Method Article

FACS-Isolation and Culture of Fibro-Adipogenic Progenitors and Muscle Stem Cells from Unperturbed and Injured Mouse Skeletal Muscle

DOI:

10.3791/63983

⸱

June 8th, 2022

 ,  , 
1Laboratory of Muscle Stem Cells and Gene Regulation, National Institute of Arthritis, Musculoskeletal, and Skin Diseases (NIAMS), National Institutes of Health (NIH), 2Flow Cytometry Section, National Institute of Arthritis, Musculoskeletal, and Skin Diseases (NIAMS), National Institutes of Health (NIH)

In This Article

Summary

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

The precise identification of fibro-adipogenic progenitor cells (FAPs) and muscle stem cells (MuSCs) is critical to studying their biological function in physiological and pathological conditions. This protocol provides guidelines for the isolation, purification, and culture of FAPs and MuSCs from adult mouse muscles.

Abstract

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Fibro-adipogenic progenitor cells (FAPs) are a population of skeletal muscle-resident mesenchymal stromal cells (MSCs) capable of differentiating along fibrogenic, adipogenic, osteogenic, or chondrogenic lineage. Together with muscle stem cells (MuSCs), FAPs play a critical role in muscle homeostasis, repair, and regeneration, while actively maintaining and remodeling the extracellular matrix (ECM). In pathological conditions, such as chronic damage and muscular dystrophies, FAPs undergo aberrant activation and differentiate into collagen-producing fibroblasts and adipocytes, leading to fibrosis and intramuscular fatty infiltration. Thus, FAPs play a dual role in muscle regeneration, either by sustaining MuSC turnover and promoting tissue repair or contributing to fibrotic scar formation and ectopic fat infiltrates, which compromise the integrity and function of the skeletal muscle tissue. A proper purification of FAPs and MuSCs is a prerequisite for understanding the biological role of these cells in physiological as well as in pathological conditions. Here, we describe a standardized method for the simultaneous isolation of FAPs and MuSCs from limb muscles of adult mice using fluorescence-activated cell sorting (FACS). The protocol describes in detail the mechanical and enzymatic dissociation of mononucleated cells from whole limb muscles and injured tibialis anterior (TA) muscles. FAPs and MuSCs are subsequently isolated using a semi-automated cell sorter to obtain pure cell populations. We additionally describe an optimized method for culturing quiescent and activated FAPs and MuSCs, either alone or in coculture conditions.

Introduction

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

The skeletal muscle is the largest tissue in the body, accounting for ~40% of adult human weight, and is responsible for maintaining posture, generating movement, regulating basal energy metabolism, and body temperature1. Skeletal muscle is a highly dynamic tissue and possesses a remarkable ability to adapt to a variety of stimuli, such as mechanical stress, metabolic alterations, and daily environmental factors. In addition, skeletal muscle regenerates in response to acute injury, leading to complete restoration of its morphology and functions2. Skeletal muscle plasticity mainly relies upon a population of resident muscle stem cells (MuSCs), also termed satellite cells, which are located between the myofiber plasma membrane and the basal lamina2,3. Under normal conditions, MuSCs reside in the muscle niche in a quiescent state, with only a few divisions to compensate for cellular turnover and to replenish the stem cell pool4. In response to injury, MuSCs enter the cell cycle, proliferate, and either contribute to the formation of new muscle fibers or return to the niche in a self-renewal process2,3. In addition to MuSCs, homeostatic maintenance and regeneration of the skeletal muscle rely upon the support of a population of muscle resident cells named fibro-adipogenic progenitors (FAPs)5,6,7. FAPs are mesenchymal stromal cells embedded in the muscle connective tissue and capable of differentiating along fibrogenic, adipogenic, osteogenic, or chondrogenic lineage5,8,9,10. FAPs provide structural support for MuSCs as they are a source of extracellular matrix proteins in the muscle stem cell niche. FAPs also promote long-term maintenance of the skeletal muscle by secreting cytokines and growth factors that provide trophic support for myogenesis and muscle growth6,11. Upon acute muscle injury, FAPs rapidly proliferate to produce a transient niche that supports the structural integrity of the regenerating muscle and provides a favorable environment to sustain MuSCs proliferation and differentiation in a paracrine manner5. As regeneration proceeds, FAPs are cleared from the regenerative muscle by apoptosis, and their numbers gradually return to basal level12. However, in conditions favoring chronic muscle injury, FAPs override pro-apoptotic signaling and accumulate in the muscle niche, where they differentiate into collagen-producing fibroblasts and adipocytes, leading to ectopic fat infiltrates and fibrotic scar formation12,13.

Due to their multipotency and their regenerative abilities, FAPs and MuSCs have been identified as prospective targets in regenerative medicine for the treatment of skeletal muscle disorders. Therefore, to investigate their function and therapeutic potential, it is important to establish efficient and reproducible protocols for the isolation and culture of FAPs and MuSCs.

Fluorescence-activated cell sorting (FACS) can identify different cell populations based on morphological characteristics such as size and granularity, and permits cell-specific isolation based on the use of antibodies directed against cell surface markers. In adult mice, MuSCs express the vascular cell adhesion molecule 1 (VCAM-1, also known as CD106)14,15 and α7-Integrin15, while FAPs express the platelet-derived growth factor receptor α (PDGFRα) and the stem cell antigen 1 (Sca1 or Ly6A/E)5,6,9,12,16,17. In the protocol described here, MuSCs were identified as CD31-/CD45-/Sca1-/VCAM-1+/α7-Integrin+, while FAPs were identified as CD31-/CD45-/Sca1+/VCAM-1-/α7-Integrin-. Alternatively, PDGFRαEGFP mice were employed to isolate FAPs as CD31-/CD45-/PDGFRα+/VCAM-1-/α7-Integrin- events18,19. Furthermore, we compared the overlapping between the fluorescent signal of PDGFRα-GFP+ cells to cells identified by the surface marker Sca1. Our analysis showed that all GFP-expressing cells were also positive for Sca1, indicating that either approach can be employed for the identification and isolation of FAPs. Finally, staining with specific marker antibodies confirmed the purity of each cell population.

Access restricted. Please log in or start a trial to view this content.

Protocol

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

All animal experiments performed were conducted in compliance with institutional guidelines approved by the Animal Care and Use Committee (ACUC) of the National Institute of Arthritis, Musculoskeletal, and Skin Diseases (NIAMS). Investigators performing this protocol must adhere to their local animal ethics guidelines.

NOTE: This protocol describes in detail how to isolate FAPs and MuSCs from hind limb and injured tibialis anterior (TA) muscles of adult male and female mice (3-6 months) and provides guidelines for coculturing FAPs and MuSCs. An overview of the experimental procedure is shown in Figure 1. All steps of this protocol should be performed in sterile conditions and at room temperature (RT) unless otherwise specified.

1. Reagent setup

  1. Wash Medium (WM): Prepare this solution by adding 10% (vol/vol) horse serum and 1x penicillin/streptomycin to Ham's F-10 Nutrient Mix cell media. WM can be prepared in advance and stored at 4 °C.
  2. Muscle Dissociation Buffer (MDB): Prepare this buffer by dissolving Collagenase II in WM. If collecting whole hind limb muscles, dissolve 1000 U/mL Collagenase II in 10 mL of WM for each mouse (to be prepared into a 50 mL conical tube). If collecting TA muscles, dissolve 800 U/mL Collagenase II in 7 mL of WM for each mouse (to be prepared into a 15 mL conical tube). For TA muscles, this will help to better visualize cell pellets and obtain a higher yield of FAPs and MuSCs. Prepare MDB fresh before collecting the muscles and keep it on ice until needed.
  3. Collagenase II solution stock: Prepare this solution by dissolving Collagenase II in sterile 1x phosphate-buffered saline (PBS) to a final concentration of 1000 U/mL. Filter the solution through a 0.45 μm syringe filter and aliquot into 1 mL stocks. Store the solution at -20 °C.
  4. Dispase solution stock: prepare this solution by dissolving Dispase in sterile 1x PBS to a final concentration of 11 U/mL. Filter the solution through a 0.45 μm syringe filter and aliquot into 1 mL stocks. Store the solution at -20 °C.
  5. Prepare FAP's culture medium by supplementing DMEM with 10% (vol/vol) fetal bovine serum, 1x penicillin/streptomycin, and 2.5 ng/mL FGF. Store the medium at 4 °C.
  6. Prepare MuSC's culture medium supplementing F10 with 10% (vol/vol) horse serum, 1x penicillin/streptomycin, and 2.5 ng/mL FGF. Store the medium at 4 °C.
  7. Prepare the coculture medium by supplementing DMEM with 10% (vol/vol) fetal bovine serum, 10% (vol/vol) horse serum, 1x penicillin/streptomycin, and 2.5 ng/mL FGF. Store the medium at 4 °C.

2. Hind limb muscle harvesting

  1. Add 2-3 mL of WM in a 6 cm dish (one dish per mouse) and place it on ice.
  2. Perform euthanasia by asphyxiation: place the mouse in a CO2 chamber and introduce 100% CO2. Use cervical dislocation to confirm death.
  3. Place the mouse supine on a dissection pad and spray it with 70% (vol/vol) ethanol to avoid contamination.
  4. Use forceps to pinch the center of the mouse belly skin and cut a ~1 cm opening horizontally. Grasp the wound edges and deskin the mouse by pulling the opening in opposite directions to uncover the muscles underneath. Expose one side of the mouse's hind limb at the time.
  5. Before collecting muscles, remove intermuscular fat between the hamstrings and the proximal end of the quadriceps. This will improve the isolation of FAPs and MuSCs.
  6. Without damaging the tissue, use sharp forceps to break and peel off the fascia to expose the underneath TA and extensor digitorum longus (EDL) muscles. Run the sharp tip of the forceps underneath the TA/EDL muscles to detach the muscles from the tibia.
  7. Cut off the tendons attaching the TA/EDL to the ankle and, while holding it with forceps, trim the muscles with scissors along the longitudinal line of the TA/EDL. Cut the tendons around the knee to detach the whole TA/EDL muscles. Place the muscles in a 6 cm dish kept on ice.
  8. Proceed to isolate the gastrocnemius and soleus muscles by cutting all tendons at the ankle and detaching the muscles from the tibia and fibula. Cut around the knee and transfer the muscles to the 6 cm dish.
  9. Peel off the fascia around the quadriceps and separate it from the femur by running the sharp tip of the forceps between the muscle and the bone. Cut the tendons around the knee and, while holding the quadriceps with forceps at the distal end, trim the rest of the muscle by cutting along the femur. Place the muscles in the 6 cm dish.
  10. Detach the hamstrings and remaining muscles around the femur and transfer those to the 6 cm dish. Collect the hind limb muscles in a 6 cm dish kept on ice.
    NOTE: Avoid damaging blood vessels, when possible, to prevent formation of blood clumps that could interfere with the downstream isolation. If bleeding occurs, blot the excised vessel immediately with a sterile gauze to absorb the blood.
  11. Collect TA, EDL, gastrocnemius, soleus, quadricep, and hamstring muscles from the contralateral limb.
    ​NOTE: When isolating the whole hind limb muscles, do not pool two or more mice. If isolating TA muscles, up to three TA muscles can be pooled.

3. Mechanical and enzymatic muscle digestion

  1. Aspirate WM from the 6 cm dish and add 1-2 mL of MDB to keep the muscles moist.
  2. Mince the muscles thoroughly until obtaining a slurry paste of well-minced tissue. To do so, use forceps to hold one end of a piece of muscle, then use a scalpel to tear and slice the muscle until less tight.
  3. Using scissors, keep cutting the muscle into small pieces for 1-2 min. Repeat this step for each group of muscles until obtaining a well-minced muscle sludge.
    NOTE: This is a very critical step to maximize the yield of FAPs and MuSCs. It is recommended to spend 8-10 min (4-5 min if isolating TA muscles only) in this step to ensure a proper muscle mincing.
  4. Transfer the minced muscles from each mouse to the conical tube containing MDB.
  5. Seal the tubes with laboratory film and incubate in a 37 °C water bath with agitation (75 rpm) for 1 h. Place the tubes horizontally along the shaking path and use weights to keep the tubes submerged in water.
  6. Thaw 1 mL of Collagenase II stock and 1 mL of dispase stock per mouse. Before use, spin the dispase stock in a swinging bucket rotor at 10,000 x g for 1 min at 4 °C. Use only the supernatant.
  7. If whole hind limb muscles were collected, proceed as follows:
    1. After 1 h, remove the tube from the shaker and fill it to 50 mL with WM. Gently invert a couple of times to ensure mixing.
    2. Centrifuge the cells in a swinging bucket rotor at 250 x g for 5 min at 4 °C. Transfer 42 mL of supernatant into two new tubes (~21 mL in each tube) and leave ~8 mL in the original tube.
      1. Fill the new tubes containing 21 mL of the supernatant up to 50 mL with WM. Centrifuge the cells again in a swinging bucket rotor at 350 x g for 8 min at 4 °C. Aspirate all the supernatant and keep the pellets on ice.
    3. Add 1 mL of collagenase II stock and 1 mL of dispase stock in the original tube containing ~8 mL of MDB.
    4. Using a 5 mL serological pipette, resuspend the pellet 10 times without clogging. Eject the solution toward the wall of the tube, avoiding the formation of bubbles. If clogging occurs during resuspension, push the tip of the pipette against the bottom of the tube to mechanically disrupt the muscle chunks. Proceed to step 3.9.
  8. If TA muscles were collected, proceed as follows:
    1. After 1 h, remove the tube from the shaker and fill them to 15 mL with WM. Gently invert a couple of times to ensure mixing.
    2. Centrifuge the cells in a swinging bucket rotor at 250 x g for 5 min at 4 °C. Transfer 13 mL of supernatant into one new 15 mL tube and leave ~2 mL in the original tube.
      1. Fill the new tube containing 13 mL of the supernatant up to 15 mL with WM. Centrifuge the cells again in a swinging bucket rotor at 350 x g for 8 min at 4 °C. Aspirate all the supernatant and keep the pellet on ice.
    3. Using a 1000 µL pipette tip, resuspend the pellet in the original tube up and down without clogging.
    4. Add 1 mL of Collagenase II stock and 1 mL of dispase stock in the original tube containing 2 mL of MDB and fill up to 10 mL with WM. Proceed to step 3.9.
  9. Seal the tubes with laboratory film and incubate in a 37 °C water bath with agitation (75 rpm) for 30 min. Place the tubes as in step 3.5.

4. Generation of mononucleated cells

NOTE: If working with TA muscles collected in a 15 mL conical tube, transfer the suspension into a 50 mL conical tube before proceeding with step 4.1.

  1. After 30 min, remove the tube from the shaker. Aspirate and eject the muscle suspension through a 10 mL syringe with a 20 G needle successfully 10 times.
    NOTE: Eject muscle suspension toward the wall of the tube to avoid bubbles and foaming. Small pieces of undigested tendons or cartilages might clog the needle during the first rounds of aspiration. If clogging occurs, use sterile forceps to remove the clog from the tip of the needle.
  2. Fill each tube up to 50 mL with WM and gently invert a couple of times to ensure mixing.
  3. Centrifuge the cells in a swinging bucket rotor at 250 x g for 5 min at 4 °C. Transfer supernatant into a new tube (~42 mL) and leave ~8 mL in the original tube.
    1. Fill the new tube containing 42 mL of the supernatant up to 50 mL with WM. Centrifuge the cells again in a swinging bucket rotor at 350 x g for 8 min at 4 °C. Aspirate all the supernatant and keep the pellet on ice.
  4. Place a 40 μm nylon cell strainer in the original 50 mL conical tube containing 8 mL of WM and pre-wet the cell strainer with 1-2 mL of WM.
  5. While holding the 40 μm nylon cell strainer, use a 10 mL pipette to gently resuspend the pellet 5-10 times. Filter the pellets through the cell strainer back into the same tube to minimize cell loss.
  6. At this step, ensure that there are additional tubes with cell pellets on ice. Filter those pellets back into the original tube by adding 4-5 mL of WM to each tube to resuspend the pellets and transfer the solution to the cell strainer positioned in the original tube. Allow filtering by gravity.
  7. After collecting all the pellets into the original 50 mL conical tube, rinse the cell strainer with another 4-5 mL of WM. Use a 1000 μL pipette to collect all the liquid from the underside of the cell strainer.
  8. Fill each tube with WM up to 50 mL and gently invert a couple of times to ensure mixing.
  9. Centrifuge the cells in a swinging bucket rotor at 250 x g for 5 min at 4 °C. Immediately aspirate all the supernatant after centrifugation without disturbing the pellet.
  10. Resuspend the pellet in 600 μL of WM and transfer it to a 2 mL microcentrifuge tube.

5. Antibody staining for flow cytometry

NOTE: For each experiment, set up the following controls: i) unstained control, ii) viability control to select for the live cell population, iii) single stained compensation controls to correct for fluorochrome emission spillover, and iv) fluorescence minus one (FMO) controls to set gating boundaries by accounting for spillover spread. Refer to Table 1 for a full list of staining controls.

  1. To prepare unstained control, transfer 10 μL of the cell suspension into a 2 mL microcentrifuge tube containing 190 μL of WM. Resuspend the cell suspension and filter through a 5 mL polystyrene round-bottom tube with a 35 μm cell-strainer cap. Allow the suspension to filter by gravity.
  2. Use a 200 μL pipette to collect all the liquid from the underside of the cell strainer. Seal with a cap and leave it on ice, protected from light.
  3. Prepare viability, FMO, and single stained controls: label 10 2 mL microcentrifuge tubes and add 190 μL of WM into each tube and 10 μL of cell suspension. Refer to Table 1 for a full list of controls. Add antibodies to the appropriate tube depending on the experimental control. Refer to Table 1 for information regarding antibody combination and concentration.
  4. Transfer the rest of the cell suspension (500 μL) into a 2 mL microcentrifuge tube (experimental tube) and add the following antibodies: CD31-APC, CD45-APC, Sca1-Pacific Blue, VCAM-1-biotin, and α7-Integrin-PE. Refer to Table 1 for information regarding antibody combination and concentration.
  5. Gently mix well each tube to ensure uniform distribution and incubate the cells in a rotating shaker at 4 °C for 45 min protected from light.
  6. After 45 min, fill all microcentrifuge tubes up to 2 mL with WM. Gently invert the tubes a couple of times to ensure complete mixing. Centrifuge the tubes in a refrigerated centrifuge with a fixed angle rotor at 250 x g for 5 min at 4 °C. Aspirate the supernatant without disturbing the pellet.
  7. Resuspend the cells in all microcentrifuge tubes in 300 μL of WM.
  8. Add streptavidin antibody into appropriate tubes. Refer to Table 1 for information regarding antibody combination and concentration. Gently mix well each tube to ensure uniform distribution and incubate the cells in a rotating shaker at 4 °C for 20 min protected from light. Leave the remaining tubes on ice, protected from the light.
  9. After 20 min, fill microcentrifuge tubes containing streptavidin antibody to 2 mL with WM. Gently invert the tubes a few times to ensure complete mixing. Centrifuge the tubes in a refrigerated centrifuge with a fixed angle rotor at 250 x g for 5 min at 4 °C. Aspirate the supernatant completely and resuspend cells in 300 μL of WM. 
  10. Pre-wet the 35 μm cell strainer cap of 10 5 mL polystyrene round-bottom tubes with 200 μL of WM. Filter cells from control tubes through the appropriate 5 mL polystyrene round-bottom tubes. Use a 200 μL pipette to collect all the liquid from the underside of the cell strainer. Seal with caps, leave the tubes on ice, and protect them from light.
  11. Resuspend the cells in the experimental tube with an additional 200 μL of WM for a total of 500 μL of WM. Pre-wet a cell strainer cap of a 5 mL polystyrene round-bottom tube with 200 μL of WM. Transfer the cell suspension from the experimental tube into the 5 mL polystyrene round-bottom tube and allow it to filter by gravity.
  12. Rinse the 2 mL microcentrifuge tube containing the experimental sample suspension with 300 μL of WM and pass it through the same strainer cap.Use a 200 μL pipette to collect all the liquid from the underside of the cell strainer. Seal with a cap, leave tubes on ice, and protect them from light.
    NOTE: If clogging of the 35 μm cell-strainer cap occurs, gently tap the tube on the bench to facilitate flow-through.
  13. Prepare and label the collection tubes for FAPs and MuSCs. If isolating cells from whole hind limb muscles, sort the cells in 5 mL polypropylene round-bottom tubes with up to 1 mL of either FAPs or MuSCs culture medium supplemented with 2x serum. If isolating cells from TA muscles, sort FAPs and MuSCs in 2 mL microcentrifuge tubes containing up to 400 μL of culture medium supplemented with 2x serum.

6. Fluorescence-activated cell sorting (FACS)

NOTE: This protocol employs a compact benchtop research flow cytometer equipped with a 100 μm nozzle and featuring a three-laser configuration (488 nm, 640 nm, 405 nm) with the capability to analyze up to nine different fluorochromes (11 parameters including the forward and side scatter). The fluorochromes used in this protocol and their associated detector bandpass filters are as follows: PE 586/42; PE-Cy7 783/56; APC 660/10; Pacific Blue 448/45; 7-Aminoactinomycin D (7-AAD) 700/54, GFP 527/32. Cells are sorted at 4 °C and remain on ice following the sort. Before operating this instrument, ensure that the user is properly trained by a technical applications specialist.

  1. Set up and performance check the cell sorter according to the manufacturer's specifications.
  2. Set up a hierarchical gating strategy to identify FAPs and MuSCs (Figure 2).
    1. Isolate FAPs based on the following scheme: i) forward cell scatter area vs side cell scatter area (FSC-A vs. SSC-A) to separate cells versus debris, ii) side cell scatter height vs side cell scatter width (SSC-H vs. SSC-W) to discriminate singlets from doublets in the SSC range, iii) forward cell scatter height vs forward cell scatter width (FSC-H vs. FSC-W) to discriminate singlets from doublets in the FSC range, and iv) 7-AAD area vs SSC-A to distinguish live versus dead cells.
      1. If isolating FAPs through the antibody-based method, use the following scheme: v) APC-CD45/CD31 area vs Pacific Blue-Ly-6A/E (Sca1) area to exclude CD31+ and CD45+ cells from further analysis, and vi) PE-Cy7-VCAM-1 area vs PE-α7-Integrin area from the CD31-/CD45-/Sca1+ population to distinguish FAPs. FAPs are identified as CD31-/CD45-/Sca1+/VCAM-1-/α7-Integrin- events.
      2. If isolating FAPs through endogenous GFP reporter method, use the following scheme: v) APC-CD45/CD31 area vs GFP-PDGFRα area to exclude CD31+ and CD45+ cells from further analysis and isolate GFP+ cells. FAPs are identified as CD31-/CD45-/PDGFRα+/VCAM-1-/α7-Integrin- events.
    2. Isolate MuSCs based on the following scheme: i) forward cell scatter area vs side cell scatter area (FSC-A vs. SSC-A) to separate cells versus debris, ii) side cell scatter height vs side cell scatter width (SSC-H vs. SSC-W) to discriminate singlets from doublets in the SSC range, iii) forward cell scatter height vs forward cell scatter width (FSC-H vs. FSC-W) to discriminate singlets from doublets in the FSC range, iv) 7-AAD area vs SSC-A to distinguish live vs dead cells, v) APC-CD45/CD31 area vs Pacific Blue-Ly-6A/E (Sca1) area to exclude CD31+ and CD45+ cells from further analysis, and vi) PE-Cy7-VCAM-1 area vs PE-α7-Integrin area from the CD31-/CD45-/Sca1- population to distinguish MuSCs. MuSCs are identified as CD31-/CD45-/Sca1-/VCAM-1+/α7-Integrin+ events.
  3. Run the unstained control and viability control to ensure that the cell population is properly positioned in the SSC-A vs FSC-A plot and to properly gate on live single cells.
  4. Acquire all single stained controls and generate the spillover compensation matrix.
  5. Run all FMO controls and determine the cut-off point between background fluorescence spread and the positively stained population.
  6. Approximately 5-10 min before acquisition of the experimental sample, add 7-AAD viability dye and mix gently.
  7. Once all controls have been processed, set the sort gates in accordance with the FMO controls; acquire and sort the experimental samples.
  8. After all samples have been processed, clean the cytometer according to the manufacturer's specification. Export all .fcs data for analysis.

7. Culture of FAPs and MuSCs

NOTE: Sorted cells should be cultured immediately after sorting, in an appropriate medium on collagen I coated plates.

  1. Centrifuge the collection tubes in a fixed angle rotor at 250 x g for 5 min at 4 °C.
  2. Aspirate the supernatant without disturbing the cell pellet.
  3. For MuSC's culture, resuspend the cells in an appropriate volume of MuSC culture medium and incubate them in standard conditions at 37 °C and 5% CO2.
    1. Plate freshly isolated MuSCs at a density of 20,000 cells/cm2 and activated MuSCs at a density of 15,000 cells/cm2.
    2. After plating, cells appear small, spherical, and translucent. Within 36 h, MuSCs adhere to the surface of the plate and slowly increase their size for the first 48 h.
    3. Replace the medium every 2 days.
      NOTE: MuSCs are very sensitive to cell density. To keep MuSCs in their proliferating state, grow them until they are 60%-70% confluent. Passage the cells into new collagen I coated dishes or plates and add new fresh medium every 2 days. If MuSCs are kept in the same dish or plate for longer than 3-4 days, they will acquire an elongated form and will align with neighboring cells until fusing together.
  4. For FAP's culture, resuspend the cells in an appropriate volume of MuSC culture medium and incubate them in standard conditions at 37 °C and 5% CO2.
    1. Plate the FAPs isolated from unperturbed muscle at a density of 15,000 cells/cm2 and FAPs isolated from injured muscle at 9,000 cells/cm2.
      NOTE: After plating, FAPs completely adhere to the surface of the plate within 48 h, while activated FAPs attach to the plate within a few hours. Once they are attached, FAPs acquire their characteristic shape, with a small cell body and elongated cell processes, and they rapidly proliferate.
    2. Replace the medium every 2 days.
      NOTE: FAPs are very sensitive to cell density. Seeding FAPs at low densities can lead to poor cell growth and cell death. On the contrary, plating FAPs at high seeding density will boost cell survival and improve their expansion. FAPs deriving from damaged muscle usually require lower cell density than undamaged muscle, as they are already activated. It is recommended to adjust FAP plating density based on one's experimental conditions and on the source of the samples.
  5. For coculture, resuspend FAPs and MuSCs in the coculture medium at a ratio of 1:1 and incubate those in standard conditions at 37 °C and 5% CO2. Allow the cells to attach for 48 h and replace the medium every 2 days.

8. Immunofluorescence analysis of cultured FAPs and MuSCs

  1. Aspirate the cell medium and perform two or three washes with 1x PBS.
  2. Fix the cells with 2% paraformaldehyde (PFA) in 1x PBS for 15 min at RT.
  3. Aspirate 2% PFA and quickly wash the cells two or three times with 1x PBS.
  4. Perform cell permeabilization and blocking:
    1. For immunostaining of freshly isolated FAPs, perform cell permeabilization and blocking by incubating cells with 1x PBS + 0.1% Triton X-100 (PBST) + 1% bovine serum albumin (BSA) + 5% Normal Donkey Serum (NDS) for 30 min at RT.
    2. For immunostaining of freshly isolated MuSCs, perform cell permeabilization and blocking by incubating cells with PBST + 1% BSA + 5% Normal Goat Serum (NGS) for 30 min at RT.
  5. Apply primary antibodies:
    1. For immunostaining of freshly isolated FAPs, apply goat anti PDGFRα primary antibody (1:300) diluted in PBST + 1% BSA + 5% NDS overnight at 4 °C.
    2. For immunostaining of freshly isolated MuSCs, apply mouse anti Pax7 primary antibody (1:10) diluted in PBST + 1% BSA + 5% NGS for 45 min at RT.
  6. Wash the cells two or three times with 1x PBS to remove the primary antibody solution.
  7. Apply secondary antibodies:
    1. For immunostaining of freshly isolated FAPs, apply donkey anti-goat IgG (H+L) Alexa Fluor 488 secondary antibody (1:500) diluted in PBST + 1% BSA for 1 h at RT.
    2. For immunostaining of freshly isolated MuSCs, apply goat anti-mouse IgG1 Alexa Fluor 555 secondary antibody (1:500) diluted in PBST + 1% BSA for 45 min at RT.
  8. Wash the cells two or three times with 1x PBS to remove the primary antibody solution.
  9. Perform nuclear counter-staining by incubating cells with DAPI in PBST (1:1000) for 5 min at RT.
  10. Wash the cells three times with 1x PBS to remove the DAPI solution.
    CAUTION: DAPI is toxic and hazardous; handle it with care and dispose it in the hazardous waste bottle.
  11. Store the cells at 4 °C protected from the light.

Access restricted. Please log in or start a trial to view this content.

Results

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

This protocol allows the isolation of approximately one million FAPs and up to 350,000 MuSCs from uninjured hind limbs of wild-type adult mice (3-6 months), corresponding to a yield of 8% for FAPs and 3% for MuSCs of total events. When sorting cells from damaged TA 7 days post-injury, two to three TA muscles are pooled to obtain up to 300,000 FAPs and 120,000 MuSCs, which correspond to a yield of 11% and 4%, respectively. Post-sort purity values are usually above 95% for FAPs and MuSCs.

The ga...

Access restricted. Please log in or start a trial to view this content.

Discussion

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Establishing efficient and reproducible protocols for the identification and isolation of pure adult stem cell populations is the first and most critical step toward understanding their function. Isolated FAPs and MuSCs can be used to conduct multiomics analysis in transplantation experiments as a potential treatment for muscular diseases or can be genetically modified for disease modeling in stem cell therapy.

The protocol described here provides standardized guidelines for the identificatio...

Access restricted. Please log in or start a trial to view this content.

Acknowledgements

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

We would like to thank Tom Cheung (The Hong Kong University of Science & Technology) for advice on MuSC isolation. This work was funded by the NIAMS-IRP through NIH grants AR041126 and AR041164.

Access restricted. Please log in or start a trial to view this content.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
5 mL Polypropylene Round-Bottom TubeFalcon352063
5 mL Polystyrene Round-Bottom Tube with Cell-Strainer CapFalcon352235
20 G BD Needle 1 in. single use, sterileBD Biosciences 305175
anti-Alpha 7 Integrin PE (clone:R2F2) (RatIgG2b)The University of British Columbia53-0010-01
APC anti-mouse CD31 AntibodyBioLegend102510
APC anti-mouse CD45 AntibodyBioLegend103112
BD FACSMelody Cell SorterBD Biosciences 
BD Luer-Lok tip control syringe, 10-mLBD Biosciences 309604
Biotin anti-mouse CD106 AntibodyBioLegend105703
C57BL/6J  mouse (Female and Male)The Jackson Laboratory000664
B6.129S4-Pdgfratm11(EGFP)Sor/J mouseThe Jackson Laboratory007669
Corning BioCoat Collagen I 6-well Clear Flat Bottom TC-treated Multiwell PlateCorning356400
Corning BioCoat Collagen I 12-well Clear Flat Bottom TC-treated Multiwell PlateCorning356500
Corning BioCoat Collagen I 24-well Clear Flat Bottom TC-treated Multiwell PlateCorning356408
DAPI Solution (1 mg/mL)ThermoFisher Scientific62248
Disposable Aspirating Pipets, Polystyrene, SterileVWR414004-265
Donkey anti-Goat IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 488ThermoFisher ScientificA-11055
Falcon 40 µm Cell Strainer, Blue, SterileCorning352340
Falcon 60 mm TC-treated Cell Culture Dish, SterileCorning353002
Falcon Centrifuge Tubes, Polypropylene, Sterile, Corning, 15-mLVWR352196
Falcon Centrifuge Tubes, Polypropylene, Sterile, Corning, 50-mLCorning352070
Falcon Round-Bottom Tubes, Polypropylene, CorningVWR60819-728
Falcon Round-Bottom Tubes, Polystyrene, with 35um Cell Strainer Cap CorningVWR21008-948
Fibroblast Growth Factor, Basic, Human, Recombinant (rhFGF, Basic)PromegaG5071
FlowJo 10.8.1
Gibco Collagenase, Type II, powderThermoFisher Scientific17101015
Gibco Dispase, powderThermoFisher Scientific17105041
Gibco DMEM, high glucose, HEPESThermoFisher Scientific12430054
Gibco Fetal Bovine Serum, certified, United StatesThermoFisher Scientific16000044
Gibco Ham's F-10 Nutrient MixThermoFisher Scientific11550043
Gibco Horse Serum, New Zealand originThermoFisher Scientific16050122
Gibco PBS, pH 7.4ThermoFisher Scientific10010023
Gibco PBS (10x), pH 7.4ThermoFisher Scientific70011044
Gibco Penicillin-Streptomycin-Glutamine (100x)ThermoFisher Scientific10378016
Goat anti-Mouse IgG1 cross-absorbed secondary antibody, Alexa Fluor 555ThermoFisher ScientificA-21127
Hardened Fine ScissorsFine Science Tools Inc14090-09
Invitrogen 7-AAD (7-Aminoactinomycin D)ThermoFisher ScientificA1310
Mouse PDGF R alpha AntibodyR&D SystemsAF1062
Normal Donkey SerumFisher ScientificNC9624464
Normal Goat SerumThermoFisher Scientific31872
Pacific Blue anti-mouse Ly-6A/E (Sca 1) AntibodyBioLegend108120
Paraformaldehyde, 16%Fisher ScientificNCC0528893
Pax7 mono-clonal mouse antibody (IgG1) (supernatant)Developmental Study Hybridoma BankN/A
PE/Cyanine7 StreptavidinBioLegend405206
Student Vannas Spring ScissorsFine Science Tools Inc91500-09
Student Dumont #5 ForcepsFine Science Tools Inc91150-20
Triton X-100Sigma-AldrichT8787

References

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,
  1. Baskin, K. K., Winders, B. R., Olson, E. N. Muscle as a "mediator" of systemic metabolism. Cell Metabolism. 21 (2), 237-248 (2015).
  2. Dumont, N. A., Bentzinger, C. F., Sincennes, M. C., Rudnicki, M. A. Satellite cells and skeletal muscle regeneration. Comprehensive Physiology. 5 (3), 1027-1059 (2015).
  3. Relaix, F., et al. Perspectives on skeletal muscle stem cells. Nature Communications. 12 (1), 692(2021).
  4. Pawlikowski, B., Pulliam, C. J., Betta, N. D., Kardon, G., Olwin, B. B. Pervasive satellite cell contribution to uninjured adult muscle fibers. Skeletal Muscle. 5, 42(2015).
  5. Joe, A. W. B., et al. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nature Cell Biology. 12 (2), 153-163 (2010).
  6. Wosczyna, M. N., et al. Mesenchymal stromal cells are required for regeneration and homeostatic maintenance of skeletal muscle. Cell Reports. 27 (7), 2029-2035 (2019).
  7. Theret, M., Rossi, F. M. V., Contreras, O. Evolving roles of muscle-resident fibro-adipogenic progenitors in health, regeneration, neuromuscular disorders, and aging. Frontiers in Physiology. 12, 673404(2021).
  8. Uezumi, A., Fukada, S. I., Yamamoto, N., Takeda, S., Tsuchida, K. Mesenchymal progenitors distinct from satellite cells contribute to ectopic fat cell formation in skeletal muscle. Nature Cell Biology. 12 (2), 143-152 (2010).
  9. Wosczyna, M. N., Biswas, A. A., Cogswell, C. A., Goldhamer, D. J. Multipotent progenitors resident in the skeletal muscle interstitium exhibit robust BMP-dependent osteogenic activity and mediate heterotopic ossification. Journal of Bone and Mineral Research. 27 (5), 1004-1017 (2012).
  10. Eisner, C., et al. Murine tissue-resident PDGFRα+ fibro-adipogenic progenitors spontaneously acquire osteogenic phenotype in an altered inflammatory environment. Journal of Bone and Mineral Research. 35 (8), 1525-1534 (2020).
  11. Biferali, B., Proietti, D., Mozzetta, C., Madaro, L. Fibro-adipogenic progenitors cross-talk in skeletal muscle: The social network. Frontiers in Physiology. 10, 1074(2019).
  12. Lemos, D. R., et al. Nilotinib reduces muscle fibrosis in chronic muscle injury by promoting TNF-mediated apoptosis of fibro/adipogenic progenitors. Nature Medicine. 21 (7), 786-794 (2015).
  13. Malecova, B., et al. Dynamics of cellular states of fibro-adipogenic progenitors during myogenesis and muscular dystrophy. Nature Communications. 9, 3670(2018).
  14. Liu, L., Cheung, T. H., Charville, G. W., Rando, T. A. Isolation of skeletal muscle stem cells by fluorescence-activated cell sorting. Nature Protocols. 10 (10), 1612-1624 (2015).
  15. Maesner, C. C., Almada, A. E., Wagers, A. J. Established cell surface markers efficiently isolate highly overlapping populations of skeletal muscle satellite cells by fluorescence-activated cell sorting. Skeletal Muscle. 8, 6-35 (2016).
  16. Low, M., Eisner, C., Rossi, F. M. V. Fibro/adipogenic progenitors (FAPs): Isolation by FACS and culture. Methods and Protocols. 1556, 179-189 (2017).
  17. Judson, R. N., Low, M., Eisner, C., Rossi, F. M. V. Isolation, culture, and differentiation of fibro/adipogenic progenitors (FAPs) from skeletal muscle. Methods in Molecular Biology. 1668, 93-103 (2017).
  18. Hamilton, T. G., Klinghoffer, R. A., Corrin, P. D., Soriano, P. Evolutionary divergence of platelet-derived growth factor alpha receptor signaling mechanisms. Molecular and Cellular Biology. 23 (11), 4013-4025 (2003).
  19. Contreras, O., et al. Cross-talk between TGF-β and PDGFRα signaling pathways regulates the fate of stromal fibro-adipogenic progenitors. Journal of Cell Science. 132 (19), (2019).
  20. Holmes, C., Stanford, W. L. Concise review: Stem cell antigen-1: Expression, function, and enigma. Stem Cells. 25 (6), 1339-1347 (2007).
  21. Rodgers, J. T., Schroeder, M. D., Chanthia, M., Rando, T. A. HGFA Is an injury-regulated systemic factor that induces the transition of stem cells into GAlert. Cell Reports. 19 (3), 479-486 (2017).
  22. García-Prat, L., et al. FoxO maintains a genuine muscle stem-cell quiescent state until geriatric age. Nature Cell Biology. 22 (11), 1307-1318 (2020).
  23. Yamamoto, T., et al. Deterioration and variability of highly purified collagenase blends used in clinical islet isolation. Transplantation. 84 (8), 997-1002 (2007).
  24. Walls, P. L. L., et al. Quantifying the potential for bursting bubbles to damage suspended cells. Scientific Reports. 7 (1), 1-9 (2017).
  25. Kafadar, K. A., et al. Sca-1 expression is required for efficient remodeling of the extracellular matrix during skeletal muscle regeneration. Developmental Biology. 326 (1), 47-59 (2009).

Access restricted. Please log in or start a trial to view this content.

Reprints and Permissions

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Request permission to reuse the text or figures of this JoVE article

Request Permission

Tags

FACS IsolationFibro adipogenic ProgenitorsMuscle Stem CellsSkeletal MuscleFluorescence Activated SortingBiological RolePhysiological ConditionsPathological ConditionsMuscle DissectionIntermuscular FatTibialis AnteriorExtensor Digitorum LongusGastrocnemius MusclesQuadricepsHamstringsTissue Dissection