The authors wish to thank Claire Cryer and Fiona Rossi at the University of Edinburgh for their expert assistance with flow cytometry. We also wish to thank the personnel of the Murrayfield hospital who contributed by providing tissue specimens.
This work was supported by grants from the British Heart Foundation and Lipogems, which supplied adipose tissue processing kits. Human adult tissue samples were procured with full ethics permission of the South East Scotland Research Ethics Committee (reference: 16/SS/0103).

Ethical approval for the use of human tissues in this research was obtained from the South East Scotland Research Ethics Committee (reference: 16/SS/0103).
1. Subcutaneous Abdominal Adipose Tissue Collection
NOTE: All instruments used in the manual lipo-aspiration procedure are provided by the manufacturer of the micro-fragmentation device.
<ol>
	<li>Maintain sterility for all fluids, containers, instruments, and the operational area throughout the experiment.</li>
	<li>Place the abdominoplasty sample (procured by the surgical team in a sterile bag without any solution) on top of a surgical cloth with the skin facing upwards. No washing is required. For optimal use, keep the sample at 4 &#176;C and process the adipose sample within 16 h of harvesting.</li>
	<li>Inject 50 to 100 mL of 37 &#176;C 0.9% NaCl solution, depending on the size of the tissue sample(use 50 mL for maximum a 15 cm2 adipose tissue surface and increase the volume accordingly), using a disposable tissue infiltration cannula, into the subcutaneous adipose tissue. Handle the specimen by the cutaneous part.</li>
	<li>Connect a 10 mL Luer lock syringe to a disposable liposuction cannula.</li>
	<li>Carefully introduce the cannula inside the adipose tissue from the edges. Once inside, create the vacuum inside the syringe by pulling the plunger. Keep the plunger in position by hand to secure the vacuum suction.</li>
	<li>Make radial movements inside the adipose tissue sample until the syringe is full of lipoaspirate. Carefully remove the cannula from the tissue and disconnect the syringe. Connect a new empty syringe.</li>
	<li>Repeat the procedure until 60 mL of lipoaspirate have been collected. 
	NOTE: The maximum amount of lipoaspirate that can be processed changes according to the type of device used. Please refer to the manufacturer instructions (Table of Materials).</li>
</ol>
2. Micro-fragmentation of the lipoaspirate
NOTE: This protocol is meant for the research use only. Micro-fragmentation is performed with the help of a commercially available device (Table of Materials).
<ol>
	<li>Remove the device from the package and verify that the main unit is connected to the waste bag. Place the waste collection bag on the ground and make sure that the valves are secured to the process unit caps.</li>
	<li>Connect the terminal spike of the input line to the saline bag by piercing the bag connection port. Keep the saline bag higher than the processing unit. 
	NOTE: For the type of device used in this protocol, a 2,000 mL bag of sterile saline is recommended.</li>
	<li>Verify that all 5 clamps connected to the tubes of the circuits are open. Place the processing unit in a vertical position with the gray cap upwards.</li>
	<li>Allow the saline solution to fill the processing unit. Check that the flow reaches the waste bag. Remove all air bubbles from the processing unit by shaking it. 
	NOTE:&#160;All the following passages must be performed in the absence of air in the processing unit.</li>
	<li>Place the processing unit in a vertical position with the blue cap upwards and close the clamp next to the input line.</li>
	<li>Start injecting the lipoaspirate by connecting the syringe to the self-occluding valve of the blue input cap. Keep the processing unit vertical during this procedure and slowly pull the syringe plunger until all the lipoaspirate has been transferred to the unit. Repeat the process until all the lipoaspirate is inside the processing unit. 
	NOTE:&#160;For the processing unit used in the video, add maximum 30 mL of lipoaspirate at the time. The procedure can be performed maximum twice, until 60 mL of lipoaspirate have been processed.</li>
	<li>Open the input clamp to restore the saline flow.</li>
	<li>Vigorously shake the processing unit for 2 min at least, regularly checking the saline solution flow into the waste bag.</li>
	<li>When the saline solution in the processing unit turns transparent, after approximatively 2 min of shaking, place the unit with the gray cap upwards, and close the clamp located on the drain near the gray cap. The processed tissue will float at the top.</li>
	<li>Fill a 10 mL Luer lock syringe with saline solution and connect it to the loading valve of the blue cap. Close the clamp located near the blue cap. Connect an empty 10 mL Luer lock syringe, with the plunger completely inserted, to the valve of the gray cap. 
	NOTE: Use Luer lock syringes only.</li>
	<li>Firmly inject the saline into the processing unit from the blue cap. Ensure that the syringe connected to the gray valve is consequently being filled with the processed tissue. Once having injected all the saline, carefully remove both syringes.</li>
	<li>Repeat steps 2.10 and 2.11 until all the processed tissue is collected. 
	NOTE: The average yield of MAT is 30 mL from 60 mL of manual lipoaspirate. Some tissue will remain inside the processing unit and some clumps might be present attached to the blue edge inside the unit.</li>
	<li>At the end of the processing, remove all syringes, close all clamps, detach the saline bag and dispose the device according to local protocols. 
	NOTE: The processing device cannot be re-used.</li>
</ol>
3. Fluorescence Immunohistochemistry
<ol>
	<li>Transfer 300-500 &#181;L of MAT into a 1.5 mL tube. Add 1 mL of 4% buffered paraformaldehyde (PFA), mix gently by hand (no vortexing) and leave at 4 &#176;C overnight (from 16 to 24 h).</li>
	<li>Collect the specimen and transfer to a clean 1.5 mL tube containing 1 mL of PBS. Repeat the step twice to remove PFA excess.</li>
	<li>Transfer the specimen in 15% (w/v) sucrose in PBS solution and incubate at 4 &#176;C overnight (from 16 to 24 h)</li>
	<li>Transfer the specimen in a well of a 24 well plate and add an embedding solution made of 15% sucrose and 7% gelatin (w/v) in PBS. Fill the well halfway. Incubate for 4 h at 37 &#176;C.</li>
	<li>Transfer the samples to 4 &#176;C. After 2-4 h for the gelatin to solidify, fill up the well with the embedding solution and leave at 4 &#176;C overnight.</li>
	<li>Remove the samples from the wells. Remove excess embedding compound and freeze on dry ice. Store the samples at -80 &#176;C.</li>
	<li>Cut 8-10 &#181;m thick sections using a cryostat. For optimal sectioning, use the cryostat at -30 &#176;C. Slides can be stored at -80 &#176;C.</li>
	<li>Prior to proceeding with the immunofluorescence, fix the sections in 4% PFA for 7 min. Remove the PFA and wash twice with lukewarm PBS (37 &#176;C) in order to remove gelatin from the sections.</li>
	<li>Block non-specific antibody binding by incubating the slides with 10% goat serum in PBS (v/v) for 1 h at room temperature.</li>
	<li>Dilute the primary antibodies in antibody diluent or 0.2% (w/v) bovine serum albumin (BSA) in PBS (w/v): mouse anti-human-NG2 (1:100, stock 0.5 mg/mL), rabbit anti-human-PDGFR&#946; (1:100, stock 0.15 mg/mL).</li>
	<li>Remove the blocking solution and add the diluted primary antibodies. Incubate at 4 &#176;C overnight. 
	NOTE: Perform all the incubation steps from now on in the dark.</li>
	<li>Remove the primary antibodies and wash 3 times with PBS for 10 min each time.</li>
	<li>Dilute the secondary antibodies in antibody diluent or 0.2% (w/v) BSA in PBS: goat anti-mouse- 555 (1:300), goat anti-rabbit- 647 (1:300). Incubate 1 h at room temperature.</li>
	<li>Remove the secondary antibodies and wash three times with PBS for 10 min each time.</li>
	<li>If a biotin conjugated lectin is used, use the avidin/biotin blocking kit. Incubate for 10 min with each reagent provided with two washes in between with PBS.</li>
	<li>Repeat the blocking step by incubating the slides for 1 h at room temperature with 10% goat serum in PBS.</li>
	<li>Add the biotinylated lectin, biotinylated Ulex europaeus lectin (1:200, stock 2 mg/mL), diluted either in 0.2% BSA (w/v) in PBS or antibody diluent. Incubate 1 h and 30 min at room temperature.</li>
	<li>Remove excess biotinylated lectin and wash three times with PBS for 10 min each time.</li>
	<li>Add the secondary streptavidin conjugated antibody diluted in either 0.2% (w/v) BSA in PBS or antibody diluent. Incubate for 1 h.</li>
	<li>Remove the secondary antibody and wash three times with PBS for 10 min each time.</li>
	<li>Counterstain nuclei with 4&#8242;,6-diamidino-2-phenylindole (DAPI, 1:500, stock 5 mg/mL), diluted in 0.2% BSA (w/v) in PBS, for 10 min at room temperature.</li>
	<li>Remove the DAPI solution and wash twice with PBS, 10 min each time.</li>
	<li>Mount the slides with an aqueous mounting agent. Let it dry completely, either overnight on the bench in the dark or 1 h at 37 &#176;C.</li>
	<li>Keep the slides at 4 &#176;C in the dark and acquire images on an epifluorescence microscope.</li>
</ol>
4. Digestion of the micro-fragmented adipose tissue and cell isolation
<ol>
	<li>Transfer the micro-fragmented adipose tissue into a sterile container.</li>
	<li>Freshly make up the digestion medium by dissolving type-II collagenase in DMEM at the concentration of 1 mg/mL.</li>
	<li>Add the same volume of digestion medium to the micro-fragmented adipose tissue (i.e., for 30 mL of sample add 30 mL of digestion medium).</li>
	<li>Place the sealed container in a 37 &#176;C shaking water bath set to 120 rpm for 45 min. Please note that the container should allow a proper shaking of the sample. If a large container is not available, use two 50 mL tubes (30 mL of sample/digestion medium mixture in each) placed horizontally.</li>
	<li>After the incubation, block the digestion by adding an equal volume of blocking solution (2% (v/v) FCS/PBS) and filter sequentially through 100 &#181;m and 70 &#181;m cell strainers.</li>
	<li>Centrifuge the filtered suspension for 5 min at 200 x g. Discard the supernatant.</li>
	<li>Resuspend the pellet in approximately 5 mL of erythrocyte lysis buffer (155 mM NH4Cl, 170 mM Tris, pH 7.65). The volume of buffer may vary according to the erythrocyte contamination observed in the cellular pellet. Incubate at room temperature for 15 min.</li>
	<li>Add an equal volume of blocking solution and filter through a 40 &#181;m cell strainer.</li>
	<li>Centrifuge the cell suspension for 5 min at 200 x g and discard the supernatant.</li>
	<li>Resuspend the pellet in blocking solution. Mix well by pipetting. Count viable cells with Trypan blue exclusion on a hemocytometer.
	<ol>
		<li>Mix an equal volume of the cell suspension with Trypan blue stain. Aspirate 10 &#181;L of the solution and place it on a hemocytometer.</li>
		<li>Count the number of live cells (bright and round) present in at least 5 squares and get the mean cell number. In order to include the stain dilution factor, multiply the mean cell number by 2. To get the total number of live cells per 1 mL of sample multiply the obtained number by 104. 
		NOTE: The obtained single cell suspension, namely the stromal vascular fraction (SVF), can be either seeded at 20,000 cells/cm2 in perivascular cell growth medium (DMEM supplemented with 20% heat inactivated FCS, 1% penicillin/streptomycin and 1% L-glutamine) or used for flow cytometry analysis and sorting. The average yield of nucleated cells in the SVF is approximately 3 x 104 cells per mL of MAT. Sorting experiments require at least 8 x 105 cells to obtain enough perivascular cells for culturing.</li>
	</ol>
	</li>
</ol>
5. Cell labelling and sorting
<ol>
	<li>Centrifuge the cell suspension at room temperature for 5 min at 200 x g, and re-suspend the pellet in blocking medium at a concentration of 1 x 106 cells per 100 &#181;L.</li>
	<li>Aliquot at least 50,000 cells for the unstained control and the fluorescence minus one (FMO) controls into polystyrene round bottom flow cytometry tubes. Use the rest of the cells for multicolor staining by placing in another tube. 
	NOTE: Once the settings of the experiment have been established (i.e., laser intensity, gating strategy), the number of cells used for the unstained control and the FMOs can be reduced, if, the antibodies used are the same.</li>
	<li>Add CD31-V450 (1:400), CD34-PE (1:100), CD45-V450 (1:400) and CD146-BV711 (1:100) antibodies to the single cell suspension for multi-color staining. For the FMO controls, add: for the V450 FMO equivalent volumes of CD34-PE and CD146-BV711 plus V450 isotype control; for the PE FMO equivalent volumes of CD31-V450, CD45-V450 and CD146-BV711 plus PE isotype control; for the BV711 FMO equivalent volumes of CD31-V450, CD45-V450 and CD34-PE plus BV711 isotype control.</li>
	<li>Gently pipette, or vortex slowly the solution in the tube to mix and incubate the tubes at 4 &#176;C for 20 min in the dark.</li>
	<li>Prepare compensation control beads. Add 15 &#181;L of positive beads and 15 &#181;L of negative beads to 70 &#181;L of blocking medium in 3 polystyrene round-bottom flow-cytometry tubes. Add CD34-PE, CD31-V450 or CD45-V450, and CD146-BV711 antibodies, one per tube.</li>
	<li>Gently pipette, or vortex slowly, to mix the antibodies with the beads and incubate all tubes at 4 &#176;C for 20 min in the dark. 
	NOTE: The procedure described can be used either for flow cytometry analysis or cell sorting.</li>
	<li>Prepare collection tubes by pre-wetting the inner surface of the tubes with endothelial growth medium (EGM), leaving approximately 50 &#181;L of medium on the bottom of each tube.</li>
	<li>After antibody incubation, remove the excess of antibody by washing the cells and the beads with 2 mL of blocking medium.</li>
	<li>Remove the solution by centrifuging the tubes at 200 x g for 5 min and carefully aspirating the supernatant. Replicate the washing step.</li>
	<li>Resuspend the cells in blocking medium at a final concentration of 1 x 106 cells per 250 &#181;L. Resuspend the beads in 100 &#181;L of blocking medium.</li>
	<li>Transfer the cells into new polystyrene round-bottom flow cytometry tubes with cell strainer cap. This will allow the disruption of cell clumps.</li>
	<li>Transport all cell and bead suspensions to the cell sorter on ice in the dark.</li>
	<li>Run unstained control cells to establish the background fluorescence and set the voltages.</li>
	<li>Run the compensation control tubes to set the fluorescence compensation.</li>
	<li>Use forward scatter area (FSC-A) vs side scatter area (SSC-A) to identify cells, and then use forward scatter area (FSC-A) vs forward scatter height (FSC-H) to select single cells.</li>
	<li>Add DAPI, 1 &#181;L of 5 &#181;g/mL solution for each mL of sample, to the unstained control to set the gating for dead cells (Figure 2). No washing is required after DAPI staining.</li>
	<li>Run isotype controls to establish background fluorescence thresholds related to non-specific binding and set the gating.</li>
	<li>Run the multi-color stained sample. Exclude hematopoietic and endothelial cells by gating on CD31 and CD45 negative cells. Collect the pericyte (CD146+ CD34-) and adventitial cell (CD146- CD34+) populations into collection tubes (Figure 2). 
	NOTE: Optimal viable cell yields are approximately 2% of the total live cell dissociation for pericytes and 1.5% for adventitial cells.</li>
</ol>
6. Cell culture
<ol>
	<li>Seed freshly sorted pericytes and adventitial cells at a density of 30,000 to 40,000 cells/cm2 on gelatin coated culture plates</li>
	<li>To coat the culture plates, cover the plate area with sterile 0.2% (w/v) gelatin in PBS solution (approximately 100 &#181;L of solution per cm2). Incubate at room temperature for 10 min and remove the solution. Keep freshly sorted cells on ice during culture plate coating.</li>
	<li>Centrifuge freshly collected cells at 200 x g for 5 min and gently re-suspend the cell pellet in an appropriate amount of endothelial growth medium (EGM). If the number of cells collected is very low, avoid the centrifugation and plate directly the collected cells.</li>
	<li>Seed the cells on gelatin-coated plates. Incubate at 37 &#176;C in 5% CO2.</li>
	<li>Once cells have settled and adhered to the plate (after at least 72 h), exchange EGM with perivascular cell growth medium (DMEM supplemented with 20% heat inactivated FCS, 1% penicillin/streptomycin and 1% L-glutamine) for both pericytes and adventitial cells.</li>
	<li>Once pericytes and adventitial cells have reached 80%-90% confluence, dissociate cells using 0.05% trypsin-EDTA. Collect with 5% FCS/PBS, centrifuge at 200 x g for 5 min, re-suspend in perivascular cell growth medium, and then passage the cells from 1:3 to 1:5 ratio (or at approximately 7,000 cells/cm2) onto uncoated polystyrene culture plates or flasks.</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multilineage+cells+from+human+adipose+tissue:+implications+for+cell-based+therapies.">Multilineage cells from human adipose tissue: implications for cell-based therapies.</a> Tissue Engineering Journal. 7 (2), 211-228 (2001).</li><li>Sch&#228;ffler, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Concise+review:+Adipose+tissue-derived+stromal+cells-basic+and+clinical+implications+for+novel+cell-based+therapies.">Concise review: Adipose tissue-derived stromal cells-basic and clinical implications for novel cell-based therapies.</a> Stem Cells. 25 (4), 818-827 (2007).</li><li>Bianchi, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+non+enzymatic+method+and+device+to+obtain+a+fat+tissue+derivative+highly+enriched+in+pericyte-like+elements+by+mild+mechanical+forces+from+human+lipoaspirates.">A new non enzymatic method and device to obtain a fat tissue derivative highly enriched in pericyte-like elements by mild mechanical forces from human lipoaspirates.</a> Cell Transplantation. 2, 2063-2077 (2013).</li><li>Raffaini, M., Tremolada, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Micro+fractured+and+purified+adipose+tissue+graft+(Lipogems)+can+improve+the+orthognathic+surgery+outcomes+both+aesthetically+and+in+postoperative+healing.">Micro fractured and purified adipose tissue graft (Lipogems) can improve the orthognathic surgery outcomes both aesthetically and in postoperative healing.</a> CellR4. 2 (4), (2014).</li><li>Cestaro, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intersphincteric+anal+lipofilling+with+micro-fragmented+fat+tissue+for+the+treatment+of+faecal+incontinence:+preliminary+results+of+three+patients.">Intersphincteric anal lipofilling with micro-fragmented fat tissue for the treatment of faecal incontinence: preliminary results of three patients.</a> Wideochir Inne Tech Maloinwazyjne. 10 (2), 337-341 (2015).</li><li>Fantasia, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfractured+and+purified+adipose+tissue+(Lipogems+system)+injections+for+treatment+of+atrophic+vaginitis.">Microfractured and purified adipose tissue (Lipogems system) injections for treatment of atrophic vaginitis.</a> Journal of Urology Research. 3 (7), 1073-1075 (2016).</li><li>Saibene, A. 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CT is a founder of Lipogems.

This paper describes the physical fractionation, using a closed system device, of human adipose tissue into small clusters displaying normal adipose tissue microanatomy.
Manually aspirated human subcutaneous adipose tissue and saline solution are loaded into a transparent plastic cylinder containing large pinball-style metallic spheres that, upon vigorous manual shaking of the device, rupture the fat into sub-millimeter fragments. Attached filters and outlet allow eliminating debris, blood and...

Adipose tissue, long used as a filler in reconstructive and cosmetic surgery, has recently become more popular in regenerative medicine once recognized as a source for mesenchymal stem cells (MSCs)1. Lipoaspirates dissociated enzymatically into single-cell suspensions yield an adipocyte-free stromal vascular fraction (SVF) that is used unaltered in the patient or, more commonly, is cultured for several weeks into MSCs2.
However, enzyme dissociation ruptures the tissue microenvironments, secluding neighboring regulatory cells from presumptive regenerative cells that become considerably modified by in vitro culture. To avoid such experimental artifacts and consequent functional alterations, attempts have been made to process adipose tissue for therapeutic use while maintaining its native configuration as intact as possible3,4. Notably, mechanical tissue disruption has started to replace enzymatic dissociation. To this end, the full immersion closed system micro-fragments lipoaspirates into sub-millimeter, blood- and oil-free tissue clusters (e.g., Lipogems) via a sequence of sieve filtration and steel marble induced disruption3. Autologous transplantation of micro-fragmented adipose tissue, using this closed system technology, has been successful in multiple indications, spanning cosmetics, orthopedics, proctology and gynaecology4,5,6,7,8,9,10,11,12,13.
Comparison between human micro-fragmented adipose tissue (MAT) obtained with the closed system device and isogenic SVF revealed that with respect to vascular/stromal cell distribution and secretory activity in culture, MAT contains more pericytes, which are presumptive MSCs14, and secretes higher amounts of growth factors and cytokines15.
The present article illustrates the enzyme-free micro-fragmentation of human subcutaneous adipose tissue using a closed system device, and the further processing of such micronized adipose tissue for in vitro culture, immunohistochemistry and FACS analysis, in order to identify the cell types present and the soluble factors secreted (Figure 1). The described method safely generates adipose derived sub-millimeter organoids containing viable adipose tissue cell populations in an intact niche, suitable for further applications and studies.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>4% Buffered paraformaldehyde (PFA)</td>
			<td>VWR chemicals</td>
			<td>9317.901</td>
			<td></td>
		</tr>
		<tr>
			<td>0.9% NaCl Solution</td>
			<td>Baxter</td>
			<td>3KB7127</td>
			<td></td>
		</tr>
		<tr>
			<td>AlexaFluor 555 goat anti-mouse IgG&#160;</td>
			<td>Life Technologies</td>
			<td>A21422</td>
			<td></td>
		</tr>
		<tr>
			<td>AlexaFluor 647 goat anti-Rabbit IgG</td>
			<td>Life Technologies</td>
			<td>&#160;A21245</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium chloride</td>
			<td>fisher chemicals</td>
			<td>1158868</td>
			<td></td>
		</tr>
		<tr>
			<td>Antigent Diluent</td>
			<td>Life Technologies</td>
			<td>3218</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Mouse Ig, &#954;/Negative Control (BSA) Compensation Plus</td>
			<td>BD Biosciences</td>
			<td>560497</td>
			<td></td>
		</tr>
		<tr>
			<td>Avidin/Biotin Blocking Kit</td>
			<td>Life Technologies</td>
			<td>4303</td>
			<td></td>
		</tr>
		<tr>
			<td>BD LSR Fortessa 5-laser flow cytometer&#160;</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>Laser 405nm (violet)/375nm (UV) &#8211; filter V450/50 for DAPI and V450 antibodies; Laser 561nm (Yellow-green) &#8211; filter YG582/15 for PE antibodies; Laser 405nm (violet)/375nm (UV) &#8211; filter V710/50 for BV711 antibodies&#160;</td>
		</tr>
		<tr>
			<td>Biotinylated Ulex europaeus lectin</td>
			<td>Vector Laboratories</td>
			<td>Vector-B1065</td>
			<td></td>
		</tr>
		<tr>
			<td>BV711 Mouse IgG1, k Isotype Control</td>
			<td>BD Biosciences</td>
			<td>563044</td>
			<td></td>
		</tr>
		<tr>
			<td>CD146-BV711</td>
			<td>BD Biosciences</td>
			<td>563186</td>
			<td></td>
		</tr>
		<tr>
			<td>CD31-V450</td>
			<td>BD Biosciences</td>
			<td>561653</td>
			<td></td>
		</tr>
		<tr>
			<td>CD34-PE</td>
			<td>BD Biosciences</td>
			<td>555822</td>
			<td></td>
		</tr>
		<tr>
			<td>CD45-V450</td>
			<td>BD Biosciences</td>
			<td>560367</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Life Technologies</td>
			<td>D1306</td>
			<td>stock concentration: 5mg/mL</td>
		</tr>
		<tr>
			<td>Disposable liposuction cannula (LGI 13Gx185 mm &#8211; AR 13/18) &#160;</td>
			<td>Lipogems&#160;</td>
			<td></td>
			<td>provided in the Lipogems surgical kit</td>
		</tr>
		<tr>
			<td>Diva software 306 (v.6.0)</td>
			<td>BD Biosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM, high glucose, GlutaMAX without sodium pyruvate</td>
			<td>Life Technologies</td>
			<td>61965026</td>
			<td></td>
		</tr>
		<tr>
			<td>EGMTM-2 Endothelial Cell Growth Medium-2 BulletKitTM</td>
			<td>Lonza&#160;</td>
			<td>CC-3156</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Calf Serum (FCS)</td>
			<td>Sigma-Aldrich</td>
			<td>F2442</td>
			<td></td>
		</tr>
		<tr>
			<td>FlowJo (v.10.0)</td>
			<td>FlowJo</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount G</td>
			<td>SouthernBiotech</td>
			<td>0100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin</td>
			<td>Acros Organics</td>
			<td>410870025</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipogems Surgical Kit</td>
			<td>Lipogems&#160;</td>
			<td>LG SK 60</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti human- NG2</td>
			<td>BD Biosciences</td>
			<td>554275</td>
			<td>stock concentration: 0.5 mg/mL</td>
		</tr>
		<tr>
			<td>PE Mouse IgG1, &#954; Isotype Control</td>
			<td>BD Biosciences</td>
			<td>555749</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Sigma-Aldrich</td>
			<td>P4333</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS)</td>
			<td>Sigma-Aldrich</td>
			<td>D8537</td>
			<td></td>
		</tr>
		<tr>
			<td>Polystirene round bottom 5 mL tube with cell strainer snap cap&#160;</td>
			<td>BD Biosciences</td>
			<td>352235, 25/Pack</td>
			<td></td>
		</tr>
		<tr>
			<td>Polystyrene round bottom 5 mL tubes</td>
			<td>BD Biosciences</td>
			<td>352003</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti human - PDGFRb</td>
			<td>Abcam</td>
			<td>32570</td>
			<td>stock concentration: 0.15 mg/mL</td>
		</tr>
		<tr>
			<td>Streptavidin conjugated-488</td>
			<td>Life Technologies</td>
			<td>&#160;S32354</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>84100-5kg</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue infiltration cannula (17GX185 mm-VG 17/18)&#160;</td>
			<td>Lipogems&#160;</td>
			<td></td>
			<td>provided in the Lipogems surgical kit</td>
		</tr>
		<tr>
			<td>Tris base</td>
			<td>fisher chemicals</td>
			<td>BP152-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Type- II Collagenase</td>
			<td>Gibco</td>
			<td>17101-015</td>
			<td></td>
		</tr>
		<tr>
			<td>V450 Mouse IgG1, &#954; Isotype Control</td>
			<td>BD Biosciences</td>
			<td>560373</td>
			<td></td>
		</tr>
		<tr>
			<td>Widefield Zeiss observer</td>
			<td>Zeiss</td>
			<td></td>
			<td>Objective used: Plan-Apo 20x/0.8</td>
		</tr>
		<tr>
			<td>Zeiss Colibri7 LED light source ( LEDs: 385, 475, 555, 590, 630 nm)</td>
			<td>Zeiss</td>
			<td></td>
			<td>DAPI: UV, excitation 385nm; 488: Blue, excitation 475nm;&#160; 555: Green, excitation 555nm;&#160; 647:Red, excitation 630nm&#160;</td>
		</tr>
	</tbody>
</table>

human adipose tissue micro-fragmentation for cell phenotyping and secretome characterization

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. Accordingly, techniques for harvesting and processing human adipose tissue have evolved in order to quickly and efficiently obtain large amounts of tissue. Among these, the closed system technology represents an innovative and easy-to-use system to harvest, process, and re-inject refined fat tissue in a short time and in the same intervention (intra-operatively). Adipose tissue is collected by liposuction, washed, emulsified, rinsed and minced mechanically into cell clusters of 0.3 to 0.8 mm. Autologous transplantation of mechanically fragmented adipose tissue has shown remarkable efficacy in different therapeutic indications such as aesthetic medicine and surgery, orthopedic and general surgery. Characterization of micro-fragmented adipose tissue revealed the presence of intact small vessels within the adipocyte clusters; hence, the perivascular niche is left unperturbed. These clusters are enriched in perivascular cells (i.e., mesenchymal stem cell (MSC) ancestors) and in vitro analysis showed an increased release of growth factors and cytokines involved in tissue repair and regeneration, compared to enzymatically derived MSCs. This suggests that the superior therapeutic potential of microfragmented adipose tissue is explained by a higher frequency of presumptive MSCs and enhanced secretory activity. Whether these added pericytes directly contribute to higher growth factor and cytokine production is not known. This clinically approved procedure allows the transplantation of presumptive MSCs without the need for expansion and/or enzymatic treatment, thus bypassing the requirements of GMP guidelines, and reducing the costs for cell-based therapies.

Mechanical dissociation of manual lipoaspirates resulted in the production of micro-fragmented adipose tissue (MAT), consisting of an aggregate of adipocytes enveloping a microvascular network (Figure 3). Immunofluorescence analysis of gelatin-embedded and cryofixed MAT highlights this structure, showing the vascular network marked by the endothelial cell marker Ulex europaeus agglutinin 1 (UEA-1) receptor mainly consisting of small, capillary-like vessels (<...

Watch this Scientific Journal Video about Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization at JoVE.com

Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization

Here, we present human adipose tissue enzyme-free micro-fragmentation using a closed system device. This new method allows the obtainment of sub-millimeter clusters of adipose tissue suitable for in vivo transplantation, in vitro culture, and further cell isolation and characterization.

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. ...

human-adipose-tissue-micro-fragmentation-for-cell-phenotyping

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JoVE Journal

Biology

9.6K Views.  University of Edinburgh. Here, we present human adipose tissue enzyme-free micro-fragmentation using a closed system device. This new method allows the obtainment of sub-millimeter clusters of adipose tissue suitable for in vivo transplantation, in vitro culture, and further cell isolation and characterization.

Video: Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Transduction of Human Cells with Polymer-complexed Ecotropic Lentivirus for Enhanced Biosafety

Stem and tumor cell biology studies often require viral transduction of human cells with known or suspected oncogenes, raising major safety issues for laboratory personnel. Pantropic lentiviruses, such as the commonly used VSV-G pseudotype, are a valuable tool for studying gene function because they can transduce many cell types, including non-dividing cells. However, researchers may wish to avoid production and centrifugation of pantropic viruses encoding oncogenes due to higher biosafety level handling requirements and safety issues. Several potent oncogenes, including c-Myc and SV40 large T antigen, are known to enhance production of induced pluripotent stem cells (iPSC). All other known iPSC-inducing genetic changes (OCT4, SOX2, KLF4, NANOG, LIN28, and p53 loss of function) also have links to cancer, making them of relatively high safety concern as well. 
While these cancer-related viruses are useful in studying cellular reprogramming and pluripotency, they must be used safely. To address these biosafety issues, we demonstrate a method for transduction of human cells with ecotropic lentivirus, with additional emphasis on reduced cost and convenient handling. We have produced ecotropic lentivirus with sufficiently high titer to transduce greater than 90% of receptor-expressing human cells exposed to the virus, validating the efficacy of this approach. 
Lentivirus is often concentrated by ultracentrifugation; however, this process takes several hours and can produce aerosols infectious to human biomedical researchers. As an alternative, viral particles can be more safely sedimented onto cells by complexation with chondroitin sulfate and polybrene (CS/PB). This technique increases the functional viral titer up to 3-fold in cells stably expressing murine retrovirus receptor, with negligible added time and cost. Transduction of human dermal fibroblasts (HDFs) is maximally enhanced using CS/PB concentrations approximately 4-fold lower than the optimal value previously reported for cancer cell lines, suggesting that polymer concentration should be titrated for the target cell type of interest. We therefore describe the use of methylthiazolyldiphenyl-tetrazolium bromide (MTT) to assay for polymer toxicity in a new cell type. We observe equivalent viability of HDFs after viral transduction using either polymer complexation or the standard dose of polybrene (PB, 6 &mu;g/ml), indicating minimal acute toxicity.
In this protocol, we describe the use of ecotropic lentivirus for overexpression of oncogenes in human cells, reducing biosafety risks and increasing the transduction rate. We also demonstrate the use of polymer complexation to enhance transduction while avoiding aerosol-forming centrifugation of viral particles.

Lentiviruses are a valuable research tool for exploring gene function; however, researchers may wish to avoid production of pantropic lentivirus encoding known or suspected oncogenes. As an alternative, we present a safer protocol for use of ecotropic lentivirus on human cells modified to express the ecotropic receptor mSlc7a1.

Stem and tumor cell biology studies often require viral transduction of human cells with known or suspected oncogenes, raising major safety issues for ...

Tissue Triage and Freezing for Models of Skeletal Muscle Disease

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal results from functional, cellular, molecular, and pathological evaluations. Due to the subtlety of some pathological abnormalities seen in congenital muscle disorders and the potential for fixation to interfere with the recognition of these features, pathological evaluation of frozen muscle is preferable to fixed muscle when evaluating skeletal muscle for congenital muscle disease. Additionally, the potential to produce severe freezing artifacts in muscle requires specific precautions when freezing skeletal muscle for histological examination that are not commonly used when freezing other tissues. This manuscript describes a protocol for rapid freezing of skeletal muscle using isopentane (2-methylbutane) cooled with liquid nitrogen to preserve optimal skeletal muscle morphology. This procedure is also effective for freezing tissue intended for genetic or protein expression studies. Furthermore, we have integrated our freezing protocol into a broader procedure that also describes preferred methods for the short term triage of tissue for (1) single fiber functional studies and (2) myoblast cell culture, with a focus on the minimum effort necessary to collect tissue and transport it to specialized research or reference labs to complete these studies. Overall, this manuscript provides an outline of how fresh tissue can be effectively distributed for a variety of phenotypic studies and thereby provides standard operating procedures (SOPs) for pathological studies related to congenital muscle disease.

The analysis of skeletal muscle tissues to determine structural, functional, and biochemical properties is greatly facilitated by appropriate preparation. This protocol describes appropriate methods to prepare skeletal muscle tissue for a broad range of phenotyping studies.

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal ...

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial respirometric assays from isolated mitochondrial preparations allow for the assessment of mitochondrial function, as well as determination of the mechanism(s) of action of drugs and proteins that modulate metabolism. Current isolation procedures often require large quantities of tissue to yield high quality mitochondria necessary for respirometric assays. The methods presented herein describe how high quality purified mitochondria (~ 450 &#181;g) can be isolated from minimal quantities (~75-100 mg) of mouse skeletal muscle for use in high throughput respiratory measurements. We determined that our isolation method yields 92.5&#177; 2.0% intact mitochondria by measuring citrate synthase activity spectrophotometrically. In addition, Western blot analysis in isolated mitochondria resulted in the faint expression of the cytosolic protein, GAPDH, and the robust expression of the mitochondrial protein, COXIV. The absence of a prominent GAPDH band in the isolated mitochondria is indicative of little contamination from non-mitochondrial sources during the isolation procedure. Most importantly, the measurement of O2 consumption rate with micro-plate based technology and determining the respiratory control ratio (RCR) for coupled respirometric assays shows highly coupled (RCR; &#62;6 for all assays) and functional mitochondria. In conclusion, the addition of a separate mincing step and significantly reducing motor driven homogenization speed of a previously reported method has allowed the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle that results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Visualization of 3D White Adipose Tissue Structure Using Whole-mount Staining

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various techniques have been developed to study the morphology and biology of adipose tissue. However, conventional visualization methods are limited to studying the tissue in 2D sections, failing to capture the 3D architecture of the whole organ. Here we present whole-mount staining, an immunohistochemistry method that preserves intact adipose tissue morphology with minimal processing steps. Hence, the structures of adipocytes and other cellular components are maintained without distortion, achieving the most representative 3D visualization of the tissue. In addition, whole-mount staining can be combined with lineage tracing methods to determine cell fate decisions. However, this technique has some limitations to providing accurate information regarding deeper parts of adipose tissue. To overcome this limitation, whole-mount staining can be further combined with tissue clearing techniques to remove the opaqueness of tissue and allow for complete visualization of entire adipose tissue anatomy using light-sheet fluorescent microscopy. Therefore, a higher resolution and more accurate representation of adipose tissue structures can be captured with the combination of these techniques.

The focus of the present study is to demonstrate the whole-mount immunostaining and visualization technique as an ideal method for 3D imaging of adipose tissue architecture and cellular component.

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various ...

Y-27632 Enriches the Yield of Human Melanocytes from Adult Skin Tissues

The isolation and culture of primary melanocytes from skin tissues is very important for biological research and has been widely used for clinical applications. Isolating primary melanocytes from skin tissues by the conventional method usually takes about 3 to 4 weeks to passage sufficiently. More importantly, the tissues used are usually newborn foreskins and it is still a challenge to efficiently isolate primary melanocytes from adult tissues. We recently developed a new isolation method for melanocytes that adds Y-27632, a Rho kinase inhibitor, to the initial culture medium for 48 h. Compared with the conventional protocol, this new method dramatically increases the yield of melanocytes and shortens the time required to isolate melanocytes from foreskin tissues. We now describe this new method in more detail using adult epidermis to efficiently culture primary melanocytes. Importantly, we show that melanocytes obtained from adult tissues prepared by this new method can function normally. This new protocol will significantly benefit studies of pigmentation defects and melanomas using primary melanocytes prepared from easily accessed adult skin tissues.

This paper reports that the addition of Y-27632 to TIVA medium can significantly increase the yield of melanocytes from adult skin tissues.

The isolation and culture of primary melanocytes from skin tissues is very important for biological research and has been widely used for clinical ...

Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity is characterized by an increase in adipose tissue (AT) mass due to adipocyte hyperplasia and/or hypertrophia, leading to profound remodeling of its three-dimensional structure. Indeed, the maximal capacity of AT to expand during obesity is pivotal to the development of obesity-associated pathologies. This AT expansion is an important homeostatic mechanism to enable adaptation to an excess of energy intake and to avoid deleterious lipid spillover to other metabolic organs, such as muscle and liver. Therefore, understanding the structural remodeling that leads to the failure of AT expansion is a fundamental question with high clinical applicability. In this article, we describe a simple and fast clearing method that is routinely used in our laboratory to explore the morphology of mouse and human white adipose tissue by fluorescent imaging. This optimized AT clearing method is easily performed in any standard laboratory equipped with a chemical hood, a temperature-controlled orbital shaker and a fluorescent microscope. Moreover, the chemical compounds used are readily available. Importantly, this method allows one to resolve the 3D AT structure by staining various markers to specifically visualize the adipocytes, the neuronal and vascular networks, and the innate and adaptive immune cells distribution.

Here, we describe a simple, inexpensive and fast clearing method to resolve the 3D structure of both mouse and human white adipose tissue using a combination of markers to visualize vasculature, nuclei, immune cells, neurons, and lipid-droplet coat proteins by fluorescent imaging.

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity ...

Generation of Human Kidney Tubuloids from Tissue and Urine

Adult stem cell (ASC)-derived human kidney epithelial organoids, or tubuloids, can be established from healthy and diseased kidney epithelium with high efficiency. Normal kidney tubuloids recapitulate many aspects of their tissue of origin. They represent distinct nephron segments - most notably of the proximal tubule, loop of Henle, distal tubules, and collecting duct - and can be used to study normal kidney physiology. Furthermore, tubuloid technology facilitates disease modeling, e.g., for infectious diseases as well as for cancer. Obtaining kidney epithelial cells for tubuloid generation is, however, dependent on leftover surgical material (such as partial) nephrectomies) or needle biopsies. The ability to grow tubuloids from urine would provide an alternative, less invasive source of healthy kidney epithelial cells. It has been previously shown that tubuloid cultures can be successfully generated from only a few milliliters of freshly collected urine. This article describes the protocols to generate and propagate ASC-derived human kidney tubuloid cultures from tissue and urine samples.

Human kidney tubuloid cultures represent a valuable in vitro model to study kidney physiology and disease. Tubuloids can be established from kidney tissue (healthy and diseased) as well as urine, the latter representing an easily obtainable and less invasive source of research material.

Adult stem cell (ASC)-derived human kidney epithelial organoids, or tubuloids, can be established from healthy and diseased kidney epithelium with high ...

Isolation, Characterization, and Therapeutic Application of Extracellular Vesicles from Cultured Human Mesenchymal Stem Cells

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological regulators of organismal homeostasis and important indicators of pathologies; in the meantime, their immense potential to establish accessible and controllable disease therapeutics is emerging. Mesenchymal stem cells (MSCs) can release large amounts of EVs in culture, which have shown promise to jumpstart effective tissue regeneration and facilitate extensive therapeutic applications with good scalability and reproducibility. There is a growing demand for simple and effective protocols for collecting and applying MSC-EVs. Here, a detailed protocol is provided based on differential centrifugation to isolate and characterize representative EVs from cultured human MSCs, exosomes, and microvesicles for further applications. The adaptability of this method is shown for a series of downstream approaches, such as labeling, local transplantation, and systemic injection. The implementation of this procedure will address the need for simple and reliable MSC-EVs collection and application in translational research.

The present protocol describes the differential centrifugation for isolating and characterizing representative EVs (exosomes and microvesicles) from cultured human MSCs. Further applications of these EVs are also explained in this article.

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological ...