Rapid Isolation of BMPR-IB+ Adipose-Derived Stromal Cells for Use in a Calvarial Defect Healing Model

NOTE: Samples were obtained from patients who gave informed consent. All protocols were reviewed and approved by the appropriate Stanford University Institutional Review Board. While handling human tissue and cells, always adhere to Biosafety Level 2 (BSL2) precautions, as specified by your institution.

1. Preparation of Reagents

1. Prepare FACS buffer: Add 10 mL FBS, 5 mL Poloxamer 188 and 5 mL Pen-Strep to 500 mL sterile phosphate-buffered saline (PBS).
2. Prepare digest mixture: Add 0.375 g C. hemolyticum collagenase type II powder and 5 mL Poloxamer 188 to 500 mL sterile 199/EBSS medium.
3. Prepare standard medium: Add 50 mL FBS and 5 mL Pen-Strep to 500 mL Dulbecco's Modified Eagle Medium.

2. Harvesting and Isolation of ASCs

NOTE: Ensure that adequate institutional approvals are in place for using human tissue and for isolating human stem cells. Obtain human abdominal, flank, or thigh subcutaneous fat from a healthy donor undergoing elective liposuction. Keep the fat in a plastic suction canister.

1. Add sterile PBS in a 1:1 ratio to the fat in the original plastic canister. (If there is 500 mL fat, add 500 mL PBS). Mix by gentle agitation for 30 s. Allow the aqueous layer to settle to the bottom (Figure 1) and then aspirate and discard the aqueous layer with a 10 mL plastic pipette attached to suction.
2. Decant the fat into a large plastic container and add digest mixture in a 1:1 ratio. Close the container and clean the outside with 70% ethanol. Seal the cap with paraffin film.
3. Agitate this container in an orbital shaker at 180 rpm at 37 °C for 30 min.
4. Neutralize the digestion by adding standard medium in a 1:1 ratio. (If there is 1,000 mL fat mixture, add 1,000 mL standard medium).
5. Distribute the mixture equally into an even number of 250 mL plastic conical centrifuge tubes and centrifuge the mixture at 300 x g for 20 min at 4° C.
6. Aspirate the supernatant and be careful to leave the pellet intact. Resuspend pellets in 5 mL standard medium for each conical tube.
7. Filter the suspension through a 100 micron filter into one 50 mL conical centrifuge tube. Then, centrifuge at 300 x g for 15 min at 4 °C.
8. Aspirate the supernatant and resuspend the pellet in 5 mL room temperature RBC lysis buffer. Allow the mixture to sit for 5 min at room temperature then centrifuge at 300 x g for 15 min at room temperature (RT).
9. Aspirate the supernatant and resuspend the pellet in 15 mL standard medium. Again filter through a 100 micron filter into a 50 mL conical centrifuge tube.
10. In a new 50 mL conical, add 15 mL polysucrose solution designed for density-based separation (see list of reagents). Then, holding the tube at a 45° angle, carefully pipette the cells onto the side of the tube so that the cell suspension gently layers on top of the polysucrose solution (Figure 2).
    NOTE: It is important to pipette cells onto the surface of the solution. Do not add polysucrose solution to a suspension of cells as this will create an irreversible mixture. Similarly, do not pipette the cells into the center of the polysucrose solution.
11. Centrifuge this mixture at 300 x g for 10 min at RT. Use low acceleration (2 – 3 out of 10) and set the brake function of the centrifuge to zero for this step.
    NOTE: Three layers will be visible: clear on the bottom, cloudy in the middle, and salmon-colored on the top. The middle layer contains the cells of interest.
12. Using a 10 mL pipette carefully remove the middle layer and transfer to a new 50 mL conical tube.
    NOTE: It is better to completely transfer the middle layer and in the process take some of the top and bottom layers than to leave behind some of the middle layer.
13. Count the cells using a hemocytometer and determine total live cell number.
    NOTE: We typically use trypan blue added to the cell mixture to a final concentration of 0.8% to assist in counting cells. Viable live cells will not take up the blue dye and will appear white.

3. FACS Sorting for BMPR-IB Positive Cells and Preparation of Cell-containing Scaffolds

1. Centrifuge the cell suspension at 300 x g for 5 min at 4 °C. Resuspend the cells in FACS buffer at a concentration of 1 million cells per 100 µL (based on the cell count done in step 2.13).
2. Transfer at least 10 million cells to a plastic centrifuge tube labeled "BMPR-IB." Separately, transfer one million cells in 100 µL FACS buffer to a separate plastic centrifuge tube labeled "Unstained." Add 1 mL FACS buffer to the "Unstained" tube. Keep the remainder of the cells in FACS buffer on ice while the FACS sorting portion of the experiment is being done. Label these cells "Unsorted." Use these as controls later in the experiment.
3. Add an appropriate amount of a fluorescent anti-human BMPR-IB antibody to the "BMPR-IB" tube according to the manufacturer's instructions. Pipette the mixture up and down gently to distribute the antibody.
   NOTE: We use a Human BMPR-IB/ALK-6 APC-conjugated Antibody at a 1:10 dilution (see list of reagents for details). However, antibodies from different manufacturers will have different recommended concentrations.
4. Cover the ice bucket in a way that keeps out light. Allow the cell/antibody mixture to sit for 30 min (or according to the instructions of the antibody manufacturer).
   NOTE: If the anti-BMPR-IB antibody comes as a primary unconjugated antibody that requires a secondary antibody, perform a separate staining step on ice with the secondary antibody according to the manufacturer's instructions.
5. Centrifuge both tubes at 300 x g for 5 min at 4 °C. Aspirate the supernatant, being careful not to aspirate the pellet. Resuspend the pelleted cells in 1 mL FACS buffer. Again centrifuge the tubes at 300 x g for 5 min. Carefully aspirate the supernatant and resuspend the cells in FACS buffer to a concentration of 1 million cells per 1 mL.
6. Filter the "BMPR-IB," "Unstained," and "Unsorted" cells through 40 micron cell strainers into new labeled glass FACS tubes. Rinse the filters with 500 μL FACS buffer to ensure that cells are not left behind in the filter. Also add 2 mL standard medium to two separate plastic centrifuge tubes labeled "BMPR-IB-positive" and "BMPR-IB-negative." Use these two to collect the sorted cells in the FACS machine.
7. On the FACS machine, use the unstained cells to define a negative gate for the fluorescent marker.
8. Using a 100 micron nozzle, sort BMPR-IB positive and negative cells into the respective tubes containing standard medium. Keep these tubes chilled at 4 °C during the sort if the FACS machine has this ability.
   NOTE: Please refer to the JOVE article¹³ by Sharon et al. for an excellent protocol for FACS sorting.
9. Using a hemocytometer, count the cells in each tube. Centrifuge the "BMPR-IB-positive," "BMPR-IB-negative," and "Unsorted" tubes at 300 x g for 5 min at 4 °C and aspirate the supernatant, being careful not to disturb the cell pellet. Then resuspend the cells in standard medium to a concentration of 1 million cells per 250 µL.
10. Obtain pre-cut 4 mm PLGA (poly[lactic-co-glycolic acid])scaffolds coated with hydroxyapatite. Place each scaffold into a well of a 24-well plate. Cover each scaffold with 50 µL of cell suspension (i.e., 200,000 cells) from one of the three groups: Unsorted, BMPR-IB-positive, and BMPR-IB-negative (Figure 3).
    NOTE: Please refer to the JOVE article¹⁴ by Lo et al. for details on the construction of PLGA scaffolds.
11. Partially fill the surrounding empty wells with standard medium (to prevent desiccation of the scaffolds), cover the plate with its lid, and incubate in a cell culture incubator at 37° C for 30 min.

4. Creation of Calvarial Defects and Application of ACS-containing Scaffold

NOTE: Ensure that adequate institutional approvals are in place for the creation of calvarial defects in live mice. This protocol has been approved by the Stanford University Institutional Animal Care and Use Committee.

1. Anesthetize a CD-1 nude mouse using either inhaled isoflurane gas or an injected long-acting anesthetic cocktail.
   1. To use gas alone, place the mouse into an anesthesia chamber and start the flow of oxygen at 2 – 3 L/min with a 2% concentration of isoflurane. Once the mouse is fully unconscious, place it prone onto a warm water recirculating blanket with an absorbant pad and place its snout into an anesthesia nose cone with the same concentration of oxygen and isoflurane. Carefully monitor the respiratory rate and adjust the concentration of isoflurane as needed.
   2. To use an injected long-acting anesthetic cocktail, anesthetize the mouse in an anesthesia chamber as described above and then inject the anesthetic cocktail. Remove the mouse from the anesthesia chamber and observe its behavior.
      NOTE: The mouse may briefly emerge from anesthesia but within several minutes should be again fully anesthetized under the effect of the injected anesthetic. Transfer the mouse to a warm water recirculating blanket covered with an absorbant pad.
      NOTE: For an anesthetic cocktail, we recommend a mixture containing ketamine 10 mg/mL and xylazine 1 mg/mL in normal saline, delivered intraperitoneally. The standard dose of this mixture is 300 µL for a 30 g mouse, which equates to ketamine 100 mg/kg and xylazine 10 mg/kg. See the Discussion section for more details regarding the injectable anesthetic.
   3. Use a toe pinch maneuver to determine the depth of anesthesia. An adequately anesthetized mouse will not retract its paw when it is lightly pinched by the surgeon.
   4. Apply eye ointment to prevent desiccation of the cornea.
   5. Administer 1 mg/kg buprenorphine SR subcutaneously.
      NOTE: Providing analgesia prior to the surgery results in superior pain relief.
   6. During the entire procedure, monitor the respiratory rate of the mouse.
      NOTE: A mouse properly anesthetized under isoflurane should have a respiratory rate of 50 – 100 breaths/min. An increased respiratory rate indicates that anesthesia is too light, while a decreased respiratory rate may indicate that anesthesia is too deep.
2. Prep the skin of the dorsal aspect of the skull with povidone-iodine solution followed by 70% ethanol three times. Drape with sterile drapes, leaving the surgical site exposed.
   NOTE: Wear sterile surgical gloves and maintain sterile technique during the procedure. Only touch sterile surfaces and objects such as the sterile drape and the autoclaved instruments. Have an assistant adjust lighting, open suture packets, etc.
3. Using a 15-blade scalpel, make a sagittal midline incision that extends over the majority of the dorsal skull. Using a fine toothed forceps, retract the skin on the right side of the incision to expose the right parietal bone.
4. Using a drill with an autoclaved 4 mm diamond-coated trephine drill bit, drill a defect through the parietal bone. Do not extend the defect past bone into the dura mater layer. (Figure 4)
5. Place a scaffold into the defect, and then close the skin incision with running nylon suture.
6. Monitor the mouse and provide standard postoperative care according to institutional guidelines.
   1. Keep the mouse on a clean paper towel inside a clean cage while it recovers. One half of the cage should be placed over a warm water recirculating blanket and the mouse should initially be placed in this half.
   2. Do not leave a mouse unattended until it has regained sufficient consciousness to ambulate. Do not return a mouse that has undergone surgery to a cage with other mice until it is fully recovered.
7. Repeat the above steps with a new mouse for each scaffold that is to be tested. Sterilize surgical instruments in between surgeries using a hot bead sterilizer or other suitable method.
8. Shortly after the procedure, and then at 2, 4, 6, and 8 weeks, use micro CT scanning to analyze the rate of calvarial defect closure.
   NOTE: See the article by Levi et al.⁶ for a description of micro CT scan of calvarial defects.
9. When the experiment is complete, euthanize the mice by placing them into a clean euthanasia chamber and starting a flow of carbon dioxide gas into the chamber at a rate of 2 L/min. After respirations have ceased completely (after roughly 10 min) perform cervical dislocation to confirm euthanasia.
10. Optionally, after the experiment is complete, use sectioning and histological staining to assess bone formation within the defect.
    NOTE: See the article by McArdle et al.¹² for details of preparing bone specimens for histology. For a review of histological and staining techniques, see a histological manual such as the Manual of Surgical Pathology¹⁵.