This work was supported by the National Nature Science Foundation of China (81471881, 81601702, 81671931), the Natural Science Foundation of the Guangdong Province of China (2014A030310155), and the Administrator Foundation of Nanfang Hospital (2014B009, 2015Z002, 2016Z010, 2016B001).

This research was approved by the Ethical Review Board in Nanfang Hospital, Guangzhou, China. Adipose tissue was collected from healthy donors who gave written informed consent to take part in the study. All animal experiments were approved by the Nanfang Hospital Institutional Animal Care and Use Committee and performed according to the guidelines of the National Health and Medical Research Council (China).
1. ECM/SVF-gel Preparation
<ol>
	<li>Harvest fat.
	<ol>
		<li>Perform liposuction on a human with a 3-mm multiport cannula, which contains several sharp side holes of 1 mm in diameter, at -0.75 atm of suction pressure.</li>
		<li>Collect 200 mL of lipoaspirates in a sterile bag.</li>
	</ol>
	</li>
	<li>Prepare Coleman fat.
	<ol>
		<li>Transfer the lipoaspirates into four 50 mL tubes and allow the harvested fat to stand still for 10 min.</li>
		<li>Collect the fat on the top layer into two 50 mL tubes by using a wide-tip pipette to transfer, and discard the liquid portion at the bottom layer.</li>
		<li>Using 50 mL tubes, centrifugate the fat layer at 1,200 x g at room temperature (RT) for 3 min.</li>
		<li>Define the upper layer (approximately 80 mL) as Coleman fat.</li>
		<li>Transfer the upper 2/3 of the Coleman fat to a 20 mL tube by using a wide-tip pipette, and define this portion as low-density fat.</li>
		<li>Transfer the lower 1/3 of the Coleman fat to another 20 mL tube by using a wide-tip pipette, and define this portion as high-density fat.</li>
	</ol>
	</li>
	<li>Produce ECM/SVF-gel from low-density fat.
	<ol>
		<li>Using two 20 mL syringes connected by a female-to-female Luer-Lock connector (with an internal diameter of 2.4 mm) to intershift 20 mL of low-density fat.</li>
		<li>Keep the shifting speed stable (at 20 mL/s) and repeat for 6x to 8x.</li>
		<li>Centrifugate the mixture at 2,000 x g at RT for 3 min.</li>
		<li>Collect the oil portion on the top in a 10 mL tube by using a wide-tip pipette at RT for further use.</li>
		<li>Collect the sticky substance in the middle layer, which is ECM/SVF-gel (Figure 1A), by using a wide-tip pipette, and discard the fluid at the bottom layer.</li>
	</ol>
	</li>
	<li>Produce ECM/SVF-gel from high-density fat.
	<ol>
		<li>Add 5 mL of oil (collected from step 1.3.4) to 15 mL of high-density fat.</li>
		<li>Intershift the mixed fat between syringes 6x to 8x until a flocculate is observed within the emulsion.</li>
		<li>Centrifugate the mixture at 2,000 x g at RT for 3 min.</li>
		<li>Discard the oil portion on the top.</li>
		<li>Collect the sticky substance in the middle layer (ECM/SVF-gel) by using a wide-tip pipette and discard the fluid at the bottom layer.</li>
		<li>Mix the ECM/SVF-gel from steps 1.3.5 and 1.4.5.</li>
	</ol>
	</li>
</ol>
2. Nude Mouse ECM/SVF-gel Graft Model
<ol>
	<li>Anesthetize the nude mice (8 weeks old, female) with isoflurane (1% - 3%) inhalation anesthesia in an animal operation room.</li>
	<li>Transfer the ECM/SVF-gel to a 1 mL syringe.</li>
	<li>Connect the 1 mL syringe with a blunt infiltration cannula.</li>
	<li>Insert the cannula subcutaneously into each flank of the mouse.</li>
	<li>Inject 0.3 mL of the ECM/SVF-gel.</li>
</ol>
3. Tissue Harvesting on 3, 15, and 90 Days after ECM/SVF-gel Injection
<ol>
	<li>Anesthetize the mice with isoflurane (1% - 3%) inhalation anesthesia.</li>
	<li>Sacrifice the mice by the cervical dislocation method.</li>
	<li>Make an incision at the midline of the mouse&#39;s dorsal skin with surgical scissors.</li>
	<li>Dissect and harvest the fat grafts on both sides of the mouse. Embed the fat grafts in the 4% paraformaldehyde at RT overnight.</li>
	<li>Dehydrate the tissue in increasing concentrations of ethanol: 70% ethanol, two changes, 1 h each; 80% ethanol, one change, 1 h; 95% ethanol, one change, 1 h; 100% ethanol, three changes, 1.5 h each; xylene, three changes, 1.5 h each.</li>
	<li>Infiltrate the tissue with paraffin wax (58 - 60 &#176;C), two changes, 2 h each.</li>
	<li>Embed the tissue into paraffin blocks. Cut sections at a thickness of about 4 &#181;m and put them on slides.</li>
</ol>
4. Hematoxylin and Eosin Staining
<ol>
	<li>Deparaffinize the paraffin block slides by soaking them in xylene I, II, and III (10 min each).</li>
	<li>Rehydrate the tissue sections by passing them through decreasing concentrations (100%, 100%, 95%, 80%, 70%) of ethanol baths for 3 min each.</li>
	<li>Rinse the tissue sections in distilled water (5 min).</li>
	<li>Stain the tissue sections in hematoxylin for 5 min.</li>
	<li>Rinse the tissue sections in running tap water for 20 min. Decolorize in 1% acid alcohol (1% HCl in 70% alcohol) for 5 s. Rinse in running tap water until the sections are blue again.</li>
	<li>Add two to three drops of Eosin Y dye directly onto the slides by pipette, and let the dye set for 10 min.</li>
	<li>Wash the slides in tap water for 1 - 5 min.</li>
	<li>Dehydrate the slides in increasing concentrations (70%, 80%, 95%, 100%, 100%) of ethanol for 3 min each.</li>
	<li>Clear the slides in xylene I and II for 5 min each.</li>
	<li>Mount the slides in mounting media.</li>
</ol>
5. Immunofluorescent Staining
<ol>
	<li>Deparaffinize the tissue sections in xylene I, II, and III (5 min each).</li>
	<li>Rehydrate the tissue sections by passing them through different concentrations (100%, 100%, 95%, 95%, 70%) of alcohol baths for 3 min each.</li>
	<li>Incubate the sections in a 3% H2O2 solution in methanol at RT for 10 min.</li>
	<li>Rinse the slides 2x with distilled water, 5 min each.</li>
	<li>Drop the slides in a slide basket. Add 300 mL of 10 mM citrate buffer (pH 6.0) and incubate the slides at 95 - 100 &#176;C for 10 min.</li>
	<li>Cool the slides in RT for 20 min.</li>
	<li>Rinse the slides 2x with phosphate-buffered saline (PBS), 5 min each.</li>
	<li>Add 100 &#181;L of 10% fetal bovine serum onto the slides and incubate in a humidified chamber at RT for 1 h.</li>
	<li>Incubate the sections with primary antibody solution (guinea pig anti-mouse Perilipin, 1:400) at 4 &#176;C overnight.</li>
	<li>Rinse the slides with PBS 3x, 5 min each.</li>
	<li>Incubate the sections with secondary antibody solution (goat anti-guinea pig-488 IgG) for 2 h at RT.</li>
	<li>Rinse the slides with PBS 3x, 5 min each.</li>
	<li>Wipe off the water around the section with clean tissue paper.</li>
	<li>Drop 4&#8242;,6-diamidino-2-phenylindole (DAPI) and Alexa Fluor 488-conjugated isolectin into the circle to cover the section on the slide.</li>
	<li>Mount the coverslips and let them dry in the dark.</li>
	<li>Observe the slides with a fluorescent microscope.</li>
</ol>

<ol>
	<li>Bateman, M. E., 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=Using+Fat+to+Fight+Disease:+A+Systematic+Review+of+Non-Homologous+Adipose-Derived+Stromal/Stem+Cell+Therapies.">Using Fat to Fight Disease: A Systematic Review of Non-Homologous Adipose-Derived Stromal/Stem Cell Therapies.</a> Stem Cells. 36 (9), 1311-1328 (2018).</li><li>Baer, P. C., Geiger, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose-derived+mesenchymal+stromal/stem+cells:+tissue+localization,+characterization,+and+heterogeneity.">Adipose-derived mesenchymal stromal/stem cells: tissue localization, characterization, and heterogeneity.</a> Stem Cells International. , 812693 (2012).</li><li>Gimble, J. M., Katz, A. J., Bunnell, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose-derived+stem+cells+for+regenerative+medicine.">Adipose-derived stem cells for regenerative medicine.</a> Circulation Research. 100 (9), 1249-1260 (2017).</li><li>Halme, D. G., Kessler, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FDA+regulation+of+stem-cell-based+therapies.">FDA regulation of stem-cell-based therapies.</a> New England Journal of Medicine. 355 (16), 1730-1735 (2006).</li><li>Cheng, N. C., Wang, S., Young, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+spheroid+formation+of+human+adipose-derived+stem+cells+on+chitosan+films+on+stemness+and+differentiation+capabilities.">The influence of spheroid formation of human adipose-derived stem cells on chitosan films on stemness and differentiation capabilities.</a> Biomaterials. 33 (6), 1748-1758 (2012).</li><li>van Dongen, J. 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=Comparison+of+intraoperative+procedures+for+isolation+of+clinical+grade+stromal+vascular+fraction+for+regenerative+purposes:+a+systematic+review.">Comparison of intraoperative procedures for isolation of clinical grade stromal vascular fraction for regenerative purposes: a systematic review.</a> Journal of Tissue Engineering and Regenerative Medicine. 12 (1), 261-274 (2018).</li><li>van Dongen, J. 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=The+fractionation+of+adipose+tissue+procedure+to+obtain+stromal+vascular+fractions+for+regenerative+purposes.">The fractionation of adipose tissue procedure to obtain stromal vascular fractions for regenerative purposes.</a> Wound Repair and Regeneration. 24 (6), 994-1003 (2016).</li><li>Mashiko, T., 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=Mechanical+Micronization+of+Lipoaspirates:+Squeeze+and+Emulsification+Techniques.">Mechanical Micronization of Lipoaspirates: Squeeze and Emulsification Techniques.</a> Plastic and Reconstructive Surgery. 139 (1), 79-90 (2017).</li><li>Feng, 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=Micronized+cellular+adipose+matrix+as+a+therapeutic+injectable+for+diabetic+ulcer.">Micronized cellular adipose matrix as a therapeutic injectable for diabetic ulcer.</a> Regenerative Medicine. 10 (6), 699-708 (2015).</li><li>Zhang, P., 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=Ischemic+flap+survival+improvement+by+composition-selective+fat+grafting+with+novel+adipose+tissue+derived+product+-+stromal+vascular+fraction+gel.">Ischemic flap survival improvement by composition-selective fat grafting with novel adipose tissue derived product - stromal vascular fraction gel.</a> Biochemistry and Biophysics Research Communication. 495 (3), 2249-2256 (2018).</li><li>Tonnard, P., 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=Nanofat+grafting:+basic+research+and+clinical+applications.">Nanofat grafting: basic research and clinical applications.</a> Plastic and Reconstructive Surgery. 132 (4), 1017-1026 (2013).</li><li>Yao, Y., 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=Adipose+Extracellular+Matrix/Stromal+Vascular+Fraction+Gel:+A+Novel+Adipose+Tissue-Derived+Injectable+for+Stem+Cell+Therapy.">Adipose Extracellular Matrix/Stromal Vascular Fraction Gel: A Novel Adipose Tissue-Derived Injectable for Stem Cell Therapy.</a> Plastic and Reconstructive Surgery. 139 (4), 867-879 (2017).</li><li>Yao, Y., 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=Adipose+Stromal+Vascular+Fraction+Gel+Grafting:+A+New+Method+for+Tissue+Volumization+and+Rejuvenation.">Adipose Stromal Vascular Fraction Gel Grafting: A New Method for Tissue Volumization and Rejuvenation.</a> Dermatologic Surgery. 44 (10), 1278-1286 (2018).</li><li>Zhang, Y., 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=Improved+Long-Term+Volume+Retention+of+Stromal+Vascular+Fraction+Gel+Grafting+with+Enhanced+Angiogenesis+and+Adipogenesis.">Improved Long-Term Volume Retention of Stromal Vascular Fraction Gel Grafting with Enhanced Angiogenesis and Adipogenesis.</a> Plastic and Reconstructive Surgery. 141 (5), 676-686 (2018).</li><li>Sun, B., 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=Applications+of+stem+cell-derived+exosomes+in+tissue+engineering+and+neurological+diseases.">Applications of stem cell-derived exosomes in tissue engineering and neurological diseases.</a> Reviews in the Neurosciences. 29 (5), 531-546 (2018).</li><li>Allen, R. 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=Grading+lipoaspirate:+is+there+an+optimal+density+for+fat+grafting.">Grading lipoaspirate: is there an optimal density for fat grafting.</a> Plastic and Reconstructive Surgery. 131 (1), 38-45 (2013).</li><li>Qiu, L., 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=Identification+of+the+Centrifuged+Lipoaspirate+Fractions+Suitable+for+Postgrafting+Survival.">Identification of the Centrifuged Lipoaspirate Fractions Suitable for Postgrafting Survival.</a> Plastic and Reconstructive Surgery. 137 (1), 67-76 (2016).</li></ol>

The authors have nothing to disclose.

Stem cell-based regenerative therapy has shown a great potential benefit in different diseases. ASCs are outstanding therapeutic candidates because they are easy to obtain and have the capacity for tissue repair and the regeneration of novel tissues15. However, there are limitations to expanding its clinical application, since it requires complicated procedures to isolate cells and collagenase for processing6. Thus, it is essential to develop a simple technique to obtain st...

Stem cell therapies provide a paradigm shift for tissue repair and regeneration so that they may offer an alternative therapeutic regimen for various diseases1. Stem cells (e.g., induced pluripotent stem cells and embryonic stem cells) have great therapeutic potential but are limited due to cell regulation and ethical considerations. Adipose-derived mesenchymal stromal/stem cells (ASCs) are easy to obtain from lipoaspirates and not subject to the same restrictions; thus, it has become an ideal cell type for practical regenerative medicine2. In addition, they are nonimmunogenic and have abundant resources from autologous fat3.
Currently, ASCs are obtained mainly by collagenase-mediated digestion of the adipose tissue. The stromal vascular fraction (SVF) of adipose tissue contains ASCs, endothelial progenitor cell, pericytes, and immune cells. Although obtaining a high density of SVF/ASCs enzymatically was shown to have beneficial effects, the legislation in several countries strictly regulates the clinical application of cell-based products using collagenase4. Digesting the adipose tissue with collagenase for 30 min to 1 h to obtain SVF cells increases the risk of both exogenous material in the preparation and biological contamination. The adherent culture and the purification of ASCs, which takes days to weeks, require specific laboratory equipment. Moreover, in most studies, SVF cells and ASCs are used in suspension. Without the protection of extracellular matrix (ECM) or another carrier, free cells are vulnerable, cause a poor cell retention after injection, and compromise the therapeutic result5. All of these reasons limit the further application of stem cell therapy.
To obtain ASCs from adipose tissue without collagenase-mediated digestion, several mechanical processing procedures, including centrifugation, mechanical chopping, shredding, pureeing, and mincing, have been developed6,7,8,9. These methods are thought to condense tissue and ASCs by mechanically disrupting mature adipocytes and their oil-containing vesicles. Moreover, these preparations, containing high concentrations of ASCs, showed considerable therapeutic potential as regenerative medicine in animal models8,9,10.
In 2013, Tonnard et al. introduced the nanofat grafting technique, which involves producing emulsified lipoaspirates by intersyringe processing11. The shearing force created by intersyringe shifting can selectively break mature adipocytes. Based on their findings, we developed a purely mechanical processing method that removes most of the lipid and fluid in the lipoaspirates, leaving only SVF cells and fractionated ECM, which is ECM/SVF-gel12. Herein, we describe the details of the mechanical process of human-derived adipose tissue to produce the ECM/SVF-gel.

<table><tbody><tr><td>Alexa Fluor 488-conjugated isolectin GS-IB4</td><td>Molecular Probes</td><td>I21411</td><td></td></tr><tr><td>guinea pig anti-mouse perilipin</td><td>Progen</td><td>GP29</td><td></td></tr><tr><td>DAPI</td><td>Thermofisher</td><td>D1306</td><td></td></tr><tr><td>wide tip pipet</td><td>Celltreat</td><td>229211B</td><td></td></tr><tr><td>Confocal microscope&amp;nbsp;</td><td>Leica&amp;nbsp;</td><td>TCS SP2</td><td></td></tr><tr><td>nude nice&amp;nbsp;</td><td>Southern Mdical University</td><td>/</td><td></td></tr><tr><td>light microscope&amp;nbsp;</td><td>Olympus</td><td>/</td><td></td></tr><tr><td>50 mL tube</td><td>Cornig</td><td>430828</td><td></td></tr><tr><td>sterile bag</td><td>Laishi</td><td>/</td><td></td></tr><tr><td>microtome</td><td>Leica&amp;nbsp;</td><td>CM1900</td><td></td></tr><tr><td>centrifuge</td><td>Heraus</td><td></td><td></td></tr></tbody></table>

mechanical micronization of lipoaspirates for regenerative therapy

Stromal vascular fraction (SVF) has become a regenerative tool for various diseases; however, legislation strictly regulates the clinical application of cell products using collagenase. Here, we present a protocol to generate an injectable mixture of SVF cells and native extracellular matrix from adipose tissue by a purely mechanical process. Lipoaspirates are put into a centrifuge and spun at 1,200 x g for 3 min. The middle layer is collected and separated into two layers (high-density fat at the bottom and low-density fat on the top). The upper layer is directly emulsified by intersyringe shifting, at a rate of 20 mL/s for 6x to 8x. The emulsified fat is centrifuged at 2,000 x g for 3 min, and the sticky substance under the oil layer is collected and defined as the extracellular matrix (ECM)/SVF-gel. The oil on the top layer is collected. Approximately 5 mL of oil is added to 15 mL of high-density fat and emulsified by intersyringe shifting, at a rate of 20 mL/s for 6x to 8x. The emulsified fat is centrifuged at 2,000 x g for 3 min, and the sticky substance is also ECM/SVF-gel. After the transplantation of the ECM/SVF-gel into nude mice, the graft is harvested and assessed by histologic examination. The result shows that this product has the potential to regenerate into normal adipose tissue. This procedure is a simple, effective mechanical dissociation procedure to condense the SVF cells embedded in their natural supportive ECM for regenerative purposes.

After processing the Coleman fat to ECM/SVF-gel, the volume of discarded oil takes up 80% of the final volume, and only 20% of adipose tissue preserved under the oil layer is regarded as ECM/SVF-gel (Figure 1A). ECM/SVF-gel has a smooth liquid-like texture that enables it to go through a 27 G fine needle; however, Coleman fat is comprised of an integral adipose structure with large fibers and can only go through an 18 G cannula (<strong class...

Watch this Scientific Journal Video about Mechanical Micronization of Lipoaspirates for Regenerative Therapy at JoVE.com

Mechanical Micronization of Lipoaspirates for Regenerative Therapy

Here, we present a protocol to obtain stromal vascular fraction from adipose tissue through a series of mechanical processes, which include emulsification and multiple centrifugations.

Stromal vascular fraction (SVF) has become a regenerative tool for various diseases; however, legislation strictly regulates the clinical application of ...

mechanical-micronization-of-lipoaspirates-for-regenerative-therapy

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7.5K Views.  Southern Medical University. Here, we present a protocol to obtain stromal vascular fraction from adipose tissue through a series of mechanical processes, which include emulsification and multiple centrifugations.

Video: Mechanical Micronization of Lipoaspirates for Regenerative Therapy

Constant Pressure-controlled Extrusion Method for the Preparation of Nano-sized Lipid Vesicles

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform to study protein-protein and protein-lipid interactions3, monitor drug delivery4,5, and encapsulation4. Phospholipids naturally create curved lipid bilayers, distinguishing itself from a micelle.6 Liposomes are traditionally classified by size and number of bilayers, i.e. large unilamellar vesicles (LUVs), small unilamellar vesicles (SUVs) and multilamellar vesicles (MLVs)7. In particular, the preparation of homogeneous liposomes of various sizes is important for studying membrane curvature that plays a vital role in cell signaling, endo- and exocytosis, membrane fusion, and protein trafficking8. Several groups analyze how proteins are used to modulate processes that involve membrane curvature and thus prepare liposomes of diameters &lt;100 - 400 nm to study their behavior on cell functions3. Others focus on liposome-drug encapsulation, studying liposomes as vehicles to carry and deliver a drug of interest9. Drug encapsulation can be achieved as reported during liposome formation9. Our extrusion step should not affect the encapsulated drug for two reasons, i.e. (1) drug encapsulation should be achieved prior to this step and (2) liposomes should retain their natural biophysical stability, securely carrying the drug in the aqueous core. These research goals further suggest the need for an optimized method to design stable sub-micron lipid vesicles.
Nonetheless, the current liposome preparation technologies (sonication10, freeze-and-thaw10, sedimentation) do not allow preparation of liposomes with highly curved surface (i.e. diameter &lt;100 nm) with high consistency and efficiency10,5, which limits the biophysical studies of an emerging field of membrane curvature sensing. Herein, we present a robust preparation method for a variety of biologically relevant liposomes.
Manual extrusion using gas-tight syringes and polycarbonate membranes10,5 is a common practice but heterogeneity is often observed when using pore sizes &lt;100 nm due to due to variability of manual pressure applied. We employed a constant pressure-controlled extrusion apparatus to prepare synthetic liposomes whose diameters range between 30 and 400 nm. Dynamic light scattering (DLS)10, electron microscopy11 and nanoparticle tracking analysis (NTA)12 were used to quantify the liposome sizes as described in our protocol, with commercial polystyrene (PS) beads used as a calibration standard. A near linear correlation was observed between the employed pore sizes and the experimentally determined liposomes, indicating high fidelity of our pressure-controlled liposome preparation method. Further, we have shown that this lipid vesicle preparation method is generally applicable, independent of various liposome sizes. Lastly, we have also demonstrated in a time course study that these prepared liposomes were stable for up to 16 hours. A representative nano-sized liposome preparation protocol is demonstrated below.

This protocol describes an extrusion method for preparing lipid vesicles of sub-micron sizes with a high degree of homogeneity. This method uses a pressure-controlled system with controlled nitrogen flow rates for liposome preparation. The lipid preparation1,2, liposome extrusion, and size characterization will be presented herein.

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform ...

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. ...

Intra-lymph Node Injection of Biodegradable Polymer Particles

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these foreign molecules to T and B lymphocytes. Lymph nodes have thus become critical targets for new vaccines and immunotherapies. A recent strategy for targeting these tissues is direct lymph node injection of soluble vaccine components, and clinical trials involving this technique have been promising. Several biomaterial strategies have also been investigated to improve lymph node targeting, for example, tuning particle size for optimal drainage of biomaterial vaccine particles. In this paper we present a new method that combines direct lymph node injection with biodegradable polymer particles that can be laden with antigen, adjuvant, or other vaccine components. In this method polymeric microparticles or nanoparticles are synthesized by a modified double emulsion protocol incorporating lipid stabilizers. Particle properties (e.g.&#160;size, cargo loading) are confirmed by laser diffraction and fluorescent microscopy, respectively. Mouse lymph nodes are then identified by peripheral injection of a nontoxic tracer dye that allows visualization of the target injection site and subsequent deposition of polymer particles in lymph nodes. This technique allows direct control over the doses and combinations of biomaterials and vaccine components delivered to lymph nodes and could be harnessed in the development of new biomaterial-based vaccines.

Lymph nodes are the immunological tissues that orchestrate immune response and are a critical target for vaccines. Biomaterials have been employed to better target lymph nodes and to control delivery of antigens or adjuvants. This paper describes a technique combining these ideas to inject biocompatible polymer particles into lymph nodes.

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these ...

Stromal Vascular Fraction-enriched Fat Grafting for the Treatment of Symptomatic End-neuromata

The purpose of this study was to methodically illustrate and highlight the crucial steps of stromal vascular fraction (SVF)-enriched fat grafting as a novel treatment of symptomatic end-neuromata of peripheral sensory nerves, and in this study, specifically of the superficial branch of the radial nerve (SBRN). Despite a multitude of existing treatments, persistent postoperative pain and common pain relapse are still very common, independent of the procedure assessed. The neuroma is microsurgically excised accordingly to standardized protocol. Instead of the relocation of the regenerating nerve stump in neighboring anatomical structures, such as muscle or bone, a fat graft is applied perifocally and acts as a mechanical barrier. In order to reduce the fat resorption rate and boost the regenerative potential of the graft, the highly concentrated SVF is integrated in the grafting. The SVF is isolated from subcutaneous fat by enzymatic and mechanic separation of the lipoaspirate by a specific commercial isolation system. The SVF-enriched fat graft provides both a mechanical barrier and various biological effects at the cellular level, including improving angiogenesis, inflammation, and fibrosis. Both mechanical and biologic effects help to reduce the disorganized axonal outgrowth of the nerve stump during nerve regeneration and hence prevent the recurrence of painful end-neuromata.

The purpose of the study is to illustrate the technical procedure of stromal vascular fraction (SVF)-enriched fat grafting as a novel treatment of symptomatic end-neuromata. This technique provides advantages of both the mechanical barrier and the biological action at the cellular level by the processed and concentrated SVF.

The purpose of this study was to methodically illustrate and highlight the crucial steps of stromal vascular fraction (SVF)-enriched fat grafting as a ...

Formulating and Characterizing Lipid Nanoparticles for Gene Delivery using a Microfluidic Mixing Platform

Lipid-based drug carriers have been used for clinically and commercially available delivery systems due to their small size, biocompatibility, and high encapsulation efficiency. Use of lipid nanoparticles (LNPs) to encapsulate nucleic acids is advantageous to protect the RNA or DNA from degradation, while also promoting cellular uptake. LNPs often contain multiple lipid components including an ionizable lipid, helper lipid, cholesterol, and polyethylene glycol (PEG) conjugated lipid. LNPs can readily encapsulate nucleic acids due to the ionizable lipid presence, which at low pH is cationic and allows for complexation with negatively charged RNA or DNA. Here LNPs are formed by encapsulating messenger RNA (mRNA) or plasmid DNA (pDNA) using rapid mixing of the lipid components in an organic phase and the nucleic acid component in an aqueous phase. This mixing is performed using a precise microfluidic mixing platform, allowing for nanoparticle self-assembly while maintaining laminar flow. The hydrodynamic size and polydispersity are measured using dynamic light scattering (DLS). The effective surface charge on the LNP is determined by measuring the zeta potential. The encapsulation efficiency is characterized using a fluorescent dye to quantify entrapped nucleic acid. Representative results demonstrate the reproducibility of this method and the influence that different formulation and process parameters have on the developed LNPs.

Lipid nanoparticles are developed using a microfluidic mixing platform approach for mRNA and DNA encapsulation.

Lipid-based drug carriers have been used for clinically and commercially available delivery systems due to their small size, biocompatibility, and high ...

Clinical Protocol of Producing Adipose Tissue-Derived Stromal Vascular Fraction for Potential Cartilage Regeneration

Osteoarthritis (OA) is one of the most common debilitating disorders.&#160;Recently, numerous attempts have been made to improve the functions of the knees by using different forms of mesenchymal stem cells (MSCs). In Korea, bone marrow concentrates and cord blood-derived stem cells have been approved by the Korean Food and Drug Administration (KFDA) for cartilage regeneration. In addition, an adipose tissue-derived stromal vascular fraction (SVF) has been allowed by the KFDA for joint injections in human patients. Autologous adipose tissue-derived SVF contains extracellular matrix (ECM) in addition to mesenchymal stem cells. ECM excretes various cytokines that, along with hyaluronic acid (HA) and platelet-rich plasma (PRP) activated by calcium chloride, may help MSCs to regenerate cartilage and improve knee functions. In this article, we presented a protocol to improve knee functions by regenerating cartilage-like tissue in human patients with OA. The result of the protocol was first reported in 2011 followed by a few additional publications. The protocol involves liposuction to obtain autologous lipoaspirates that are mixed with collagenase. This lipoaspirates-collagenase mixture is then cut and homogenized to remove large fibrous tissue that may clog up the needle during the injection. Afterwards, the mixture is incubated to obtain adipose tissue-derived SVF. The resulting adipose tissue-derived SVF, containing both adipose tissue-derived MSCs and remnants of ECM, is injected into knees of patients, combined with HA and calcium chloride activated PRP. Included are three cases of patients who were treated with our protocol resulting in improvement of knee pain, swelling, and range of motion along with MRI evidence of hyaline cartilage-like tissue.

Here, we present a protocol to produce an adipose tissue-derived stromal vascular fraction and its application to improve knee functions by regenerating cartilage-like tissue in human patients with osteoarthritis.

Osteoarthritis (OA) is one of the most common debilitating disorders.&#160;Recently, numerous attempts have been made to improve the functions of the ...

Manual Isolation of Adipose-derived Stem Cells from Human Lipoaspirates

In 2001, researchers at the University of California, Los Angeles, described the isolation of a new population of adult stem cells from liposuctioned adipose tissue that they initially termed Processed Lipoaspirate Cells or PLA cells. Since then, these stem cells have been renamed as Adipose-derived Stem Cells or ASCs and have gone on to become one of the most popular adult stem cells populations in the fields of stem cell research and regenerative medicine. Thousands of articles now describe the use of ASCs in a variety of regenerative animal models, including bone regeneration, peripheral nerve repair and cardiovascular engineering. Recent articles have begun to describe the myriad of uses for ASCs in the clinic. The protocol shown in this article outlines the basic procedure for manually and enzymatically isolating ASCs from large amounts of lipoaspirates obtained from cosmetic procedures. This protocol can easily be scaled up or down to accommodate the volume of lipoaspirate and can be adapted to isolate ASCs from fat tissue obtained through abdominoplasties and other similar procedures.

In 2001, researchers at UCLA described the isolation of a population of adult stem cells, termed Adipose-derived Stem Cells or ASCs, from adipose tissue. This article outlines the isolation of ASCs from lipoaspirates using a manual, enzymatic digestion protocol using collagenase.

In 2001, researchers at the University of  California, Los Angeles, described the isolation of a new population of adult  stem cells from liposuctioned ...

Biology

Isolation of Adipose Derived Regenerative Cells for the Treatment of Erectile Dysfunction Following Radical Prostatectomy

Stem cells are used in many research areas within regenerative medicine in part because these treatments can be curative rather than symptomatic. Stem cells can be obtained from different tissues and several methods for isolation have been described. The presented method for the isolation of adipose-derived regenerative cells (ADRCs) can be used within many therapeutic areas because the method is a general procedure and, therefore, not limited to erectile dysfunction (ED) therapy. ED is a common and serious side effect to radical prostatectomy (RP) since ED often is not well treated with conventional therapy. Using ADRC&#8217;s as treatment for ED has attracted great interest due to the initial positive results after a single injection of cells into the corpora cavernosum. The method used for the isolation of ADRC&#8217;s is a simple, automated process, that is reproducible and ensures a uniform product. Furthermore, the sterility of the isolated product is ensured because the entire process takes place in a closed system. It is important to minimize the risk of contamination and infection since the stem cells are used for injection in humans. The whole procedure can be done within 2.5-3.5 hours and does not require a classified laboratory which eliminates the need for shipping tissue to an off-site. However, the procedure has some limitations since the minimum amount of drained lipoaspirate for the isolation device to function is 100 g.

Precise disclosure of methods and protocols is crucial for large scale uptake of stem cell therapies. Here, we present a protocol to isolate adipose-derived regenerative cells, used for a single intracavernous injection as treatment of erectile dysfunction (ED) following radical prostatectomy (RP).

Stem cells are used in many research areas within regenerative medicine in part because these treatments can be curative rather than symptomatic. Stem ...

Technique for Obtaining Mesenchymal Stem Cell from Adipose Tissue and Stromal Vascular Fraction Characterization in Long-Term Cryopreservation

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible source for regenerative clinical applications. Different protocols have been used to obtain adipose-derived stem cells. This article describes different steps of an improved time-saving protocol to obtain a more significant amount of ADSC, showing how to cryopreserve and thaw ADSC to obtain viable cells for culture expansion. One hundred milliliters of lipoaspirate were collected, using a 26 cm three-hole and 3 mm caliber syringe liposuction, from the abdominal area of nine patients who subsequently underwent elective abdominoplasty. The stem cells isolation was carried out with a series of washes with Dulbecco&#39;s Phosphate Buffered Saline (DPBS) solution supplemented with calcium and the use of collagenase. Stromal Vascular Fraction (SVF) cells were cryopreserved, and their viability was checked by immunophenotyping. The SVF cellular yield was 15.7 x 105 cells/mL, ranging between 6.1-26.2 cells/mL. Adherent SVF cells reached confluence after an average of 7.5 (&#177;4.5) days, with an average cellular yield of 12.3 (&#177; 5.7) x 105 cells/mL. The viability of thawed SVF after 8 months, 1 year, and 2 years ranged between 23.06%-72.34% with an average of 47.7% (&#177;24.64) with the lowest viability correlating with cases of two-year freezing. The use of DPBS solution supplemented with calcium and bag resting times for fat precipitation with a shorter time of collagenase digestion resulted in an increased stem cell final cellular yield. The detailed procedure for obtaining high yields of viable stem cells was more efficient regarding time and cellular yield than the techniques from previous studies. Even after a long period of cryopreservation, viable ADSC cells were found in the SVF.

The present protocol describes an improved methodology for ADSC isolation resulting in a tremendous cellular yield with time gain compared to the literature. This study also provides a straightforward method for obtaining a relatively large number of viable cells after long-term cryopreservation.

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible ...

Mechanical Stromal Vascular Fraction in Fibrin Hydrogel for Regenerative Medicine

The Combination of Mechanically Isolated Stromal Vascular Fraction and Fibrin Hydrogel: A Processing Protocol

The regenerative potential of adipose-derived stromal cells (ASCs) has gained significant attention in regenerative and translational research. In the past, the extraction of these cells from adipose tissue required a multistep enzyme-based process, resulting in a heterogenous cell mix consisting of ACSs and other cells, which are jointly termed the stromal vascular fraction (SVF). More recently introduced mechanical SVF (mSVF) isolation protocols are less time-consuming and bypass regulatory concerns. We recently proposed a protocol that generates mSVF rich in stromal cells based on a combination of emulsification and centrifugation. One current issue in mSVF application for wound therapy application is the lack of a scaffold providing protection from mechanical manipulation and desiccation. Fibrin hydrogels have been shown to be a useful adjunct in cell transfer for wound healing purposes in the past. In the work herein, we delineate the preparation steps of an mSVF-fibrin hydrogel construct as a novel approach for translational research and clinical application.

Author Spotlight: Exploring the Potential of Fat-Derived Stromal Vascular Fraction for Wound Healing

Mechanically isolated stromal vascular fraction (SVF) in combination with a fibrin hydrogel offers an easy and efficient carrier for viable adipose-derived stromal cells for various indications, including tissue engineering and or wound healing purposes. Here, we present the preparation of a mechanical SVF (mSVF)-fibrin hydrogel construct for translational research and clinical application.

The regenerative potential of adipose-derived stromal cells (ASCs) has gained significant attention in regenerative and translational research. In the ...