C.D.M. was supported by the American College of Surgeons (ACS) Resident Research Scholarship. M.S.H. was supported by the California Institute for Regenerative Medicine (CIRM) Clinical Fellow training grant TG2-01159. M.S.H., H.P.L., and M.T.L. were supported by the American Society of Maxillofacial Surgeons (ASMS)/Maxillofacial Surgeons Foundation (MSF) Research Grant Award. H.P.L. was supported by NIH grant R01 GM087609 and a gift from Ingrid Lai and Bill Shu in honor of Anthony Shu. H.P.L. and M.T.L. were supported by the Hagey Laboratory for Pediatric Regenerative Medicine and The Oak Foundation. M.T.L. was supported by NIH grants U01 HL099776, R01 DE021683-01, and RC2 DE020771. D.C.W. was supported by NIH grant 1K08DE024269, the Hagey Laboratory for Pediatric Regenerative Medicine, and the Stanford University Child Health Research Institute Faculty Scholar Award.

NOTE: Samples were obtained from patients who gave informed consent. All protocols were reviewed and approved by the appropriate Stanford University Institutional Review Board.&#160;While handling human tissue and cells, always adhere to Biosafety Level 2 (BSL2) precautions, as specified by your institution.
1. Preparation of Reagents
<ol>
	<li>Prepare FACS buffer: Add 10 mL FBS, 5 mL Poloxamer 188 and 5 mL Pen-Strep to 500 mL sterile phosphate-buffered saline (PBS).</li>
	<li>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.</li>
	<li>Prepare standard medium: Add 50 mL FBS and 5 mL Pen-Strep to 500 mL Dulbecco&#39;s Modified Eagle Medium.</li>
</ol>
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.
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Agitate this container in an orbital shaker at 180 rpm at 37 &#176;C for 30 min.</li>
	<li>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).</li>
	<li>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&#176; C.</li>
	<li>Aspirate the supernatant and be careful to leave the pellet intact. Resuspend pellets in 5 mL standard medium for each conical tube.</li>
	<li>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 &#176;C.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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&#176; 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
3. FACS Sorting for BMPR-IB Positive Cells and Preparation of Cell-containing Scaffolds
<ol>
	<li>Centrifuge the cell suspension at 300 x g for 5 min at 4 &#176;C. Resuspend the cells in FACS buffer at a concentration of 1 million cells per 100 &#181;L (based on the cell count done in step 2.13).</li>
	<li>Transfer at least 10 million cells to a plastic centrifuge tube labeled &#34;BMPR-IB.&#34; Separately, transfer one million cells in 100 &#181;L FACS buffer to a separate plastic centrifuge tube labeled &#34;Unstained.&#34; Add 1 mL FACS buffer to the &#34;Unstained&#34; 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 &#34;Unsorted.&#34; Use these as controls later in the experiment.</li>
	<li>Add an appropriate amount of a fluorescent anti-human BMPR-IB antibody to the &#34;BMPR-IB&#34; tube according to the manufacturer&#39;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.</li>
	<li>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&#39;s instructions.</li>
	<li>Centrifuge both tubes at 300 x g for 5 min at 4 &#176;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.</li>
	<li>Filter the &#34;BMPR-IB,&#34; &#34;Unstained,&#34; and &#34;Unsorted&#34; cells through 40 micron cell strainers into new labeled glass FACS tubes. Rinse the filters with 500 &#956;L 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 &#34;BMPR-IB-positive&#34; and &#34;BMPR-IB-negative.&#34; Use these two to collect the sorted cells in the FACS machine.</li>
	<li>On the FACS machine, use the unstained cells to define a negative gate for the fluorescent marker.</li>
	<li>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 &#176;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.</li>
	<li>Using a hemocytometer, count the cells in each tube. Centrifuge the &#34;BMPR-IB-positive,&#34; &#34;BMPR-IB-negative,&#34; and &#34;Unsorted&#34; tubes at 300 x g for 5 min at 4 &#176;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 &#181;L.</li>
	<li>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 &#181;L 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.</li>
	<li>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&#176; C for 30 min.</li>
</ol>
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.
<ol>
	<li>Anesthetize a CD-1 nude mouse using either inhaled isoflurane gas or an injected long-acting anesthetic cocktail.
	<ol>
		<li>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&#160;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.</li>
		<li>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&#160;covered with an absorbant pad. 
		NOTE:&#160;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 &#181;L 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.</li>
		<li>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.</li>
		<li>Apply eye ointment to prevent desiccation of the cornea.</li>
		<li>Administer 1 mg/kg buprenorphine SR subcutaneously. 
		NOTE: Providing analgesia prior to the surgery results in superior pain relief.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>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)</li>
	<li>Place a scaffold into the defect, and then close the skin incision with running nylon suture.</li>
	<li>Monitor the mouse and provide standard postoperative care according to institutional guidelines.
	<ol>
		<li>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&#160;and the mouse should initially be placed in this half.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

<ol><li>Silber, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Donor+site+morbidity+after+anterior+iliac+crest+bone+harvest+for+single-level+anterior+cervical+discectomy+and+fusion%5BTitle%5D)+AND+%22Spine+%28Phila+Pa+1976%29%22%5BJournal%5D)">Donor site morbidity after anterior iliac crest bone harvest for single-level anterior cervical discectomy and fusion.</a> Spine (Phila Pa 1976). 28 (2), 134-139 (2003).</li>
<li>Giannoudis, P. V., Dinopoulos, H., Tsiridis, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bone+substitutes%3A+an+update%5BTitle%5D)+AND+%22Injury%22%5BJournal%5D)">Bone substitutes: an update.</a> Injury. 36, S20-S27 (2005).</li>
<li>Laurencin, C., Khan, Y., El-Amin, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bone+graft+substitutes%5BTitle%5D)+AND+%22Expert+Rev+Med+Devices%22%5BJournal%5D)">Bone graft substitutes.</a> Expert Rev Med Devices. 3 (1), 49-57 (2006).</li>
<li>Walmsley, G. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Nanotechnology+in+bone+tissue+engineering%5BTitle%5D)+AND+%22Nanomedicine%22%5BJournal%5D)">Nanotechnology in bone tissue engineering.</a> Nanomedicine. 11 (5), 1253-1263 (2015).</li>
<li>Chapekar, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tissue+engineering%3A+challenges+and+opportunities%5BTitle%5D)+AND+%22J+Biomed+Mater+Res%22%5BJournal%5D)">Tissue engineering: challenges and opportunities.</a> J Biomed Mater Res. 53 (6), 617-620 (2000).</li>
<li>Levi, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Human+adipose+derived+stromal+cells+heal+critical+size+mouse+calvarial+defects%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">Human adipose derived stromal cells heal critical size mouse calvarial defects.</a> PLoS One. 5 (6), e11177(2010).</li>
<li>Gimble, J. M., Katz, A. J., Bunnell, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Adipose-derived+stem+cells+for+regenerative+medicine%5BTitle%5D)+AND+%22Circ+Res%22%5BJournal%5D)">Adipose-derived stem cells for regenerative medicine.</a> Circ Res. 100 (9), 1249-1260 (2007).</li>
<li>Levi, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CD105+protein+depletion+enhances+human+adipose-derived+stromal+cell+osteogenesis+through+reduction+of+transforming+growth+factor+beta1+%28TGF-beta1%29+signaling%5BTitle%5D)+AND+%22J+Biol+Chem%22%5BJournal%5D)">CD105 protein depletion enhances human adipose-derived stromal cell osteogenesis through reduction of transforming growth factor beta1 (TGF-beta1) signaling.</a> J Biol Chem. 286 (45), 39497-39509 (2011).</li>
<li>Chung, M. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CD90+%28Thy-1%29-positive+selection+enhances+osteogenic+capacity+of+human+adipose-derived+stromal+cells%5BTitle%5D)+AND+%22Tissue+Eng+Part+A%22%5BJournal%5D)">CD90 (Thy-1)-positive selection enhances osteogenic capacity of human adipose-derived stromal cells.</a> Tissue Eng Part A. 19 (7-8), 989-997 (2013).</li>
<li>Wozney, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Novel+regulators+of+bone+formation%3A+molecular+clones+and+activities%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Novel regulators of bone formation: molecular clones and activities.</a> Science. 242 (4885), 1528-1534 (1988).</li>
<li>Wan, D. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Osteogenic+differentiation+of+mouse+adipose-derived+adult+stromal+cells+requires+retinoic+acid+and+bone+morphogenetic+protein+receptor+type+IB+signaling%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Osteogenic differentiation of mouse adipose-derived adult stromal cells requires retinoic acid and bone morphogenetic protein receptor type IB signaling.</a> Proc Natl Acad Sci U S A. 103 (33), 12335-12340 (2006).</li>
<li>McArdle, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Positive+selection+for+bone+morphogenetic+protein+receptor+type-IB+promotes+differentiation+and+specification+of+human+adipose-derived+stromal+cells+toward+an+osteogenic+lineage%5BTitle%5D)+AND+%22Tissue+Eng+Part+A%22%5BJournal%5D)">Positive selection for bone morphogenetic protein receptor type-IB promotes differentiation and specification of human adipose-derived stromal cells toward an osteogenic lineage.</a> Tissue Eng Part A. 20 (21-22), 3031-3040 (2014).</li>
<li>Sharon, Y., Alon, L., Glanz, S., Servais, C., Erez, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isolation+of+normal+and+cancer-associated+fibroblasts+from+fresh+tissues+by+Fluorescence+Activated+Cell+Sorting+%28FACS%29%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Isolation of normal and cancer-associated fibroblasts from fresh tissues by Fluorescence Activated Cell Sorting (FACS).</a> J Vis Exp. (71), e4425(2013).</li>
<li>Lo, D. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Repair+of+a+critical-sized+calvarial+defect+model+using+adipose-derived+stromal+cells+harvested+from+lipoaspirate%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Repair of a critical-sized calvarial defect model using adipose-derived stromal cells harvested from lipoaspirate.</a> J Vis Exp. (68), (2012).</li>
<li>Lester, S. C. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aLester%2C+SC+%22Manual+of+surgical+pathology%22">Manual of surgical pathology</a>. , Saunders/Elsevier. (2010).</li>
<li>Bunnell, B. A., Flaat, M., Gagliardi, C., Patel, B., Ripoll, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Adipose-derived+stem+cells%3A+isolation%2C+expansion+and+differentiation%5BTitle%5D)+AND+%22Methods%22%5BJournal%5D)">Adipose-derived stem cells: isolation, expansion and differentiation.</a> Methods. 45 (2), 115-120 (2008).</li>
</ol>

The authors have no disclosures to make.

Critical Steps within the Protocol
During the harvest of ASCs, the critical step is adequate digestion of fat with collagenase. Inadequate digestion will result in a low yield of ASCs. During FACS sorting of BMPR-IB+ cells, it is important to carefully define the gate for positivity. Defining gates too loosely may result in sorted populations that are not pure. During creation of the calvarial defect, it is critical to drill the defect through the bone of the skull but to not advance into the dur...

Major bone defects resulting from injury, infection, or invasive cancer have a significant impact on a patient&#39;s recovery and quality of life. Techniques exist to fill these defects with healthy bone from elsewhere in the patient&#39;s own body, but this transfer carries its own morbidity and risk of complications1,2,3. Furthermore, some defects are so large or complex that sufficient donor bone is not available to fill the defect. Prosthetic devices are a potential option for filling bony defects but these are associated with several disadvantages including infection risk, hardware failure, and foreign body reaction4.
For these reasons there is great interest in the possibility of engineering biological bone substitutes using a patient&#39;s own cells5. Adipose-derived stromal cells (ASCs) have potential for this application because they are abundantly available in the patient&#39;s own fat tissue and they have demonstrated the ability to heal bone defects by generating new bone tissue6,7. ASCs are a diverse population of cells and several studies have shown that selecting for specific cell surface markers can produce cell populations with enhanced osteogenic activity8,9. Selecting ASCs with the highest osteogenic potential would increase the likelihood that a scaffold seeded with these cells could regenerate a large bone defect.
Bone morphogenetic protein (BMP) signaling is critical for regulating bone differentiation and formation10 and the BMP Receptor type IB (BMPR-IB) is known to be important for osteogenesis in ASCs11. Recently, we have shown that expression of BMPR-IB can be used to select for ASCs with enhanced osteogenic activity12. Here we demonstrate a protocol for the isolation of BMPR-IB-expressing ASCs from human fat followed by an assay of their osteogenic activity using an in vivo calvarial defect model.

<table><tbody><tr><td>100 micron cell strainer</td><td>Falcon</td><td>352360</td><td></td></tr><tr><td>15 blade scalpel</td><td>Miltex</td><td>4-515</td><td></td></tr><tr><td>24 well plate</td><td>Corning</td><td>3524</td><td></td></tr><tr><td>40 micron cell strainer</td><td>Falcon</td><td>352340</td><td></td></tr><tr><td>50 mL conical centrifuge tubes</td><td>Falcon</td><td>352098</td><td></td></tr><tr><td>6-0 Ethilon nylon suture, 18&quot;, P-3 needle,&amp;nbsp;</td><td>Ethicon</td><td>1698G</td><td></td></tr><tr><td>Anti-BMPR-IB primary antibody</td><td>R&amp;amp;D systems</td><td>FAB5051A</td><td></td></tr><tr><td>BioGel PI surgical gloves</td><td>M&amp;ouml;lnlycke Health Care</td><td>ALA42675Z</td><td></td></tr><tr><td>Buprenorphine SR</td><td>ZooPharm</td><td></td><td></td></tr><tr><td>Castro-Viejo needle driver</td><td>Fine Science Tools</td><td>12565-14</td><td></td></tr><tr><td>CD1 nude mouse</td><td>Charles River</td><td>086</td><td></td></tr><tr><td>Collagenase Type II powder</td><td>Gibco</td><td>17101-015</td><td></td></tr><tr><td>DMEM medium</td><td>Gibco</td><td>10564-011</td><td></td></tr><tr><td>Drill: Circular knife 4.0 mm</td><td>Xemax Surgical</td><td>CK40</td><td></td></tr><tr><td>Drill: Z500 Brushless Micromotor</td><td>NSK</td><td>NSKZ500</td><td></td></tr><tr><td>FBS</td><td>Gicbo</td><td>10437-077</td><td></td></tr><tr><td>Fisherbrand Absorbent Underpads, 20&quot; x 24&quot;</td><td>Fisher Scientific</td><td>14-206-62</td><td></td></tr><tr><td>Fisherbrand Sterile cotton gauze pad, 4&quot; x 4&quot;</td><td>Fisher Scientific</td><td>22-415-469</td><td></td></tr><tr><td>Heating pad</td><td>Kent Scientific</td><td>DCT-20</td><td></td></tr><tr><td>Hyclone 199/EBSS medium</td><td>GE&amp;nbsp; Life Sciences</td><td>SH30253.01</td><td></td></tr><tr><td>Isothesia isoflurane</td><td>Henry Schein&amp;nbsp;</td><td>050033</td><td></td></tr><tr><td>Micro Forceps with teeth</td><td>Roboz</td><td>RS-5150</td><td></td></tr><tr><td>Micro Forceps with teeth</td><td>Roboz</td><td>RS-5150</td><td></td></tr><tr><td>Paraffin film (Parafilm)</td><td>Bemis</td><td>PM996</td><td></td></tr><tr><td>PBS</td><td>Gibco</td><td>10010-023</td><td></td></tr><tr><td>Pen-Strep</td><td>Gibco</td><td>15140-122</td><td></td></tr><tr><td>PLGA scaffolds</td><td>Proprietary Formulation</td><td></td><td></td></tr><tr><td>Poloxamer 188, 10%</td><td>Sigma</td><td>P5556-100ML</td><td></td></tr><tr><td>Polylined Sterile Field, 18&quot; x 24&quot;</td><td>Busse Hospital Disposables</td><td>696</td><td>Cut a rectangular hole of the appropriate size</td></tr><tr><td>Polysucrose Solution: Histopaque 1119</td><td>Sigma</td><td>11191</td><td></td></tr><tr><td>Povidone Iodine Prep Solution</td><td>Medline</td><td>MDS093944H</td><td></td></tr><tr><td>Puralube petrolatum ophthalmic ointment, 1/8 oz. tube</td><td>Dechra Veterinary Products</td><td></td><td></td></tr><tr><td>RBC lysis buffer</td><td>Sigma</td><td>11814389001</td><td></td></tr><tr><td>Webcol alcohol prep swabs</td><td>Covidien</td><td>6818</td><td></td></tr></tbody></table>

rapid isolation of bmpr-ib+ adipose-derived stromal cells for use in a calvarial defect healing model

Invasive cancers, major injuries, and infection can cause bone defects that are too large to be reconstructed with preexisting bone from the patient's own body. The ability to grow bone de novo using a patient's own cells would allow bony defects to be filled with adequate tissue without the morbidity of harvesting native bone. There is interest in the use of adipose-derived stromal cells (ASCs) as a source for tissue engineering because these are obtained from an abundant source: the patient's own adipose tissue. However, ASCs are a heterogeneous population and some subpopulations may be more effective in this application than others. Isolation of the most osteogenic population of ASCs could improve the efficiency and effectiveness of a bone engineering process. In this protocol, ASCs are obtained from subcutaneous fat tissue from a human donor. The subpopulation of ASCs expressing the marker BMPR-IB is isolated using FACS. These cells are then applied to an in vivo calvarial defect healing assay and are found to have improved osteogenic regenerative potential compared with unsorted cells.

Micro CT scan done on the day of surgery will clearly show the skull defect. At this time there will be no ingrowth into the 4 mm defect. Subsequent scans are obtained over time to quantify the size of the defect over time compared with the baseline. Defects seeded with BMPR-IB+ cells should demonstrate more rapid closure of the defect when compared with BMPR-IB- and Unsorted cells (Figure 5). In addition, the portion of the skull containing the defect can be decalcified ...

Watch this Scientific Journal Video about Rapid Isolation of BMPR-IB+ Adipose-Derived Stromal Cells for Use in a Calvarial Defect Healing Model at JoVE.com

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

Adipose-derived stromal cells may be useful for engineering new tissue from a patient's own cells. We present a protocol for the isolation of a subpopulation of human adipose-derived stromal cells (ASCs) with increased osteogenic potential, followed by application of the cells in an in vivo calvarial healing assay.

Invasive cancers, major injuries, and infection can cause bone defects that are too large to be reconstructed with preexisting bone from the patient's own ...

rapid-isolation-bmpr-ib-adipose-derived-stromal-cells-for-use

Research

JoVE Journal

Developmental Biology

6.7K Views.  Stanford University School of Medicine. Adipose-derived stromal cells may be useful for engineering new tissue from a patient's own cells. We present a protocol for the isolation of a subpopulation of human adipose-derived stromal cells (ASCs) with increased osteogenic potential, followed by application of the cells in an in vivo calvarial healing assay.

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

Isolation and Enrichment of Human Adipose-derived Stromal Cells for Enhanced Osteogenesis

Bone marrow-derived mesenchymal stromal cells (BM-MSCs) are considered the gold standard for stem cell-based tissue engineering applications. However, the process by which they must be harvested can be associated with significant donor site morbidity. In contrast, adipose-derived stromal cells (ASCs) are more readily abundant and more easily harvested, making them an appealing alternative to BM-MSCs. Like BM-MSCs, ASCs can differentiate into osteogenic lineage cells and can be used in tissue engineering applications, such as seeding onto scaffolds for use in craniofacial skeletal defects. ASCs are obtained from the stromal vascular fraction (SVF) of digested adipose tissue, which is a heterogeneous mixture of ASCs, vascular endothelial and mural cells, smooth muscle cells, pericytes, fibroblasts, and circulating cells. Flow cytometric analysis has shown that the surface marker profile for ASCs is similar to that for BM-MSCs. Despite several published reports establishing markers for the ASC phenotype, there is still a lack of consensus over profiles identifying osteoprogenitor cells in this heterogeneous population. This protocol describes how to isolate and use a subpopulation of ASCs with enhanced osteogenic capacity to repair critical-sized calvarial defects.

The transcriptional heterogeneity within human adipose-derived stromal cells can be defined on the single cell level using cell surface markers and osteogenic genes. We describe a protocol utilizing flow cytometry for the isolation of cell subpopulations with increased osteogenic potential, which may be used to enhance craniofacial skeletal reconstruction.

Bone marrow-derived mesenchymal stromal cells (BM-MSCs) are considered the gold standard for stem cell-based tissue engineering applications. However, the ...

Use of Human Perivascular Stem Cells for Bone Regeneration

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are a FACS-sorted population of 'pericytes' (CD146+CD34-CD45-) and 'adventitial cells' (CD146-CD34+CD45-), each of which we have previously reported to have properties of mesenchymal stem cells. PSCs, like MSCs, are able to undergo osteogenic differentiation, as well as secrete pro-osteogenic cytokines1,2. In the present protocol, we demonstrate the osteogenicity of PSCs in several animal models including a muscle pouch implantation in SCID (severe combined immunodeficient) mice, a SCID mouse calvarial defect and a femoral segmental defect (FSD) in athymic rats. The thigh muscle pouch model is used to assess ectopic bone formation. Calvarial defects are centered on the parietal bone and are standardly 4 mm in diameter (critically sized)8. FSDs are bicortical and are stabilized with a polyethylene bar and K-wires4. The FSD described is also a critical size defect, which does not significantly heal on its own4. In contrast, if stem cells or growth factors are added to the defect site, significant bone regeneration can be appreciated. The overall goal of PSC xenografting is to demonstrate the osteogenic capability of this cell type in both ectopic and orthotopic bone regeneration models.

Human perivascular stem cells (PSCs) are a novel stem cell class for skeletal tissue regeneration similar to mesenchymal stem cells (MSCs). PSCs can be isolated by FACS (fluorescence activated cell sorting) from adipose tissue procured during standard liposuction procedures, then combined with an osteoinductive scaffold to achieve bone formation in vivo.

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are ...

Bioengineering

Repair of a Critical-sized Calvarial Defect Model Using Adipose-derived Stromal Cells Harvested from Lipoaspirate

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. Adipose-derived stromal cells (ASCs) have proven to be an abundant source of multipotent stem cells capable of undergoing osteogenic, chondrogenic, adipogenic, and myogenic differentiation. Many studies have explored the osteogenic potential of these cells in vivo with the use of various scaffolding biomaterials for cellular delivery. It has been demonstrated that by utilizing an osteoconductive, hydroxyapatite-coated poly(lactic-co-glycolic acid) (HA-PLGA) scaffold seeded with ASCs, a critical-sized calvarial defect, a defect that is defined by its inability to undergo spontaneous healing over the lifetime of the animal, can be effectively show robust osseous regeneration. This in vivo model demonstrates the basis of translational approaches aimed to regenerate the bone tissue - the cellular component and biological matrix. This method serves as a model for the ultimate clinical application of a progenitor cell towards the repair of a specific tissue defect.

This protocol describes the isolation of adipose-derived stromal cells from lipoaspirate and the creation of a 4 mm critical-sized calvarial defect to evaluate skeletal regeneration.

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. ...

Medicine

Visualizing Angiogenesis by Multiphoton Microscopy In Vivo in Genetically Modified 3D-PLGA/nHAp Scaffold for Calvarial Critical Bone Defect Repair

The reconstruction of critically sized bone defects remains a serious clinical problem because of poor angiogenesis within tissue-engineered scaffolds during repair, which gives rise to a lack of sufficient blood supply and causes necrosis of the new tissues. Rapid vascularization is a vital prerequisite for new tissue survival and integration with existing host tissue. The de novo generation of vasculature in scaffolds is one of the most important steps in making bone regeneration more efficient, allowing repairing tissue to grow into a scaffold. To tackle this problem, the genetic modification of a biomaterial scaffold is used to accelerate angiogenesis and osteogenesis. However, visualizing and tracking in vivo blood vessel formation in real-time and in three-dimensional (3D) scaffolds or new bone tissue is still an obstacle for bone tissue engineering. Multiphoton microscopy (MPM) is a novel bio-imaging modality that can acquire volumetric data from biological structures in a high-resolution and minimally-invasive manner. The objective of this study was to visualize angiogenesis with multiphoton microscopy in vivo in a genetically modified 3D-PLGA/nHAp scaffold for calvarial critical bone defect repair. PLGA/nHAp scaffolds were functionalized for the sustained delivery of a growth factor pdgf-b gene carrying lentiviral vectors (LV-pdgfb) in order to facilitate angiogenesis and to enhance bone regeneration. In a scaffold-implanted calvarial critical bone defect mouse model, the blood vessel areas (BVAs) in PHp scaffolds were significantly higher than in PH scaffolds. Additionally, the expression of pdgf-b and angiogenesis-related genes, vWF and VEGFR2, increased correspondingly. MicroCT analysis indicated that the new bone formation in the PHp group dramatically improved compared to the other groups. To our knowledge, this is the first time multiphoton microscopy was used in bone tissue-engineering to investigate angiogenesis in a 3D bio-degradable scaffold in vivo and in real-time.

Visualizing Angiogenesis by Multiphoton Microscopy In Vivo in Genetically Modified 3D-PLGA/nHAp Scaffold for Calvarial Critical Bone Defect Repair

Here, we present a protocol to visualize blood vessel formation in vivo and in real-time in 3D scaffolds by multiphoton microscopy. Angiogenesis in genetically modified scaffolds was studied in a murine calvarial critical bone defect model. More new blood vessels were detected in the treatment group than in controls.

The reconstruction of critically sized bone defects remains a serious clinical problem because of poor angiogenesis within tissue-engineered scaffolds ...

Biological Compatibility Profile on Biomaterials for Bone Regeneration

Large non-union bone fractures are a significant challenge in orthopedic surgery. Although auto- and allogeneic bone grafts are excellent for healing such lesions, there are potential complications with their use. Thus, material scientists are developing synthetic, biocompatible biomaterials to overcome these problems. In this study, we present a multidisciplinary platform for evaluating biomaterials for bone repair. We combined expertise from bone biology and immunology to develop a platform including in vitro osteoclast (OC) and osteoblast (OB) assays and in vivo mouse models of bone repair, immunogenicity, and allergenicity. We demonstrate how to perform the experiments, summarize the results, and report on biomaterial biocompatibility. In particular, we tested OB viability, differentiation, and mineralization and OC viability and differentiation in the context of &#946;-tricalcium phosphate (&#946;-TCP) disks. We also tested a &#946;-TCP/Collagen (&#946;-TCP/C) foam which is a commercially available material used clinically for bone repair in a critical-sized calvarial bone defect mouse model to determine the effects on the early phase of bone healing. In parallel experiments, we evaluated immune and allergic responses in mice. Our approach generates a biological compatibility profile of a bone biomaterial with a range of parameters necessary for predicting the biocompatibility of biomaterials used for bone healing and repair in patients.

The number of novel biomaterials engineered for repairing large bone lesions is continuously expanding with the aim to enhance bone healing and overcome the complications associated with bone transplantation. Here, we present a multidisciplinary strategy for pre-clinical biocompatibility testing of biomaterials for bone repair.

Large non-union bone fractures are a significant challenge in orthopedic surgery. Although auto- and allogeneic bone grafts are excellent for healing such ...

Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows assessment of bone substitution materials for oral and craniofacial bone regeneration in terms of bone growth and neovascularization support. Once animals are anesthetized and ventilated (endotracheal intubation), four cylinders made of polyether ether ketone (PEEK) are screwed onto the skull, on both sides of the median and coronal sutures. Five intramedullary holes are drilled within the bone area delimited by each cylinder, allowing influx of bone marrow cells. The material samples are placed into the cylinders which are then closed. Finally, the surgical site is sutured, and animals are awaken. Bone growth may be assessed on live animals by using microtomography. Once animals are euthanized, bone growth and neovascularization may be evaluated by using microtomography, immune-histology and immunofluorescence. As the evaluation of a material requires maximum standardization and calibration, the calvarial model appears ideal. Access is very easy, calibration and standardization are facilitated by the use of defined cylinders and four samples may be assessed simultaneously. Furthermore, live tomography may be used and ultimately a large decrease in animals to be euthanized may be anticipated.

Here we present a surgical protocol in rabbits with the aim to assess bone substitution materials in terms of bone regeneration capacities. By using PEEK cylinders fixed onto rabbit skulls, osteoconduction, osteoinduction, osteogenesis and vasculogenesis induced by the materials may be evaluated either on live or euthanized animals.

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows ...

Isolation and Culture of Bone Marrow Mesenchymal Stem Cells from the Human Mandible

Human mesenchymal stem cells (hMSCs) have shown great potential in bone regeneration, immune modulation, and treating refractory chronic diseases. Various origins have been found to obtain hMSCs recently, while bone marrow was still considered the main source. Bone marrow-derived MSCs (BMSCs) from different donor bone sites have distinct characteristics due to microenvironmental factors. Studies have shown that BMSCs from maxillofacial bone may have greater proliferative and osteogenic capacities than BMSCs from long bones or the iliac crest. And maxillofacial BMSCs were considered more suitable for stem cell therapy in the maxillofacial tissues. The mandible, especially the ascending ramal area with sufficient marrow, was a feasible donor site for harvesting BMSCs. This study described a protocol for harvesting, isolating, and culturing human mandibular bone marrow-derived MSCs (hmBMSCs). Furthermore, immunophenotyping of hmBMSCs, proliferation assays, and in vitro induction of osteogenic, adipogenic, and chondrogenic differentiation was performed to identify the cultured cells. Applying this protocol can help the researchers successfully obtain enough high-quality hmBMSCs, which is necessary for further studies of the biological function, microenvironmental effects, and clinical applications.

The present protocol describes an efficient procedure for isolating and culturing of human mandibular bone marrow-derived mesenchymal stem cells using the whole bone marrow adherence method. The cultured cells were identified by cell proliferation assays, flow cytometry, and multilineage differentiation induction.

Human mesenchymal stem cells (hMSCs) have shown great potential in bone regeneration, immune modulation, and treating refractory chronic diseases. Various ...

Isolation and Cultivation of Mandibular Bone Marrow Mesenchymal Stem Cells in Rats

Here we present an efficient method for isolating and culturing mandibular bone marrow mesenchymal stem cells (mBMSCs) in vitro to rapidly obtain numerous high-quality cells for experimental requirements. mBMSCs could be widely used in therapeutic applications as tissue engineering cells in case of craniofacial diseases and cranio-maxillofacial regeneration in the future due to the excellent self-renewal ability and multi-lineage differentiation potential. Therefore, it is important to obtain mBMSCs in large numbers.
In this study, bone marrow was flushed from the mandible and primary mBMSCs were isolated through whole bone marrow adherent cultivation. Furthermore, CD29+CD90+CD45&#8722; mBMSCs were purified through fluorescent cell sorting. The second generation of purified mBMSCs were used for further study and displayed potential in differentiating into osteoblasts, adipocytes, and chondrocytes. Utilizing this in vitro model, one can obtain a high number of proliferative mBMSCs, which may facilitate the study of the biological characteristics, the subsequent reaction to the microenvironment, and other applications of mBMSCs.

This article presents a method that combines whole bone marrow adherence and flow cytometry sorting for isolating, cultivating, sorting, and identifying bone marrow mesenchymal stem cells from rat mandibles.

Here we present an efficient method for isolating and culturing mandibular bone marrow mesenchymal stem cells (mBMSCs) in vitro to rapidly obtain numerous ...

Standardizing Rat Calvarial Suture-Bony Defect Model for Regenerative Studies

The Establishment of Calvarial Suture-Bony Composite Defects in Rats: A Standardized Model for Suture-Regenerative Therapy Investigation

Large-scale calvarial defects often coincide with cranial suture disruption, leading to impairments in calvarial defect restoration and skull development (the latter occurs in the developing cranium). However, the lack of a standardized model hinders progress in investigating suture-regenerative therapies and poses challenges for conducting comparative analyses across distinct studies. To address this issue, the current protocol describes the detailed modeling process of calvarial suture-bony composite defects in rats.
The model was generated by drilling full-thickness rectangular holes measuring 4.5 mm &#215; 2 mm across the coronal sutures. The rats were euthanized, and the cranium samples were harvested postoperatively at day 0, week 2, week 6, and week 12. &#181;CT results from samples collected immediately post-surgery confirmed the successful establishment of the suture-bony composite defect, involving the removal of the coronal suture and the adjacent bone tissues.
Data from the 6th and 12th postoperative weeks demonstrated a natural healing tendency for the defect to close. Histological staining further validated this trend by showing increased mineralized fibers and new bone at the defect center. These findings indicate progressive suture fusion over time following calvarial defects, underscoring the significance of therapeutic interventions for suture regeneration. We anticipate that this protocol will facilitate the development of suture-regenerative therapies, offering fresh insights into the functional restoration of calvarial defects and reducing adverse outcomes associated with suture loss.

Author Spotlight: Development and Evaluation of a Standardized Rat Model for Calvarial Suture-Bony Composite Defects

We describe the detailed surgical procedures of calvarial suture-bony composite defects in rats, alongside the investigations into the short-term and long-term prognoses of the model. We aim to construct a standardized model for developing suture-regenerative therapies.

Large-scale calvarial defects often coincide with cranial suture disruption, leading to impairments in calvarial defect restoration and skull development ...

Niche-Based Isolation of Rat Jaw Bone Marrow Mesenchymal Stem Cells

Technique for Isolation and Culture of Rat Jaw Bone Marrow Mesenchymal Stem Cells

Bone marrow mesenchymal stem cells (BMMSCs) are a type of stem cell with multi-directional differentiation potential. Compared with BMMSCs derived from appendicular bones, BMMSCs derived from the jaw have greater proliferative and osteogenic differentiation ability, gradually becoming important seed cells for jaw defect repair. However, the mandible has a complex bony structure and less cancellous content than appendicular bones. It is difficult to acquire a large number of high-quality jaw-derived marrow mesenchymal stem cells using traditional methods. This study presents a &#39;niche-based approach on stemness&#39; for isolating and culturing rat jaw bone marrow mesenchymal stem cells (JBMMSCs). Primary rat JBMMSCs were isolated and cultured using the whole bone marrow adherent method combined with the bone slice digestion method. The isolated cells were identified as JBMMSCs through cell morphology observation, detection of cell surface markers, and multi-directional differentiation induction. The cells extracted by this method exhibit a &#39;fibroblast-like&#39; spindle shape. The cells are long, spindle-shaped and fibroblast-like. The flow cytometry analysis shows these cells are positive for CD29, CD44, and CD90 but negative for CD11b/c, CD34, and CD45, which is congruent with BMMSCs characteristics. The cells show strong proliferation capacity and can undergo osteogenic, adipogenic, and chondrogenic differentiation. This study provides an effective and stable method for obtaining enough high-quality JBMMSCs with strong differentiation ability in a short time, which could facilitate further studies of the exploration of biological function, regenerative medicine, and related clinical applications.

Mesenchymal stem cells from the jaw bone marrow have significant functions in diverse differentiation, self-renewal, and immune modulation. They have emerged as a crucial reservoir of precursor cells in gene therapy, tissue engineering, and regenerative medicine. Here, we present a unique method for isolating jaw bone marrow mesenchymal stem cells in rats.

Bone marrow mesenchymal stem cells (BMMSCs) are a type of stem cell with multi-directional differentiation potential. Compared with BMMSCs derived from ...

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

Summary

Explore More Videos

Isolation and Enrichment of Human Adipose-derived Stromal Cells for Enhanced Osteogenesis

Use of Human Perivascular Stem Cells for Bone Regeneration

Repair of a Critical-sized Calvarial Defect Model Using Adipose-derived Stromal Cells Harvested from Lipoaspirate

Visualizing Angiogenesis by Multiphoton Microscopy In Vivo in Genetically Modified 3D-PLGA/nHAp Scaffold for Calvarial Critical Bone Defect Repair

Biological Compatibility Profile on Biomaterials for Bone Regeneration

Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials

Isolation and Culture of Bone Marrow Mesenchymal Stem Cells from the Human Mandible

Isolation and Cultivation of Mandibular Bone Marrow Mesenchymal Stem Cells in Rats

The Establishment of Calvarial Suture-Bony Composite Defects in Rats: A Standardized Model for Suture-Regenerative Therapy Investigation

Technique for Isolation and Culture of Rat Jaw Bone Marrow Mesenchymal Stem Cells