Our overarching vision is to better understand and grasp cell communication and cell behavior, leading to tissue homeostasis and how the cellular processes are misregulated in FAP repair. To do so, we are using various models, of which one is the spiny mouse. One of the main issues with this new mouse model is the lack of tools to study it.
Here we provide a new protocol, which contains reliable enzyme digestion and antibody staining, allowing us to successfully sort fibro-adipogenic progenitors and culture them in vitro for further analysis of their fate. We have tested various enzyme combinations and concentrations as well as multiple antibodies, and our method provides references for the most reliable and robust protocol. As we now have a reliable enzymatic digestion we will continue to test for other markers and antibodies to successfully analyze endothelial cells, pericytes, immune cells, and myogenic cells.
To begin, place a euthanized mouse on a dissection stage in a supine position. Thoroughly spray the mouse's fur with 70%ethanol. Make a two to three centimeter long vertical incision at the ventral midline.
Expose the ribcage and the abdominal muscle. Then, use tissue scissors to cut through the muscle at the xiphoid process and expose the diaphragm. Now, cut the diaphragm and rib cage on both sides.
Clamp and peel back the cut ribs with a hemostat. Nick the right atrium with tissue scissors. Then insert a 23-gauge needle attached to a 20 mL syringe into the apex of the heart.
Slowly inject 20 mL of 2 mM PBS EDTA into the heart. Excise it and place it in a 60 mm Petri dish containing cold PBS on ice. Remove the atria and rinse out as much blood as possible.
Next, remove the skin above the knee joint of the mouse. With a pair of fine forceps, remove the fascia. Use scissors to isolate the quadriceps with the femur as a guide.
Then transfer the quadricep muscle to a separate Petri dish containing cold PBS. To begin, add two mL of digestion buffer to a 5 mL transport vial containing the mouse heart tissue, and 4 mL of digestion buffer into a 15 mL centrifuge tube containing the mouse skeletal tissue. Place the heart and quadricep muscle tissue isolated from a spiny mouse onto the lid of a Petri dish.
With a pair of tissue scissors, mince it into pieces smaller than 1 cubic mm. Transfer the minced tissue into the respective digestion tubes. Vortex the contents of the tubes for two seconds at low speed.
After 20 minutes, remove the tubes off the rotator. Allow the tissue pieces to settle. Use a P1000 pipette to aspirate the supernatant.
Transfer the supernatant into 50 mL centrifuge tubes containing 20 mL of ice cold FACS buffer. Top up the digestion tubes with the respective volumes of digestion buffer. Incubate them on the rotator again.
After subjecting the muscles to another round of digestion, add ice cold FACS buffer to quench the digestion. Place a 40 micrometer cell strainer on top of a 50 mL centrifuge tube. Filter the digestion suspension and supernatant through the strainer.
Centrifuge the cell suspension at 500 G for eight minutes at 4 C.After removing the supernatant, add 3 mL of ACK lysis buffer to the pellet and resuspend. Incubate the suspension for five minutes on ice. Top up the volume to 40 mL with FACS buffer.
Centrifuge the samples again at 500 G for five minutes at 4 C.After decanting the supernatant, resuspend the pellet in 0.5 mL of FACS buffer. Finally, filter the suspension through a 40 micrometer cell strainer cap placed on a 5 mL polystyrene round bottom tube. To begin, prepare 30 L of staining buffer at 2x working concentrations for single color and FMO controls.
Transfer the single color and FMO control staining buffer to the wells of a 96-well plate. Top up the volume with 20 L of FACS buffer. To stain the sample, prepare 0.5 mL of 2x FACS staining buffer in 1.5 mL centrifuge tubes.
Pipette 10 L of the cell suspension to single color and FMO control wells. Then add 2x FACS staining buffer to the rest of the cell suspension and incubate. Next, add 100 L of FACS buffer into the wells and two mL of the buffer into the sample tubes.
Centrifuge at 500 G for five minutes at 4 C.Aspirate the supernatant, then resuspend the cell pellet in viability buffer. Add 300 L of proliferation media into 1.5 mL centrifuge tubes. These will act as collection tubes for cell sorting.
After sorting the cells using FACS, centrifuge the collected cells at 800 G for 10 minutes at 4 C.Use a P1000 pipette to carefully remove the supernatant without disturbing the pellet. Resuspend the pellet in proliferation media to a final density of 100 cells per microliter. Seed the cell suspension into the wells of a 48-well tissue culture plate.
Finally, transfer the plate to an incubator at 37 C under respective gas supplementation until cells are confluent. Confluent cells with typical fibroblast morphology were obtained within five days of culture.
