As FAPs are mediators of muscle regeneration and pathological fibrosis, our protocol allows for characterization of FAP dynamics post muscle injury and isolation of FAPs for in vitro or ex vivo studies. To date, studies have been performed exclusively on FAPs isolated from mice. Our protocol enables effective isolation of FAPs from the larger rat, providing much greater tissue availability for downstream assays.
Prolonged tissue processing time can negatively impact FAPs viability. If multiple samples are being processed simultaneously, it is recommended that two operators perform the protocol in tandem to maximize isolated cell viability. Begin by placing the harvested muscle tissue in a sterile 10-centimeter cell culture dish.
Gently tear and mince the tissue with forceps and remove connective tissue to obtain approximately three to four-millimeter cubed pieces. Transfer the minster tissue to a sterile 50-milliliter conical tube containing six milliliters of DMEM and 1%penicillin-streptomycin. Next, activate 365 microliters of collagenase II solution by adding 10 microliters of 300-millimolar calcium chloride solution.
Add the activated collagenase II solution to the tissue slurry for a final concentration of 250 units per milliliter. Incubate the tubes in a shaker for one hour and at 37 degrees Celsius at 240 x g with manual agitation every 15 minutes to dislodge tissue adhered to the side of the tube. After one hour of incubation, add 100 microliters of collagenase II and 50 microliters of dispase to the sample.
Then, pipette samples 15 to 20 times with a serological pipette until the solution is homogenous. Incubate the sample again for 30 minutes at 37 degrees Celsius and 240 x g with manual agitation every 15 minutes. Slowly shear the muscle solution samples through a 20-milliliter syringe with a 20-gauge needle for 10 cycles.
Then, place a 40-micron cell strainer on a sterile 50-milliliter conical tube and wet it by pipetting five milliliters of DMEM supplemented with 10%FBS and 1%penicillin-streptomycin. Pipette the sample one milliliter at a time through the strainer. Once the whole sample is filtered, wash the cell strainer with DMEM supplemented with 10%FBS and 1%penicillin-streptomycin to bring the total volume of sample to 25 milliliters.
Split the sample volume equally into two 15-milliliter conical tubes and centrifuge at 15 degrees Celsius, 400 x g for 15 minutes. After centrifugation, aspirate the supernatant and resuspend the pellet in one milliliter of RBC lysis buffer at room temperature for seven minutes. Then, add nine milliliters of wash buffer to bring the volume to 10 milliliters and centrifuge at 15 degrees Celsius, 400 x g for 15 minutes.
After centrifugation, aspirate the supernatant and resuspend the pellet in one milliliter of wash buffer. Transfer an appropriate volume of cells to a separate 1.5-milliliter microcentrifuge tube and mix it with trypan blue dye. Count live cells on a light microscope using a hemocytometer.
For flow cytometry, transfer one to 2 million cells per experimental sample to a sterile 1.5-milliliter microcentrifuge tube. Bring the sample volume to one milliliter with wash buffer, and place the tube on ice. Set up all controls for the experiment.
For cell controls, aliquot 500, 000 to 1 million cells in one milliliter of wash buffer in a 1.5-milliliter microcentrifuge tube and place it on ice. If the experiment is being performed for the first time, single-stained cell suspensions should also be included. To set up bead controls, add around 150, 000 positive compensation beads to each labeled 1.5-milliliter centrifuge tube.
Prepare the viability control by transferring half of the cell volume from the viability tube to refresh 1.5-milliliter microcentrifuge tube labeled dead. Incubate the dead tube at 65 degrees Celsius for two to three minutes, then place it on ice. After the incubation, return the dead cells to the viability tube.
Centrifuge the single-cell suspensions, including experimental and control samples at 500 x g and four degrees Celsius for five minutes, and resuspend the cell pellets in 100 microliters of wash buffer after aspirating the supernatant. Add antibodies according to control or experimental condition, and gently flick the sample to ensure complete mixing. Then, incubate them on ice in the dark for 15 minutes.
For compensation beads, incubate the tube at room temperature in the dark for 15 minutes. Bring the volume of each sample to one milliliter by adding 900 microliters of wash buffer to the single-cell suspension, and 900 microliters of PBS for compensation bead controls. Centrifuge single-cell suspensions at 500 x g and four degrees Celsius for five minutes, and compensation bead controls at 300 x g and four degrees Celsius for five minutes.
After centrifugation of the samples, aspirate the supernatant and resuspend the cell pellet in 300 microliters of wash buffer, and bead controls in 300 microliters of PBS and 150, 000 negative compensation beads. Keep the cell samples on ice and bead samples at room temperature under aluminum foil. Proceed with flow cytometry acquisition by setting up the gating strategy to identify fibro-adipogenic progenitors and myogenic progenitors.
In the representative analysis, successful conjugation of the Sca-1 APC Antibody with single staining of compensation beads and cell suspensions generated from healthy rat gastrocnemius muscle was confirmed. Five different concentrations of Sca-1 APC were titrated on single-cell suspensions, and the optimal concentration of antibody was identified based on greatest fluorescence intensity with minimal background staining. Flow cytometric identifications of FAPs and MPs in rat gastrocnemius was performed using the gating strategy shown here.
Samples were first gated to exclude debris in counting beads. Cells were then gated to exclude doublets by both front scatter and side scatter characteristics. The viability of resulting cells was assessed by staining with SYTOX Blue.
SYTOX Blue negative singlets were assessed for CD31 and CD45 to exclude Lin+fractions. The Lin-population was assessed. Cells that were single positive for Sca-1 were designated FAPs, and cells that were single positive for VCAM-1 were designated MPs.
With co-immunostaining immediately after sorting, the freshly isolated population of FAPs displayed positive staining for PDGFRalpha with no contamination by Pax7 positive cells. Conversely, the sorted population of MPs stained positive for Pax7 with an absence of PDGFRalpha-positive cells. FAPs demonstrated differentiated fibroblasts and adipocytes at day 12 in adipogenic differentiation media with expression of fibroblast-specific protein 1 and perilipin 1, respectively.
The presence of neutral triglycerides and lipids from mature adipocytes was evident with Oil Red O staining. Additionally, the FAPs demonstrated a presence of collagen type I and an absence of contaminating myocytes. MPs grown in myogenic differentiation media on day 12 displayed the presence of mature myocytes and fused multinucleated myotubes, and were clear of fibroblasts and adipocyte contamination.
When using this technique to isolate live cells for long-term culture, researchers must ensure to use aseptic technique while simultaneously working efficiently to ensure good cell viability.
