BMPR-IB +快速分离脂肪来源的使用基质细胞在颅骨缺损愈合模型

The overall goal of this protocol is to isolate specific cell populations of adipose-derived stromal cells, and test their osteogenic activity using an in vivo assay. Method can be used to answer key questions in the field of craniofacial regenerative biology, such as how adipose-derived stem cells can be used to create fully functional bone tissue in damaged bone defects. The main advantage of this technique is that the cells are rapidly isolated from fresh human tissue, this way the cells avoid spending time in cell culture, which is known to alter gene and cell service marker expression.

The implications of this technique extend to patients that have large bone defects, as caused by cancer, trauma, or infection. In the future it might be possible to isolate cell populations of ASCs and use them in a regenerative fashion. Visual demonstration of this technique is critical, as creation of the calvarial defect is difficult to learn, mainly because the defect is created on a domed skull, and because it's very close to the dura mater.

For this protocol, have abdominal, flank, or thigh subcutaneous fat collected from a healthy human donor, and stored in a plastic suction canister. To the fat, add an equal volume of sterile PBS, and mix it with gentle agitation for 30 seconds. Allow the aqueous layer to settle, and then remove it using a 10 milliliter plastic pipette.

Then, decant the fat into a large plastic container, and add an equal volume of digest mixture. Clean the sealed fat container with 70%ethanol, and seal it with paraffin. Then, shake the container at 180 rotations per minute, in a 37 degrees Celsius incubator for 30 minutes.

Next, neutralize the digestion by doubling the reaction volume with standard medium. Spin down the cells, carefully remove the supernatant, and resuspend each pellet in 5 milliliters of standard medium. Then, pass all the suspensions through a 100 micron filter into a 50 milliliter conical.

Centrifuge the filtrate at four degrees Celsius for 15 minutes at 300 G.Remove the supernatant and resuspend the pellet in five milliliters of room temperature RBC lysis buffer. Once in a suspension, allow for five minutes of lysis at room temperature. Then, spin the tube down at room temperature for 15 minutes, and complete the separation steps as per the text protocol.

Proper cell sorting is critical to this protocol. First, resuspend the cells in FACS buffer at one million cells per 100 microliters, and keep them on ice. Then, transfer at least 10 million cells to a plastic centrifuge tube on ice labeled BMPR-1B.

Transfer one million cells to a tube on ice labeled unstained, and add one milliliter FACS buffer to them. Label the remaining cells as unsorted. Next, add the antibody to the BMPR-1B tube and mix with gentle trituration to mix.

Allow this mixture to incubate for the amount of time suggested by the manufacturer. Then, spin down the cells for FACS sorting, and carefully resuspend them in one milliliter FACS buffer. Then, do it again and resuspend the cells at one million cells per milliliter.

Now, pass all three cell populations through 40 micron cell strainers into glass FACS tubes. Use 500 microliters FACS buffer to rinse the last cells off the filters. Now, prepare two collection vials, for the antibody positive and negative populations.

Load each with two milliliters of standard cell culture medium, at the FACS machine, first run the unstained cells, and use the results to define a negative gate for the fluorescent marker. Then, using a 100 micron nozzle, sort the stained cells into their respective tubes containing standard medium. If possible, keep these tubes chilled at four degrees Celsius during the sort.

Proceed by loading the cells into the scaffold as described in the text protocol. After anesthetizing the animal, clip any hair from the surgical site and scrub the exposed skin with povidone iodine solution followed by 70%ethanol, three times. Then, drape with the animal, leaving the surgical site exposed.

Now, using 15 blade scalpel, make a sagittal midline incision that extends over the majority of the dorsal skull. Then, with fine-toothed forceps, retract the skin on the right side of the incision to expose the right parietal bone. Set up the drill with an autoclaved four millimeter diamond-coated trephine drill bit.

Then, drill a defect through the parietal bone, but do not extend the defect past the bone, leave the dura mater layer untouched. Now, place a scaffold containing cells or control solution into the defect, and close the skin incision with running nylon suture. Then, proceed with the post-operative mouse care protocols as directed in the text protocol.

Micro CT scans of the skull defect performed on the day of surgery, clearly show the new defect. Repeat scans at later time points show the defects seeded with BMPR-1B positive cells close more rapidly than BMPR-1B negative, or unsorted cells. When the healing is quantified as a percentage of complete wound closure, the difference is easily appreciated.

Next, portions of the skulls containing the defects were decalcified and stained to identify new and oncontrol bone regenerate by using Movat's pentachrome, which distinguishes cartilage from bone, and all the maturation stages in between. Compared to unsorted cells, BMPR-1B positive cells had strong impact on the regenerative process, most notable along the defect borders. After watching this video, you should have a good understanding of the clinical potential that this experiment holds, in isolating patient-specific ASCs and using them to heal bone defects.

Many downstream experiments can come from this to answer even more osteology healing questions, like FACS for use of isolation of ASCs with different lineage commitments, micro CT to analyze how different cell types and tissues heal over time, and the cranial defect model in this experiment to brainstorm for more osteology healing experiments. Don't forget that working with human tissue can be extremely hazardous. Make sure to take appropriate precautions including wearing gloves, using eye protection, and disposing of all hazardous waste properly.