (e.g., binding of collagen V to the calcitonin receptors on the MuSCs)26.

2. Preparation (0.5 h; the day of dissection):

  1. Preparation of additional solutions and workspace
    1. Prepare sterile wash medium by adding 50 mL of horse serum and 5 mL of pen/strep to 445 mL of HAM's F10 medium. Work in a laminar flow hood to prevent contamination.
    2. Calculate and weigh the appropriate amount of collagenase II needed to prepare the dissociation buffer. (For each mouse, 26,000 units of collagenase II is used to make 40 mL of dissociation buffer at 650 U/mL collagenase II). Add the weighed powder to a 50 mL conical tube and store it on ice.
    3. Prepare two 10 cm dishes for muscle isolation. Pour 3 mL of wash medium (Table 1) into each Petri dish. Bring the 10 cm dishes outside the laminar flow hood for later use.
    4. Prepare the materials for muscle digestion. Based on the number of samples, label the appropriate number of 50 mL conical tubes (eight per mouse, one for each muscle) and leave them on the bench. Spray down the needles, 10 mL syringes, and 40 µm cell strainers with 70% ethanol (eight of each per mouse, one for each muscle) and bring them inside the laminar flow hood. Attach needles to the syringes.
    5. Prepare the materials for sorting. Label 5 mL round bottom tubes with cell strainer caps based on the number of samples. Cover the tubes with aluminum foil and leave them in the laminar flow hood.
    6. Corresponding to the number of samples and populations being sorted, prepare 1.5 mL microcentrifuge tubes with 500 µL of wash medium for cell collection (16 per mouse, one tube for each cell population per muscle tissue). Store these tubes on ice.

3. Muscle dissection (20-30 min)

NOTE: This section of the protocol is carried out in a non-sterile environment. The procedure can be carried out using one or several mice. However, one mouse is sufficient to prepare both samples for sorting and controls for setting up compensation and FACS gates.

  1. Initiating muscle isolation
    1. Prepare the non-sterile workspace for dissection and muscle isolation. Disinfect the workstation with 70% ethanol. Place a sterile protective disposable underpad on the workstation.
    2. Using a permanent marker, draw a box for each muscle on top of a Petri dish lid to later place the isolated muscle.
    3. Spray the surgical instruments with 70% ethanol.
    4. Euthanize the mouse by CO2 inhalation and/or cervical dislocation.
    5. Isolate the individual muscles (from one mouse at a time if using several mice). Spray the mouse with 70% ethanol to wet the fur and disinfect the skin. Place the mouse on the underpad with the abdomen facing up. An overview of all eight muscles and their respective anatomical location is shown in Figure 2A.
  2. Isolating the gracilis muscles
    1. Use a pair of scissors to make a 0.5 cm horizontal incision into the abdominal skin.
    2. Remove the skin: Using both hands, use the thumb and index finger to grab onto the upper and lower part of the incision. Pull at each side of the incision to tear and part the skin of the torso and lower extremities. Pull down the skin of the lower extremity to expose both hindlimbs (downward from hip to toes). Likewise, pull upward to expose the torso.
    3. Locate the gracilis muscle (inner thigh). Using curved forceps, grab onto the gracilis and slightly lift the muscle (Figure 2B). With scissors, make a 0.5 cm incision (Figure 2C), from which the gracilis is cut out and fully isolated (Figure 2D). Repeat this for the other leg to isolate the second gracilis muscle.
  3. Isolating the TA and EDL muscles (lower hindlimb, ventral side)
    1. Using a scalpel, cut the fascia by making a 0.5 cm incision along the lateral side of the tibia (the thickest lower hindlimb bone). Use curved forceps to grab the fascia and pull to remove it.
      NOTE: When the fascia has been fully removed, the tendons at the distal end of the hind leg are visible and can be separated.
    2. Use straight forceps with superfine tips to go in between the distal tendon of the TA (lateral side of the tibia) and the EDL (below the TA) (Figure 2E-G).
      NOTE: With experience, the blunt end of a scalpel blade can be used instead of the straight forceps.
    3. Slide the forceps toward the proximal end of the muscle to separate the muscles.
    4. Bring the straight forceps back to the distal end. Cut the distal tendon.
    5. Gently grab the distal tendon of the muscle with curved forceps. Lift the muscle up and over its proximal attachment and carefully cut the proximal tendon as close to its attachment point as possible. Cut the other end and transfer the TA to a Petri dish.
    6. Use straight forceps with superfine tips to go underneath the distal tendon of the EDL.
    7. Slide the forceps toward the proximal end of the muscle to separate the muscles.
    8. Bring the straight forceps back to the distal end. Cut the distal tendon without damaging the muscle.
    9. Gently grab the distal tendon of the EDL with curved forceps. Lift the muscle up and over its proximal attachment and carefully cut the proximal tendon as close to its attachment point as possible. Cut the other end and transfer the EDL to a Petri dish. Repeat for the other hindlimb to isolate the second TA and EDL muscles.
  4. Isolating the GA and soleus muscles (lower hindlimb, dorsal side)
    1. Use straight forceps with superfine tips to go in between the Achilles tendon and the lower hindlimb bones.
    2. Slide the forceps toward the proximal end of the muscle to separate the muscle.
    3. Bring the straight forceps back to the distal end. Cut the distal tendon.
      NOTE: In order to avoid damaging the soleus muscle, which lies underneath the GA, cut as close to its distal Achilles tendon attachment as possible, leaving a chunk of the tendon attached to the GA (Figure 2H).
    4. To reveal and isolate the soleus muscle, pull the GA up and over the fibula (the thinnest bone of the two bones in the lower hindlimb).
      NOTE: The soleus muscle is distinguished by its characteristic dark red color relative to the GA (Figure 2I).
    5. Locate the proximal soleus tendon. With straight forceps, go in between the soleus and GA.
    6. Move the forceps toward the distal end of the muscle to separate the soleus from the GA.
    7. Isolate the soleus first. Cut its proximal tendon, grab the tendon with curved forceps, and carefully lift the soleus to access its distal tendon. Cut the distal tendon to isolate the soleus from the GA. Place the soleus in the Petri dish with wash medium (Figure 2J)
    8. Cut the GA and place it in the Petri dish. Repeat this for the other leg to isolate the second soleus and GA muscles.
  5. Isolating the triceps muscles (upper forelimb, dorsal side)
    1. Use straight forceps with superfine tips to go in between the triceps muscle and the humerus bone (the main bone of the upper foreleg) (Figure 2K-M).
    2. Slide the forceps toward the proximal end of the muscle to separate the muscle. Cut the proximal end of the muscle.
    3. Grab the proximal end of the triceps with curved forceps and pull it up and over the elbow to access the distal tendon. Cut the distal tendon of the triceps, transfer the muscle into a Petri dish, and repeat the procedure for the other forelimb.
  6. Isolating the masseter muscles
    1. Remove the fur and skin from the jaw. Make a horizontal incision of 0.5 cm with scissors. Using both hands, pinch onto each side of the incision using the thumb and index fingers. Remove the skin by pulling upward and downward.
    2. Locate the major tendon of the masseter (caudal, below the eye). Insert the flat scalpel blade in between the bone and the muscle (Figure 2N). Cut the tendon.
    3. Grab the major masseter tendon with curved forceps. Cut it with a scalpel blade or scissors in the rostral direction to separate the masseter muscle from the jaw bone (Figure 2O, P). Place the isolated masseter muscle in the Petri dish. Repeat the procedure for the second masseter muscle.
  7. Isolating the diaphragm muscle
    NOTE: When isolating the diaphragm muscle, make sure to cut carefully to avoid cutting into the inner organs and intestine, as this is a source of contamination.
    1. Using scissors, make a thoracotomy (a cut in between the ribs) in the middle of the sternum (a long flat bone, situated in the middle of the chest, which connects the ribs) and cut through the sternum (Figure 2Q).
    2. Expose the diaphragm by cutting 360° through the ribcage.
    3. Separate the upper body from the lower part/abdomen. Using a pair of scissors, cut the trachea, esophagus, vena cava, and abdominal aorta.
    4. Separate the diaphragm from the lower body. Using scissors, make a laparotomy (a surgical incision into the abdominal cavity) 1 cm below the sternum and make a 360° cut (Figure 2R).
    5. Place the closed scissors between the ribcage and the abdominal organs and press down. Gently pull on the ribcage to separate it from the abdominal organs (Figure 2S).
    6. Separate the diaphragm from the rib cage. Loosely hold the diaphragm between two fingers and cut through the ribcage with scissors. Use scissors to cut the diaphragm as close to the ribs as possible in a 360° cut. Place the isolated diaphragm in a Petri dish.
      ​NOTE: During cervical dislocation, the diaphragm might rupture and collapse against the ribcage, making it difficult to locate. The muscle can still be isolated. Identify the collapsed muscle, hold it between two fingers, and cut 360° following the ribcage.

4. Muscle digestion to a single-cell suspension (~1 h 35 min)

NOTE: The following steps include non-sterile (steps 4.1-4.2) and sterile work environments (step 4.3).

  1. Mechanical digestion
    1. Place the isolated muscles in the lid of a 10 cm Petri dish.
    2. Using curved forceps, grab onto the isolated muscles one by one. For the soleus and GA, remove the remaining parts of the Achilles tendon.
    3. Using scissors, mince the isolated muscles one by one by cutting them into roughly 1 mm3 pieces.
  2. Enzymatic digestion
    1. Prepare the muscle dissociation buffer by adding 40 mL of cold wash medium to the weighed collagenase II powder (collagenase II: 650 U/mL in wash media).
      NOTE: Use freshly prepared muscle dissociation buffer to ensure optimal enzymatic activity.
    2. Transfer the minced muscles to a 15 mL conical tube containing 5 mL of dissociation buffer.
    3. Incubate the tube for 35 min in a 37 °C shaking water bath at 60 rpm.
    4. After incubation, add wash medium to a total volume of 15 mL. Spin the tube at 1,600 x g for 5 min at 4 °C.
    5. Aspirate the supernatant down to a 4 mL volume using a vacuum-based aspirator. To avoid disturbing the pellet, slowly aspirate from the top, and remove any floating fat.
    6. Thaw stock aliquots of dispase and collagenase II. Add 500 µL of the collagenase II solution (1,000 U/mL stock, -20 °C) to the remaining 4 mL sample. Next, add 500 µL of the dispase solution (11 U/mL stock, -20 °C).
      NOTE: Dispase can generate a precipitate. If this occurs, spin the dispase stock solution at 10,000 x g, 1 min before use. Transfer the now clear supernatant to the cell suspension without disturbing the pellet.
    7. Vortex the samples briefly to dissolve the pellet.
    8. Incubate the samples for 20 min in a 37 °C shaking water bath at 60 rpm.
      NOTE: Digesting a large number of muscle samples is time-consuming. For steps 4.3.1-4.3.6, estimate taking 2-5 min per sample. This step goes faster when performed in parallel by two or more researchers.
  3. Homogenization of samples
    1. Homogenize the cell suspension. Transfer the suspension from the 15 mL tube to a 50 mL tube. Use a 10 mL syringe with a 20G needle to resuspend the sample by pulling the sample up and down through the needle 5x.
      NOTE: If pieces of undigested muscle clog the needle, wipe them off on a paper tissue.
    2. Place a 40 µm cell strainer on a new 50 mL conical tube.
    3. Take up the full cell suspension in the syringe and strain the sample into a new 50 mL conical tube with a cell strainer on top. Strain the sample by dispensing the total volume directly on the filter.
    4. To retrieve all mononucleated cells, add 20 mL of the wash medium to the empty conical tube and pour this through the cell strainer into the 50 mL conical tube containing the strained sample. Retrieve the remaining volume under the cell strainer with a p1000 pipette.
    5. Discard the cell strainer and spin the sample at 1,600 x g for 5 min at 4 °C.
    6. By pipetting with a p1000 pipette, aspirate the supernatant without disturbing the pellet.
    7. By pipetting with a p1000 pipette, resuspend the diaphragm pellet in 500 µL of the wash medium and resuspend the other muscle samples in 300 µL of the wash medium each.
      ​NOTE: The diaphragm is the biggest of the muscles with the highest stem cell content and can therefore be used for the control stainings.

5. Staining and sorting (~40 min + 30 min sort/sample)

NOTE: Work in a sterile environment on ice for the following steps.

  1. Transfer a 50 μL aliquot of the diaphragm cell suspension to four new 2 mL microcentrifuge tubes for the control stains (50 µL each). Add 250 µL of the wash medium to each control tube to a final volume of 300 µL.
  2. Add 3 µL (1:100) of the fluorescence minus-one antibodies (FMO-control) to the three control staining tubes and leave one tube without antibodies (unstained control) (see Table 2).
  3. Add 3 µL (1:100) of the antibodies (VCAM1-PECy7, CD45-FITC, CD31-FITC, and SCA1-PacificBlue) to the remaining muscle single cell suspensions, including the remaining diaphragm sample.
  4. Incubate the cell suspensions for 15 min at 4 °C in a head-over-head shaker.
    NOTE: The control stainings are necessary to determine background fluorescence levels and set the gates for flow cytometry. Different antibodies can be used, but each will have a specific working concentration and incubation time that needs to be tested. If antibodies from different vendors are used, the concentration, and thus the dilution, may vary. A cell viability dye can be added to the staining mixture to eliminate any dead or dying cells during the sort.
  5. Spin the samples at 1,600 x g for 5 min at 4 °C. Discard the supernatant without disturbing the pellet by pipetting.
  6. By pipetting with a p1000 pipette, resuspend each sample in 800 µL of the wash medium.
  7. Filter the samples using a FACS tube with a cell strainer cap (40 µm) to remove any remaining cell clumps (or aggregates).
  8. Wash the cell strainer by adding an additional 800 µL of the wash medium.
  9. Keep the samples covered with aluminum foil on ice until analysis.
    NOTE: If at this point the cell suspension appears cloudy/dense due to the high concentration of cells, add an additional 800 µL of the wash medium to decrease the cell density.
  10. Start up the FACS with a 70 µm nozzle.
  11. Use the unstained and FMO controls to set the gating strategy: MuSCs are negative for CD45-FITC, CD31-FITC, and SCA1-PacBlue, and are positive for VCAM1-PECy7; The FAPs are negative for CD45-FITC, CD31-FITC, VCAM1-PECy7, and positive for SCA1-PacBlue.
  12. Sort the stained single-cell suspensions using two-way sort and collect the MuSCs and FAPs into separate collection tubes containing 500 µL of the wash medium.
    NOTE: Using FACS with four lasers (configuration: 70 µm nozzle, 405 nm, 488 nm, 561 nm, 633 nm), the chosen combination of fluorophores does not require compensation of fluorescence signal. However, if using a different flow cytometer or a different combination of fluorophores, additional single-stained control samples for compensation are recommended. From the smaller muscles (EDL, soleus, TA), yields of 3,000 MuSCs and 5,000 FAPs are to be expected. For the other larger muscles, yields of 20,000 MuSCs and FAPs are to be expected. Event counts listed by the FACS software may deviate from the actual number of viable cells in the collection tube. Cell numbers can be confirmed by cell counting using a hemocytometer.
  13. Spin down the sorted cells at 1,600 x g for 5 min at 4 °C. By pipetting, aspirate the supernatant without disturbing the cell pellet.
  14. Add 100 µL of the wash medium to each sample and carefully resuspend the cells without creating air bubbles. Transfer the 100 µL of the resuspended sample into a collagen-coated half-area 96-well plate. Plate 1,000-3,000 cells per well.
    NOTE: If the cells are not properly resuspended, the cells may stick together in clumps, confounding later microscopy analyses.
  15. Inspect the cells under the microscope and note their distribution, shape, and size.
  16. Incubate the plate at 37 °C, 5% CO2. The cells will adhere within 2 h.
    ​NOTE: It is recommended to confirm the purity of the collected cell populations, either by flow cytometry analysis (by loading an aliquot of the sorted sample on the sorter and recording a small number of events) or by antibody staining of the plated cells followed by microscopy (Pax7 is a defining marker for mouse MuSCs, PDGFRa is a defining marker for mouse FAPs). Starting at section 7 below is a protocol for the staining of cells for Pax7 and PDGFRa protein levels.

6. EdU incorporation assay

NOTE: Work under sterile conditions and use the chemical fume hood when handling paraformaldehyde (PFA) for the following steps. EdU is a nucleotide analog incorporated into the DNA as the cells go through the S-phase of the cell cycle. It is mutagenic at high concentrations. Always wear gloves when handling EdU. Check the local guidelines for handling EdU waste.

  1. EdU pulse (day 1)
    1. Prepare a working solution of 2x EdU in a fresh wash medium.
    2. Take the 96-well plate with the cells out of the incubator. Inspect the cells under the microscope to monitor confluence. Move into the laminar flow hood.
    3. Remove 50 µL of medium from the plate by pipetting. Add 50 µL of 2x EdU working solution so that the final volume in the well is 100 µL.
    4. Culture the cells in the presence of EdU for the intended period of time at 37 °C, 5% CO2.
      NOTE: The timing and duration of the EdU pulse can vary. On average, quiescent MuSCs take 2 days to fully activate and complete their first cell division19. EdU can be added to the sorted cells immediately after plating.
  2. Fixation (day 2)
    NOTE: PFA is a carcinogen. Always handle PFA with care. Familiarize with the local regulations for handling PFA and disposing of waste.
    1. Take the 96-well plate with the cells out of the incubator. Inspect the cells under the microscope and move the plate into the fume hood.
    2. By pipetting, remove the medium and fix the cells by adding 50 µL of 4% PFA to each well. Incubate the plate for 10 min at room temperature (RT) in a chemical fume hood.
    3. Remove the PFA with a pipette and discard it into an appropriate waste container.
    4. Wash the cells by adding 100 µL of PBS to all of the wells. By pipetting, aspirate the PBS and repeat the wash. Keep the cells in 100 µL of PBS.
      ​NOTE: To avoid washing out any cells, dispense the PBS to the side of the well by pipetting slowly.
  3. EdU labeling
    1. Permeabilize the cells prior to EdU detection by removing the PBS and adding 100 µL of 0.5% Triton X-100 in PBS. Incubate the plate for 5 min at 4 °C.
      NOTE: Triton X-100 is a surfactant and can cause skin irritation. Handle with care and always use gloves.
    2. After 5 min, aspirate the Triton X-100 and add 100 µL of PBS.
    3. Prepare an EdU click-chemistry reaction mix according to the manufacturer's protocol.
    4. Aspirate the PBS and add 33 µL of the EdU click-chemistry reaction mix. Incubate the plate for 30 min at RT.
    5. Aspirate the EdU click-chemistry reaction mix and wash the cells with 100 µL of PBS.
    6. Aspirate the PBS, add 100 µL of PBS with Hoechst (1:2,000), and incubate protected from light for 10 min at RT.
    7. Aspirate the Hoechst and wash twice by adding 100 µL of PBS. Store the cells in 100 µL of PBS at 4 °C in the dark.
    8. Image the cells on an inverted microscope. The cells can be stored in the dark for months as long as the wells do not dry out.
      ​NOTE: It is possible to co-stain the cells with antibodies. In this case, proceed with a blocking step followed by incubation with the primary antibody. Select a secondary antibody with a conjugated fluorophore whose emission spectrum does not overlap with that of the fluorophore used to detect EdU. Make sure not to use secondary antibodies with fluorophores overlapping with the spectral range of Hoechst.

7. Immunofluorescence staining

NOTE: This part of the protocol can be performed independently of section 6. When skipping section 6, please perform steps 6.3.1 and 6.3.2 to enable cell permeabilization before continuing with step 7.2 below.

  1. Take out the plate containing the EdU-labeled cells.
  2. Prepare 20 mL of blocking buffer by adding 2 mL of donkey serum to 18 mL of PBS.
  3. To prevent nonspecific antibody binding, remove 100 µL of the PBS from each well and add 50 µL of the blocking buffer with a p200 pipette. Incubate the plate for 30 min at RT, protected from light and covered in aluminum foil to prevent photobleaching of the fluorophore-labeled EdU.
    NOTE: The cells are fixed to the bottom of the well but can detach if too much force is applied when pipetting.
  4. While blocking the samples, prepare the primary antibody mix. Add 8.0 µL (1:100) of mouse anti-Pax7 and rabbit anti-PDGFRa to 800 µL of blocking buffer to stain 16 wells (eight tissues, two cell types).
  5. After blocking, remove the donkey serum and add 50 µL of the primary antibody mix to each well. Incubate the plate overnight at 4 °C, covered in aluminum foil.
  6. After incubation, remove the primary antibody and add 50 µL of Triton (0.5% in PBS) to each of the wells. Incubate the plate for 5 min at RT protected from light, to wash away the unbound antibody. Repeat the wash three times.
  7. Prepare the secondary antibody master mix by adding 1.0 µL (1:1,000) of donkey anti-mouse-Alexa647 and donkey anti-rabbit-Alexa555 to a microcentrifuge tube containing 1,000 µL of the blocking buffer.
  8. Add 33 µL of the secondary antibody master mix to each of the wells. Incubate the plate for 60 min at RT protected from light.
  9. Remove the excess secondary antibody by pipetting, add 50 µL of Triton (0.1% in PBS), and incubate for 5 min at RT protected from light. Repeat the wash three times.
  10. Finally, add 100 µL of PBS. Decide whether to image the plate immediately using an inverted fluorescence microscope with a cooled CCD camera, or store the plate at 4 °C until later analysis by sealing the plate edges with parafilm and wrapping the sealed plate in aluminum foil.

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Results

Following the protocol for individual skeletal muscle isolation (Figure 2), the gracilis, TA, EDL, GA, soleus, triceps, masseter, and diaphragm muscles were isolated from three Swiss male outbred mice that had been discontinued from a local breeding program (Figure 2). Following tissue dissociation and antibody staining, MuSCs and FAPs from the individual muscles were purified by FACS (Figure 3). The initial gating was obtained with...

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Discussion

Several steps are key in the execution of this protocol to achieve good yields. The individual muscles have a small volume compared to the amount of muscle used in bulk isolation protocols. This results in a risk of the muscle drying out during the dissection, which reduces yield. To prevent this, it is important to add medium to the muscles immediately after dissection. In addition, if dissection is taking longer, the skin can be removed from one limb at a time to reduce the time of exposure of the muscles to air. The s...

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Disclosures

The authors have no competing financial interests and no conflicts of interest.

Acknowledgements

Cell sorting was performed at the FACS Core Facility, Aarhus University, Denmark. Figures were created using Biorender.com. We thank Dr. J. Farup for sharing the rabbit anti-PDGFRa antibody. This work was supported by an AUFF Starting Grant to E.P. and Start Package grants from NovoNordiskFonden to E.P. (0071113) and to A.D.M. (0071116).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1.5 mL tube( PCR performance tested, PP, 30,000 xg, DNA/DNase-/RNase-free, Low DNA binding, Sterile )Sarstedt AG & Co. KG, Hounisen Laboratorieudstyr A/S72.706.7001.5 mL tube
15 mL tube (PP/HD-PE, 20,000 xg, IVD/CE, IATA, DNA/DNase-/RNase-free, Non-cytotoxic, pyrogen free, Sterile)Sarstedt AG & Co. KG, Hounisen Laboratorieudstyr A/S62.554.50215 mL tube
5 mL polystyrene round-bottom tubeFalcon, Fisher Scientific 352054FACS tube without strainer cap
5 mL polystyrene Round-bottom tube with cell-strainer capFalcon, Fisher Scientific  352235FACS tube with strainer cap
5 mL tube (PP, non sterile autoclavable)VWR collection525.09465 mL tube
50 mL tube( PP/HD-PE, 20,000 xg, IVD/CE, ADR, DNA/DNase-/RNase-free, non-cytotoxic, pyrogen free, Sterile)Sarstedt AG & Co. KG, Hounisen Laboratorieudstyr A/S62.547.25450 mL tube
Alexa Fluor 555 Donkey anti-rabbit IgG (H+L)Invitrogen, Thermo FisherLot: 2387458 (Cat # A31572)
Alexa Fluor 647 donkey-anti mouse IgG (H+L)Invitrogen, Thermo FisherLot: 2420713 (Cat#A31571)
ARIA 3BDFACS, Core facility Aarhus University
Centrifuge 5810eppendorfEP022628188Centrifuge
Click-iT EdU Cell Proliferation Kit for Imaging, Alexa Fluor 488 dyeInvitrogen, Thermo FisherLot: 2387287 (Cat# C10337)Cell Proliferation Kit
Collagen from calf-skin Bioreagent, Sigma Aldrich Source: SLCK6209 (Cat# C8919)
Collagenase type IIWorthington, Fisher Scientific Lot: 40H20248 (cat# L5004177 )Collagenase
DispaseGibco, Fisher Scientific Lot: 2309415 (cat# 17105-041 )Dispase
Donkey serum (non-sterile)Sigma Aldrich, MerckLot: 2826455 (Cat# S30-100mL)
Dumont nr. 5, 110 mmDumont, Hounisen Laboratorieudstyr A/S1606.327Straight forceps with fine tips
Dumont nr. 7, 115 mmDumont, Hounisen Laboratorieudstyr A/S1606.335Curved forceps
F-10 Nutrient mixture (Ham) (1x), +L-glutamineGibco, Fisher Scientific Lot. 2453614 (cat# 31550-023)
FITC anti-mouse CD31BioLegend, NordicBioSiteMEC13.3 (Cat # 102506)
FITC Anti-mouse CD45BioLegend, NordicBioSite30-F11 (Cat# 103108)
Glacial acetic acid (100%)EMSURE, Merck  K44104563 9Cat # 1000631000)
Head over head mini-tube rotator Fisher Scientific 15534080 (Model no. 88861052)Head over head mini-tube rotator
Horse serumGibco, Fisher Scientific Lot. 2482639 (cat# 10368902 )
Isotemp SWB 15FisherBrand, Fisher Scientific15325887Shaking water bath
MS2 mini-shaker IKA Vortex unit
Needle 20 G (0.9 mm x 25 mm)BD microlance, Fisher Scientific 30482720G needle 
Neutral formalin buffer 10%CellPath, Hounisen Laboratorieudstyr A/SLot: 03822014 (Cat # HOU/1000.1002)
Non-pyrogenic cell strainer (40 µM)Sarstedt AG & Co. KG, Hounisen Laboratorieudstyr A/S83.3945.040Cell strainer 
Pacific Blue anti-mouse Ly-6A/E (Sca-1)BioLegend, NordicBioSiteD7 (Cat# 108120)
Pax7 primary antibodyDSHBLot: 2/3/22-282ug/mL (Cat# AB 528428)
PBS 10x powder concentrateFisher BioReagents, Fisher ScientificBP665-1
PE/Cy7 anti-mouse CD106 (VCAM1)BioLegend, NordicBioSite429 (MVCAM.A) (Cat # 105720)
Pen/strepGibco, Fisher Scientific Lot. 163589 (cat# 11548876 )
Pipette tips p10Art tips, self sealing barrier, Thermo Scientific2140-05Low retention, pre-sterilized, filter tips
Pipette tips p1000Art tips, self sealing barrier, Thermo Scientific2279-05Low retention, pre-sterilized, filter tips
Pipette tips p20Art tips, self sealing barrier, Thermo Scientific2149P-05Low retention, pre-sterilized, filter tips
Pipette tips p200Art tips, self sealing barrier, Thermo Scientific2069-05Low retention, pre-sterilized, filter tips
Protective underpadAbena ACTC-7712 60 x 40cm, 8 layers
Rainin, pipet-lite XLSMettler Toledo, Thermo Scientific 2140-05, 2149P-05, 2279-05, 2069-05Pipettes (P10, P20, P200, P1000)
Recombinant anti-PDGFR-alphaRabMAb, abcamAB134123
Scalpel (shaft no. 3)Hounisen, Hounisen Laboratorieudstyr A/S1902.502Scalpel
Scalpel blade no. 11Heinz Herenz, Hounisen Laboratorieudstyr A/S1902.0911Scalpel
Scanlaf marsLabogeneclass 2 cabinet: MarsFlow bench
ScanROlympusMicroscope, Core facility Aarhus University
ScissorsFST14568-09
Series 8000 DHThermo Scientific3540-MARIncubator
Serological pipette 10 mLVWR612-3700Sterile, non-pyrogenic
Serological pipette 5 mLVWR, Avantor delivered by VWR612-3702Sterile, non-pyrogenic
Syringe 5 mL, Luer tip (6%), sterile BD Emerald, Fisher Scientific307731Syringe
TC Dish 100, standardSarstedt AG & Co. KG, Hounisen Laboratorieudstyr A/S83.3902Petri dish 
Tissue Culture (TC)-treated surface, black polystyrene, flat bottom, sterile, lid, pack of 20Corning, Sigma Aldrich376496-well Half bottom plate
Triton X-100Sigma Aldrich, MerckSource: SLCJ6163 (Cat # T8787)

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