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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol outlines a method to isolate Fibro-adipogenic progenitors (FAPs) and myogenic progenitors (MPs) from rat skeletal muscle. Utilization of the rat in muscle injury models provides increased tissue availability from atrophic muscle for the analysis and a larger repertoire of validated methods to assess muscle strength and gait in free-moving animals.

Abstract

Fibro-adipogenic Progenitors (FAPs) are resident interstitial cells in skeletal muscle that, together with myogenic progenitors (MPs), play a key role in muscle homeostasis, injury, and repair. Current protocols for FAPs identification and isolation use flow cytometry/fluorescence-activated cell sorting (FACS) and studies evaluating their function in vivo to date have been undertaken exclusively in mice. The larger inherent size of the rat allows for a more comprehensive analysis of FAPs in skeletal muscle injury models, especially in severely atrophic muscle or when investigators require substantial tissue mass to conduct multiple downstream assays. The rat additionally provides a larger selection of muscle functional assays that do not require animal sedation or sacrifice, thus minimizing morbidity and animal use by enabling serial assessments. The flow cytometry/FACS protocols optimized for mice are species specific, notably restricted by the characteristics of commercially available antibodies. They have not been optimized for separating FAPs from rat or highly fibrotic muscle. A flow cytometry/FACS protocol for the identification and isolation of FAPs and MPs from both healthy and denervated rat skeletal muscle was developed, relying on the differential expression of surface markers CD31, CD45, Sca-1, and VCAM-1. As rat-specific, flow cytometry-validated primary antibodies are severely limited, in-house conjugation of the antibody targeting Sca-1 was performed. Using this protocol, successful Sca-1 conjugation was confirmed, and flow cytometric identification of FAPs and MPs was validated by cell culture and immunostaining of FACS-isolated FAPs and MPs. Finally, we report a novel FAPs time-course in a prolonged (14 week) rat denervation model. This method provides the investigators the ability to study FAPs in a novel animal model.

Introduction

Fibro-adipogenic progenitor cells (FAPs) are a population of resident multipotent progenitor cells in skeletal muscle that play a critical role in muscle homeostasis, repair, and regeneration, and conversely, also mediate pathologic responses to muscle injury. As the name suggests, FAPs were originally identified as a progenitor population with the potential to differentiate into fibroblasts and adipocytes1 and were purported to be the key mediators of fibro-fatty infiltration of skeletal muscle in chronic injury and disease. Further study revealed that FAPs are additionally capable of osteogenesis and chondrogenesis2,3,4. Thus, they are more broadly notated in the literature as mesenchymal or stromal progenitors3,5,6,7,8. In acute skeletal muscle injury, FAPs indirectly aid in regenerative myogenesis by transiently proliferating to provide a favorable environment for activated muscle satellite cells and their downstream myogenic progenitor (MPs) counterparts1,9,10. In parallel with successful regeneration, FAPs undergo apoptosis, returning their numbers to baseline levels1,9,10,11. In contrast, in chronic muscle injury, FAPs override pro-apoptotic signals, which results in their persistence9,10,11 and abnormal muscle repair.

In vivo studies evaluating the cellular and molecular mechanisms by which FAPs mediate muscle responses have utilized murine animal models to date1,7,9,10,11,12,13,14. While genetically engineered mice are powerful tools for use in these analyses, the small size of the animal limits tissue availability for study in long-term localized injury models where muscle atrophy can be profound, such as traumatic denervation. Furthermore, measurement of muscle strength and physical function requires ex vivo or in situ measurements that necessitate termination of the mouse, or in vivo methods that require surgery and/or a general anaesthetic to permit evaluation of muscle contractile performance15,16,17,18,19,20. In rats, well validated and globally utilized muscle functional analyses, in addition to analyses for more complex motor behaviors such as gait analysis (e.g., Sciatic Function Index, CatWalk analysis) exist and are performed in awake and spontaneously moving animals21,22,23,24. This additionally optimizes the principles of minimal morbidity in animal experimentation, and numbers of research animals used. The rat thereby provides the FAPs investigator the added flexibility of greater injured muscle volume for protein and cellular analyses and the ability to undertake serial assessments of muscle complex static and dynamic functional activity and behaviors, in the alert animal.

FAPs have primarily been identified and isolated from whole muscle samples using flow cytometry and Fluorescence-activated cell sorting (FACS) respectively. These are laser-based assays that are able to identify multiple specific cell populations based on characteristic features such as size, granularity, and a specific combination of cell surface or intracellular markers25. This is highly advantageous in the study of an organ system such as skeletal muscle, as homeostasis and regeneration are complex, multifactorial processes coordinated by a plethora of cell types. A seminal study identified FAPs, as well as MPs, using flow cytometric methods in mouse skeletal muscle1. They demonstrated that FAPs are mesenchymal in nature, as they lacked surface antigens specific to cells from endothelial (CD31), hematopoietic (CD45), or myogenic (Integrin-α7 [ITGA7]) origins, but expressed the mesenchymal stem cell marker Sca-1 (Stem cell antigen 1)1 and differentiated into fibrogenic and adipogenic cells in culture. Other studies demonstrated successful isolation of mesenchymal progenitors in muscle based on the expression of an alternative stem cell marker, platelet-derived growth factor receptor alpha (PDGFRα)2,7,8 and further analysis revealed these likely to be the same cell population as FAPs3. FAPs are now commonly identified in flow cytometry using either Sca-1 or PDGFRα as a positive selection marker1,9,10,11,12,13,14,26,27,28,29,30,31. The use of PDGFRα is preferential for human tissue however, as a direct human homologue of murine Sca-1 has yet to be identified32. In addition, other cell surface proteins have been reported as markers of MPs (e.g., VCAM-1), providing a potential alternative to ITGA7 as an indicator of cells of myogenic lineage during FAPs isolation33.

While flow cytometry/FACS is a powerful methodology for studying the role and pathogenic potential of FAPs in skeletal muscle1,9,10,11,13,29, it is limited technically by the specificity and optimization of its required reagents. Since flow cytometric identification and isolation of FAPs has been developed and conducted in mouse animal models1,9,10,11,29, this poses challenges for researchers who wish to study FAPs in other model organisms. Many factors - such as optimal tissue size to be processed, as well as reagent and/or antibody specificity and availability - differ depending on the species used.

In addition to the technical barriers to studying FAPs in a novel animal model, they have largely been studied in an acute, toxic setting - usually via intramuscular chemical injection or cardiotoxin. Evaluation of the long-term dynamics of FAPs is limited primarily to assessment in Duchenne's muscular dystrophy, using the mdx mouse model9,10,11, and models of combination muscle injury such as massive rotator cuff tear where concurrent tendon transection and denervation is performed on shoulder musculature26,27,28. The response of FAPs to the sole insult of chronic traumatic denervation, a common occurrence in work-place accidents in heavy industry, agriculture, and in birth traumas (brachial plexus injury)34,35,36,37 with significant morbidity, has not been as well characterized, often limited to a short-term time frame11,38.

We describe a method for identifying and isolating FAPs and MPs from healthy as well as severely atrophic and fibrotic skeletal muscle in the rat. First, identification of CD31-/CD45-/Sca-1+/VCAM-1- FAPs and CD31-/CD45-/Sca-1-/VCAM-1+ MPs using a tissue digestion and flow cytometry staining protocol is demonstrated and subsequent validation of our findings is performed through culture and immunocytochemical staining of FACS-isolated cells. Using this method, we also report a novel FAPs time-course in a long-term isolated denervation injury model in the rat.

Protocol

Investigators conducting this protocol must receive permission from their local animal ethics board/care committee. All animal work was approved by the St. Michael's Hospital Unity Health Toronto Animal Care Committee (ACC #918) and was conducted in accordance with the guidelines set forth by the Canadian Council on Animal Care (CCAC). A schematic of the flow cytometry protocol is shown in Figure 1. If the downstream application is FACS and subsequent cell culture, all steps should be completed with proper aseptic technique.

1. Muscle harvesting

  1. Anesthetize rats using an appropriate anesthetic and sacrifice according to local vivarium and animal ethics board guidelines. This protocol harvests the gastrocnemius muscle from adult female Lewis rats (200-250 g), as an example. Rats were anesthetized using 2-3% Isoflurane and were sacrificed by intracardiac injection of T61.
  2. Once the animal has been sacrificed, shave the whole hindlimb to facilitate the location of the muscle and minimize fur contamination of the harvested tissue.
  3. Using a sterile scalpel, make two incisions in the skin: the first around the circumference of the ankle joint and the second up the midline of the medial aspect of the hindlimb from the ankle to the hip.
  4. Peel back the skin and superficial muscle layers to reveal the underlying gastrocnemius, which originates at the medial and lateral condyles of the femur and inserts at the Achilles Tendon.
  5. Use blunt dissection to separate the gastrocnemius from the surrounding tissue, handling the muscle only by the tendon to avoid crush injury.
  6. Separate the gastrocnemius from its insertion by transecting the Achilles Tendon as distally as possible with sharp scissors. Once cut, grasp the Achilles tendon with forceps and gently peel the gastrocnemius off the underlaying bone. Still holding the muscle with forceps in one hand, locate the gastrocnemius' two origins and cut at the medial and lateral femoral condyles.
  7. Blot the excised gastrocnemius gently against a sterile piece of gauze to remove as much blood as possible. Trim the muscle on a sterile surface and remove any excess connective tissue as well as the Achilles Tendon.
  8. Place muscle in a weigh-boat and weigh using a precision scale. This protocol is optimized to digest muscle with a wet weight ranging from 200-600 mg. Operators may subdivide excess harvested tissue for other downstream assays, if desired.
  9. Gently further divide harvested muscle to be used for flow cytometry into 3-4 smaller pieces (approximately 1-2 cm3) and submerge in ice-cold 1x PBS. Keep cold on ice until all samples have been harvested.

2. Muscle digestion

  1. Remove muscle from PBS and place in a sterile 10 cm cell culture dish. Gently tear and mince tissue with forceps until pieces are approximately 3-4 mm3, removing as much connective tissue as possible. Once thoroughly minced, transfer to a sterile 50 mL conical tube containing 6 mL DMEM + 1% penicillin/streptomycin (P/S).
  2. Add 10 µL of 300 mM CaCl2 solution to 365 µL of Collagenase II solution (stock concentration 4800 U/mL) to activate the collagenase enzyme. Add the activated collagenase II solution to the 50 mL conical tube containing the tissue slurry. The final Collagenase II concentration is 250 U/mL.
  3. Incubate tubes in a shaker for 1 h at 37 °C, 240 x g, making sure to manually swirl every 15 min to dislodge any tissue that has adhered to the side of the tube.
  4. After 1 h, remove tubes from shaker and add the following per sample: 100 µL of Collagenase II (4,800 U/mL) and 50 µL of Dispase (4.8 U/mL).
  5. Pipette samples using a serological pipette 15-20 times until the solution is homogenous. If processing multiple samples, use a separate sterile pipette for each sample to avoid sample cross-contamination.
  6. Incubate again in a shaker for 30 min at 37 °C and 240 x g. After 15 min, shake samples by hand to dislodge adherent tissue off the side of the tube.

3. Generation of single cell suspension

  1. Slowly shear samples through a 10 mL syringe with a 20 G needle for 10 cycles.
    NOTE: One cycle involves taking up muscle solution into syringe, and injecting it back into tube. Ensure to minimize any bubbles by completing shearing slowly, as excessive frothing can cause additional cell death39.
  2. Place a 40 µm cell strainer on a sterile 50 mL conical tube and wet it by pipetting 5 mL DMEM + 10% FBS & 1% P/S.
  3. Pipette 1 mL of the sample at a time through the cell strainer.
  4. Wash the cell strainer by pipetting DMEM with 10% FBS and 1% P/S through the strainer to bring the total volume in the tube to 25 mL.
  5. Split 25 mL of the sample equally into two 15 mL conical tubes and centrifuge at 15 °C, 400 x g for 15 min.
    NOTE: Splitting the muscle solution into two 15 mL conical tubes ensures better cell recovery after centrifugation compared to a single tube.
  6. Aspirate the supernatant and re-suspend the pellet in 1 mL 1x RBC Lysis buffer (see Supplementary File) at room temperature for 7 min to eliminate erythrocytes.
  7. Bring up the volume to 10 mL with 9 mL of wash buffer (see Supplementary File) and spin tubes at 400 x g, 15 °C for 15 min.
  8. Aspirate the supernatant and recombine pellets by re-suspending in 1 mL wash buffer.
  9. Transfer an appropriate volume of cells to a separate 1.5 mL microcentrifuge tube and mix with trypan blue dye. Count live cells on a light microscope using a hemocytometer.

4. Antibody staining for flow cytometry

NOTE: The Sca-1 antibody must be conjugated to APC prior to flow cytometry/FACS experiments, as per the manufacturer's instructions. Performance must be validated for each batch of conjugates (Figure 2). Final conjugations can be stored in 20 µL aliquots at -20 °C and are stable for three weeks. Refer to the Supplementary File for full conjugation protocol.

  1. For flow cytometry, transfer 1-2 x 106 cells per experimental sample to a sterile 1.5 mL microcentrifuge tube. Bring volume up to 1 mL with wash buffer and place on ice.
  2. For each experiment, set up the following required controls: i) unstained and ii) viability controls to accurately select for the live cell population; iii) fluorescence minus one (FMO) controls on single cell suspensions to set accurate gates for CD31-/CD45- fractions, FAPs, and MPs; and iv) single-stained compensation beads to correct for fluorescence spillover between channels.
    1. For all cell controls, aliquot 5 x 105 - 1 x 106 cells in 1 mL of wash buffer in a 1.5 mL microcentrifuge tube and place on ice.
    2. For bead controls, add 1 drop of positive compensation beads (~1.5 x 105 beads per drop) to each labeled 1.5 mL microcentrifuge tube. The full complement of controls is listed in Table 1.
      NOTE: If the experiment is being performed for the first time, run single-stained controls for each conjugated antibody on single cell suspensions (in addition to unstained, viability, single-stained compensation bead and FMO controls) to assess the positive stained population in cells and validate staining observed on compensation beads. Validate every freshly-conjugated Sca-1::APC preparation by performing single-staining on compensation beads and single cell suspensions. Refer to Table 1 for a full list of staining controls.
  3. To prepare the viability control, transfer half of the volume of cells from the "viability" tube to a new 1.5 mL microcentrifuge tube. Label this tube "Dead".
  4. Incubate "Dead" tube at 65 °C for 2-3 min to kill the cells, then place on ice. After 2-3 min, re-combine dead cells with live cells remaining in the viability control tube. This population of cells will be used to set compensation values (if needed) and properly set gates for the viability dye.
  5. Centrifuge the single cell suspensions (experimental samples and controls) at 500 x g, 4 °C for 5 min.
  6. Aspirate the supernatant and re-suspend cell pellets in 100 µL wash buffer.
  7. Add antibodies, depending on the experimental sample or control. Refer to the staining matrix (Table 2) for information on antibody combinations and amounts.
  8. Gently flick each sample to ensure complete mixing and incubate on ice in the dark for 15 min. For compensation beads, incubate at room temperature in the dark for 15 min.
  9. For single cell suspension experimental and control samples, bring up the volume to 1 mL by adding 900 µL wash buffer. For compensation bead controls, bring up the volume to 1 mL with 900 µL of 1x PBS.
  10. Centrifuge single cell suspension samples at 500 x g, 4 °C for 5 min. Centrifuge compensation bead controls at 300 x g, 4 °C for 5 min.
  11. For all single cell suspension samples, aspirate and discard supernatant and re-suspend cell pellet in 300 µL wash buffer. For compensation bead controls, aspirate and discard the supernatant, re-suspend the pellet in 300 µL of 1x PBS, then add 1 drop (~1.5 x 105) of negative compensation beads.
  12. Keep all single cell suspension samples on ice under aluminum foil and proceed to flow cytometric acquisition. Compensation bead controls should also be protected from light but can be kept at room temperature.
    NOTE: If experimental endpoint is FAPs identification by flow cytometry, please follow steps 5.1.1-5.1.11. If endpoint is cell isolation via FACS for culture and staining, please follow steps 5.2.1-5.2.9 and sections 6-7.

5. Flow cytometry and fluorescence-activated cell sorting (FACS)

  1. Flow cytometry
    NOTE: This protocol employs a benchtop flow cytometer equipped with 405 nm, 488 nm, and 640 nm lasers that are capable of simultaneously distinguishing 10 different colors. Bandpass filters and their associated fluorochromes used in this protocol are as follows: 450/50 (SYTOX Blue), 530/30 (FITC), 575/25 (PE), and 670/30 (APC). Voltages for each detector are as follows: FSC 700; SSC 475; FITC 360; PE 460; PE-Cy7 600; SYTOX Blue 360; APC 570. Ensure you are trained on the proper operation of the flow cytometer or cell sorter prior to use.
    1. Ensure the cytometer has been turned on for 10-20 min before use and has been primed by cleaning sequentially with clean, rinse, and sheath fluid solutions for 30-45 s each. Finish with a rinse with dH2O. Ensure that an adequate volume of sheath fluid has been added to the storage container to maintain proper sample flow throughout acquisition.
    2. Set up the gating strategy to identify FAPs and MPs as delineated in Figure 3.
      NOTE: FAPs and MPs are identified by the following hierarchical gating strategy: i) SSC-A vs FSC-A (side cell scatter area versus forward cell scatter area to separate cells vs debris), ii) FSC-W vs FSC-H (forward cell scatter width versus forward cell scatter height to discriminate singlets from doublets in the FSC parameter), iii) SSC-W vs SSC-H (side cell scatter width versus side cell scatter height to discriminate singlets from doublets in the SSC parameter), iv) SSC-A vs SYTOX Blue (to distinguish live versus dead singlets), v) SSC-A vs CD31/45::FITC (to exclude CD31+ and CD45+ cells from further analysis), and vi) Sca-1::APC vs VCAM-1::PE from the CD31-/CD45- (Lineage; Lin-) population (identification of FAPs and MPs). FAPs are identified as CD31-/CD45-/Sca-1+/VCAM-1- events and MPs are identified as CD31-/CD45-/Sca-1-/VCAM-1+ events.
    3. First run each single-stained compensation bead control through the cytometer on low speed to generate compensation values used to correct for any fluorescence spillover between channels. Assess compensation by comparing fluorescent signal of each control in its own detector (e.g., SSC-A vs APC for Sca-1::APC single-stained beads) as well as all other detectors. There should be two distinct populations (one with negative and one with positive signal) in the appropriate detector and only a negative population in all other detectors. Set the stopping gate to 10,000 compensation bead events and record the data.
      NOTE: In between acquisition of each sample, make sure to run dH2O through the cytometer for 10-20 sec to avoid sample-to-sample contamination.
    4. Next process the unstained and viability control samples to properly gate on live single cells. Set the stopping gate to 10,000 singlet events and record data.
      NOTE: Approximately 5 min before acquisition of each single cell suspension sample with the exception of the unstained sample, add 1 µL of SYTOX Blue viability dye (300 µM working concentration diluted from 1 mM stock solution) to each sample and flick gently to mix (final concentration 1 µM).
    5. Then acquire the remaining single cell suspension control samples. Assess each FMO control with its appropriate plot in the gating strategy (Figure 3). For example, assess the FITC signal of the CD31+CD45 FMO to ensure an accurate CD31-/CD45- gate. An optimal example is shown in Figure 3G. If the protocol is being performed for the first time, single-stained controls on cells should be run before the acquisition of FMO controls.
    6. Assess the fluorescent signal of each single-stained cell sample in its appropriate detector as well as in all other detectors to validate proper compensation. Set the stopping gate to 10,000 live singlet events and record on the software.
    7. Once all controls (single cell suspensions and beads) have been processed, prepare all experimental samples by first measuring and recording the volume of each sample. These measurements will be used to accurately quantify FAPs and MPs, as described in step 5.1.11. Then, add 50 µL of precision counting beads and gently mix by pipetting up and down 2-3 times.
    8. Briefly run the first experimental sample to validate identification of the counting bead population. This population appears as a small distinct cluster separate from the general cell population on the FSC-A vs SSC-A plot (Figure 3A, red box). Create a gate around the counting bead population. Then acquire data for each experimental sample by processing through the cytometer on low speed. Set the stopping gate to 10,000 counting bead events and record.
      NOTE: Investigators may alternatively identify counting beads by setting up an additional plot assessing SSC-A versus any of the detectors, as the counting beads are fluorescent in all detectors.
    9. After all samples have been processed, clean the cytometer using the appropriate protocols. Export all data for analysis.
    10. Open all data files on an appropriate flow cytometry analysis software. Set the gating strategy as used for data acquisition as described in step 5.1.2. Examine controls in the same order as in data acquisition (e.g., unstained, viability, single stain, then FMO controls) to re-validate the gating strategy. Once accurate gates have been set using FMO controls, apply the gates to all experimental samples. Export raw data as a spreadsheet for quantification.
    11. Calculate the number of FAPs and MPs in each experimental sample using the counting beads:
      figure-protocol-16149
      where, Acquired Cell Count is the number of recorded events of pertinent cell population (e.g. FAPs or MPs) on the acquisition software; Acquired Bead Count is the number of recorded events of counting beads on the acquisition software; Counting Beads Volume is the volume of counting bead solution added in step 5.1.7; Sample Volume is the volume of each stained experimental sample prior to addition of counting beads.; Bead Concentration is the number of beads per µL solution; this value is found on the product datasheet.
  2. FACS - sorting for cell culture
    NOTE: This protocol performs FACS on a cell sorter equipped with 4 lasers (UV, Violet, Blue, Red) that is capable of simultaneously distinguishing 11-14 colors. Follow the experimental sample staining (section 4) and flow cytometry protocol, with the exceptions of steps 1 to 3 delineated below, to optimize the FACS workflow:
    1. Increase the concentration of cells in the experimental samples to be sorted to 7 x 106 cells/mL to generate robust yields of FAPs and MPs.
    2. To account for this significant increase in the cell concentration, double all antibody concentrations in the experimental samples to be sorted.
    3. Process the final stained cell samples through a 40 µM cell strainer cap affixed to a 5 mL polystyrene tube immediately prior to sorting to reduce cell clumping and increase sort yields.
    4. Collect single, live rat FAPs and MPs directly from the cell sorter into a 5 mL polypropylene collection tube containing 1 mL of sterile, 100% Fetal Bovine Serum (FBS). Keep cells on ice until sorting is complete.
      NOTE: If conducting FACS at an off-site location, transfer all sorted cells on ice and in a secured, covered container.
    5. Working in a sterile biosafety cabinet (BSC), bring volume of sorted cells up to 7 mL with appropriate growth media (e.g., FAP growth media (FAP GM) for sorted FAPs, and MP growth media (MP GM) for sorted MPs; see Supplementary File for recipes) and centrifuge at 500 x g, 4 °C for 7 min to remove as much residual wash buffer as possible.
    6. Resuspend pellets in 1 mL of appropriate growth media and plate into a 12-well plate containing a sterile, collagen-coated 12 mm glass coverslip/well for subsequent immunostaining (see section 6).
      NOTE: If immunocytochemistry staining for collagen, plate sorted cells into a 12 well plate containing a sterile, laminin-coated 12 mm glass coverslip/well, instead of collagen-coated. If immunocytochemistry experiments of immediately isolated progenitors are required, seed FAPs and MPs at a density of 15,000 cells per cm2 and proceed directly to step 6.1. For long-term cultures to induce progenitor differentiation, seed FAPs at a density of 5,000 cells per cm2, and MPs at a density of 7,500 cells per cm2.
    7. Incubate cells at 37 °C and 5% CO2 in a cell culture incubator. After 72 h in culture, change half of the media. Change media fully every 2-4 d after.
    8. To induce myocyte development, switch MPs cultures to MP differentiation (MD) medium on Day 9 of culture. To induce adipocytes, switch FAPs cultures to FAP adipogenic differentiation (AD) medium on Day 10 of culture.
    9. To induce fibrogenesis, FAPs may be switched to fibrogenic differentiation (FD) media at variable times during culture, or alternatively, may be seeded directly into FD media following isolation (step 5.2.6) (See Supplementary File for all media recipes).

6. Immunocytochemistry of cultured FAPs and MPs

  1. To validate cell sorting and demonstrate purity of FAPs and MPs cultures, immunostain with cell-type specific markers including PDGFRα (FAPs marker), Pax-7 (muscle stem [satellite] cell marker), Fibroblast-specific protein (FSP-1, fibroblast marker), Perilipin-1 (Plin-1, adipocyte marker), Collagen type 1 (Col1a1, indicator of fibrosis), Myosin Heavy Chain (MHC, mature myocyte marker).
    1. For immunostaining of freshly sorted cells, centrifuge the 12-well plate at 200 x g for 3 min at room temperature to facilitate adherence of cells to the coverslip/well. This step is not necessary for long-term cultures. Remove culture media.
    2. For immunostaining with FSP-1, fix cells with 1 mL 100% methanol (MeOH) for 2 min at 4 °C. If immunostaining for PDGFRα, Plin-1, Pax7 or Col1a1, fix cell cultures with 1 mL 4% PFA in 1x PBS for 15 min at room temperature. MHC immunostaining tolerates either fixative.
      NOTE: For methanol-fixed cells, skip step 6.2 and proceed to step 6.3.
  2. Aspirate 4% PFA and quickly wash cell cultures 3-4 times with 1x PBS. Add 1 mL of 100 mM Glycine in 1x PBS and incubate for 10 min at room temperature to inactivate residual PFA. Aspirate and wash 1-2 times with 1x PBS.
    NOTE: Cells can be left at this stage in 2 mL of 1x PBS, wrapped in cling film and stored at 4 °C for 7-10 days maximum.
  3. After washing, add 1 mL of 0.1% Triton-X in 1x PBS and incubate for 20 min to permeabilize cell membranes.
  4. Wash wells 2-3 times with 1-2 mL of 1x PBS then block cells with 1 mL of 1x PBS + 3% BSA per well for 1 h at room temperature.
  5. Pipette 80 µL of primary antibody diluted in 1x PBS + 3% BSA (PDGFRα 1:100, Pax7 neat, FSP-1 1:50, Plin-1 1:400, Col1a1 1:250, MHC 3 µg/mL) onto a piece of parafilm taped to a mobile container. Using sterile fine forceps, carefully lift the coverslip out of the well and invert onto the drop of antibody solution. Incubate coverslip with two wet pieces of paper towel and cover container in plastic film to avoid evaporation of the antibody solution. Incubate overnight at 4 °C.
    NOTE: Staining coverslips out of the well utilizes less antibody (~80 µL) than staining inside the well (500 µL minimum).
  6. On Day 2, leave coverslips at room temperature for 30 min to warm. Using forceps carefully right and transfer coverslips back to their respective wells (cells facing up) and wash 2-3 times with 1-2 mL of 1x PBS for 2 min each to remove as much primary antibody as possible.
  7. Using the same staining technique as with primary antibody staining, stain cells with goat anti-rabbit Alexa Fluor 488 secondary antibody (1:400) to detect FSP-1, Plin-1, Col1a1, or PDGFRα and goat anti-mouse Alexa Fluor 555 secondary antibody (1:300) to detect MHC or Pax-7. Incubate cells for 1 h at room temperature and keep cells protected from light.
  8. Return cells to the well and incubate cells with Hoechst (1:10,000) for 2-4 min at room temperature. Wash cells another 2-3 times with 1x PBS for 2 min each to remove excess Hoechst.
  9. Mount coverslips onto glass slides using an anti-fade fluorescent mounting medium and leave slides to dry overnight in the dark at room temperature. Store mounted coverslips at 4 °C in the dark.

7. Oil Red O (ORO) staining of cultured FAPs and MPs

  1. Perform ORO staining on non-permeabilized cells, as permeabilization of the cell membrane can result in non-specific/undesired staining of non-adipogenic cell types. Prior to commencing staining, prepare an ORO working stock (See Supplementary File for the recipe) and incubate at room temperature for 20 min.
  2. After 20 min, filter the solution using a 0.2 µm filter in order to remove any undissolved aggregates.
  3. Aspirate media from well and add 1 mL of 10% Neutral Buffered Formalin (10% NBF). Incubate for 5 min at room temperature.
    NOTE: Cell confluency can result in lifting from the well/coverslip. Take care when aspirating/adding solutions.
  4. Aspirate and add 1 mL of fresh 10% NBF and incubate for at least 1 h at room temperature.
    NOTE: The protocol can be stopped at this point, as cells can be left in 10% NBF overnight.
  5. Quickly wash the wells once with 1 mL of 60% isopropanol, then aspirate and allow the wells to dry completely (approximately 2 min).
  6. Add 400 µL Oil Red O working stock per well and incubate for 10 min at room temperature, making sure to avoid pipetting any ORO on the walls of the plate.
  7. Remove all of the Oil Red O and quickly wash the well 4 times with dH2O.
    NOTE: If stained wells contain coverslips, mount using the same technique as described in step 6.9.
  8. Image either mounted coverslips or the stained well using a brightfield microscope.

8. Tissue staining of contralateral and denervated rat gastrocnemius sections

  1. Picrosirius Red (PSR)
    1. Perform PSR staining on 5 µm-thick, formalin-fixed paraffin embedded (FFPE) rat gastrocnemius histologic sections as previously described40.
  2. Oil Red O (ORO)
    1. Fix 5 µm-thick isopentane-frozen rat gastrocnemius histologic sections in 4% PFA for 10 min, incubate in 60% isopropyl alcohol for 1 min.
    2. Stain with ORO working stock for 12 min. Incubate in 60% isopropyl alcohol for 1 min, wash for 10 min in dH2O. Mount on coverslips using a water-soluble mounting media.
  3. Sca-1 and laminin tissue fluorescent immunohistochemistry (IHC)
    1. Perform fluorescent IHC on 5 µm-thick isopentane-frozen rat gastrocnemius histologic sections.
    2. Hydrate samples in 1x PBS for 5 min, fix in 4% PFA for 10 min then incubate samples in tissue IF blocking solution (see Supplementary File) for 90 min.
    3. Incubate with anti-Sca-1 primary antibody (1:500) diluted in 1x PBS + 0.05% Tween at 4 °C overnight.
    4. On Day 2, wash three times in 1x PBS + 0.05% Tween for 5 min each, then incubate in goat anti-rabbit Alexa Fluor 555 (1:500) for 1 h.
    5. Wash again (as before), incubate with blocking solution for 1 h, then add anti-laminin primary antibody (1:500) diluted in 1x PBS + 0.05% Tween for 1 h.
    6. Wash again (as before), then incubate in goat anti-rabbit Alexa Fluor 488 (1:500) for 1 h (for laminin).
    7. Wash again (as before) then incubate in DAPI (1:10,000) for 4 min. Wash and mount on coverslips using anti-fade mounting medium.

Results

Identifying FAPs and MPs via flow cytometry using a novel antibody panel including Sca-1 and VCAM-1
The gating strategy for identifying FAPs in rat muscle is based upon flow cytometry protocols in the mouse29, which gate on CD31 (endothelial) and CD45 (hematopoietic) positive cells (termed the lineage [Lin]) and examines the fluorescent profile of FAPs marker Sca-1 and MPs marker ITGA7 from the linage-negative (Lin-) populat...

Discussion

An optimized, validated FAPs isolation protocol for rat muscle is essential for researchers who wish to study injury models that are not feasible in the mouse for biologic or technical reasons. For example, mice are not an optimal animal model in which to study chronic local, or neurodegenerative injuries such as long-term denervation. Biologically, the short lifespan and rapid aging of mice make it difficult to accurately delineate the muscle sequalae due to denervation from the confounding factor of aging. From a techn...

Disclosures

The authors have no conflicts to disclose.

Acknowledgements

We would like to thank the Flow cytometry Core Facilities at the University of Ottawa and the Keenan Research Centre for Biomedical Sciences (KRC), St Michaels Hospital Unity Health Toronto for their expertise and guidance in optimization of the flow cytometry/FACS protocol presented in this manuscript. This work was funded by Medicine by Design New Ideas 2018 Fund (MbDNI-2018-01) to JB.

Materials

NameCompanyCatalog NumberComments
5 mL Polypropylene Round-Bottom TubeFalcon352063
5 mL Polystyrene Round-Bottom Tube with Cell-Strainer CapFalcon352235
10 cm cell culture dishesSarstedt83.3902
12-well cell culture plateThermoFisher130185
12 mm glass coverslips, No.2VWR89015-724
10 mL SyringeBeckton Dickenson302995
15 mL centrifuge tubesFroggaBio91014
20 gauge needleBeckton Dickenson305176
25mL Serological pipetteSarstedt86.1685.001
40µm cell strainerFisher Scientific22363547
50mL centrifuge tubesFroggaBioTB50
AbC Total Antibody Compensation Beads ThermoFisherA10497
Ammonium Chloride, Reagent GradeBioshopAMC303.500
APC Conjugation Kit, 50-100µgBiotium92307
Aquatex Aqueous Mounting MediumMerck108562
Biolaminin 411 LNBiolaminaLN411
Bovine Serum Albumin (BSA)BioshopALB001
Calcium ChlorideBioshopCCL444.500
Collagenase Type IIGibco17101015
CountBright Plus Absolute Counting BeadsThermoFisherC36995
DexamethasoneMillipore SigmaD4902
DispaseGibco17105041
Dulbecco’s Modified Eagle Medium (DMEM) (1X)Gibco11995-065(+)4.5 g/L D-Glucose
(+)L-Glutamine
(+)110 mg/L Sodium Pyruvate
EDTAFisherScientificS311
FACSClean SolutionBeckton Dickenson340345
FACSDiva SoftwareBeckton Dickenson--
FACSRinse SolutionBeckton Dickenson340346
Fetal Bovine SerumSigmaF1051
Flow Cytometry Sheath FluidBeckton Dickenson342003
FlowJo SoftwareBeckton Dickenson--
Fluorescent Mounting MediumDakoS302380-2
Goat anti-mouse Alexa Fluor 555 secondary antibodyInvitrogenA21424
Goat anti-rabbit Alexa Fluor 488 secondary antibodyInvitrogenA11008
Goat anti-rabbit Alexa Fluor 555 secondary antibodyInvitrogenA21429
Goat SerumGibco16210-064
Ham's F10 MediaThermoFisher 11550043(+) Phenol Red
(+) L-Glutamine
(-) HEPES
Hank’s Balanced Salt Solution (HBSS) (1X)Multicell311-513-CL
Heat Inactivated Horse SerumGibco26050-088
HemocytometerReichertN/A
HEPES, minimum 99.5% titrationSigmaH3375
Horse SerumThermoFisher16050130
Human Transforming Growth Factor β1 (hTGF-β1)Cell Signaling8915LF
Humulin RLillyHI0210
IBMXMillipore SigmaI5879Also known as 3-Isobutyl-1-methylxanthine
IsopropanolSigmaI9516Also known as 2-propanol
Lewis Rat, FemaleCharles River Kingston004 (Strain Code)200-250 grams used
LSRFortessa X-20 Benchtop CytometerBeckton Dickenson--
MicrocentrifugeEppendorfEP-5417R
MoFlo XDP Cell SorterBeckman Coulter--
Mouse Anti-CD31::FITC AntibodyAbcamab33858Clone TLD-3A12
Mouse Anti-CD45::FITC AntibodyBiolegend202205Clone OX-1
Mouse Anti-CD106::PE AntibodyBiolegend200403Also known as VCAM-1
Mouse Anti-MHC AntibodyDevelopmental Studies Hybridoma Bank (DSHB)N/AAlso known as MF20
Mouse Anti-Pax7 AntibodyDevelopmental Studies Hybridoma Bank (DSHB)N/A
Neutral Buffered Formalin, 10 %SigmaHT501128
Oil Red OMillipore SigmaO0625
PE-Cy7 Conjugation KitAbcamab102903
Penicillin-StreptomycinSigma P4333
Phosphate Buffered Saline, pH 7.4 (1X)Gibco10010-023(-)Calcium Chloride
(-)Magnesium Chloride
Potassium Bicarbonate, Reagent GradeBioshopPBC401.250
Rabbit Anti-Fibroblast Specific Protein 1 (FSP-1) AntibodyInvitrogenMA5-32347FSP-1 also known as S100A4
Rabbit Anti-Integrin-a7 AntibodyAbcamab203254
Rabbit Anti-Laminin AntibodySigmaL9393
Rabbit Anti-Perilipin-1 AntibodyAbcamab3526
Rabbit Anti-Sca-1 AntibodyMillipore SigmaAB4336
Rabbit Recombinant Anti-Collagen Type I AntibodyAbcamab260043Also known as Col1a1
Rabbit Recombinant Anti-PDGFR Alpha AntibodyAbcamab203491
Recombinant Human FGF-basicGibcoPHG0266
Sodium AzideSigmaS2002
Triton-X-100Fisher ScientificBP151
TroglitazoneMillipore SigmaT2573
Tween-20BioshopTWN510

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