Mechanical Micronization of Lipoaspirates for Regenerative Therapy

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

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 developed^6,7,8,9. 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 models^8,9,10.

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.

Protokół

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

1. Harvest fat.
   1. 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.
   2. Collect 200 mL of lipoaspirates in a sterile bag.
2. Prepare Coleman fat.
   1. Transfer the lipoaspirates into four 50 mL tubes and allow the harvested fat to stand still for 10 min.
   2. 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.
   3. Using 50 mL tubes, centrifugate the fat layer at 1,200 x g at room temperature (RT) for 3 min.
   4. Define the upper layer (approximately 80 mL) as Coleman fat.
   5. 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.
   6. 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.
3. Produce ECM/SVF-gel from low-density fat.
   1. 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.
   2. Keep the shifting speed stable (at 20 mL/s) and repeat for 6x to 8x.
   3. Centrifugate the mixture at 2,000 x g at RT for 3 min.
   4. Collect the oil portion on the top in a 10 mL tube by using a wide-tip pipette at RT for further use.
   5. 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.
4. Produce ECM/SVF-gel from high-density fat.
   1. Add 5 mL of oil (collected from step 1.3.4) to 15 mL of high-density fat.
   2. Intershift the mixed fat between syringes 6x to 8x until a flocculate is observed within the emulsion.
   3. Centrifugate the mixture at 2,000 x g at RT for 3 min.
   4. Discard the oil portion on the top.
   5. 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.
   6. Mix the ECM/SVF-gel from steps 1.3.5 and 1.4.5.

2. Nude Mouse ECM/SVF-gel Graft Model

1. Anesthetize the nude mice (8 weeks old, female) with isoflurane (1% - 3%) inhalation anesthesia in an animal operation room.
2. Transfer the ECM/SVF-gel to a 1 mL syringe.
3. Connect the 1 mL syringe with a blunt infiltration cannula.
4. Insert the cannula subcutaneously into each flank of the mouse.
5. Inject 0.3 mL of the ECM/SVF-gel.

3. Tissue Harvesting on 3, 15, and 90 Days after ECM/SVF-gel Injection

1. Anesthetize the mice with isoflurane (1% - 3%) inhalation anesthesia.
2. Sacrifice the mice by the cervical dislocation method.
3. Make an incision at the midline of the mouse's dorsal skin with surgical scissors.
4. Dissect and harvest the fat grafts on both sides of the mouse. Embed the fat grafts in the 4% paraformaldehyde at RT overnight.
5. 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.
6. Infiltrate the tissue with paraffin wax (58 - 60 °C), two changes, 2 h each.
7. Embed the tissue into paraffin blocks. Cut sections at a thickness of about 4 µm and put them on slides.

4. Hematoxylin and Eosin Staining

1. Deparaffinize the paraffin block slides by soaking them in xylene I, II, and III (10 min each).
2. Rehydrate the tissue sections by passing them through decreasing concentrations (100%, 100%, 95%, 80%, 70%) of ethanol baths for 3 min each.
3. Rinse the tissue sections in distilled water (5 min).
4. Stain the tissue sections in hematoxylin for 5 min.
5. 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.
6. Add two to three drops of Eosin Y dye directly onto the slides by pipette, and let the dye set for 10 min.
7. Wash the slides in tap water for 1 - 5 min.
8. Dehydrate the slides in increasing concentrations (70%, 80%, 95%, 100%, 100%) of ethanol for 3 min each.
9. Clear the slides in xylene I and II for 5 min each.
10. Mount the slides in mounting media.

5. Immunofluorescent Staining

1. Deparaffinize the tissue sections in xylene I, II, and III (5 min each).
2. Rehydrate the tissue sections by passing them through different concentrations (100%, 100%, 95%, 95%, 70%) of alcohol baths for 3 min each.
3. Incubate the sections in a 3% H₂O₂ solution in methanol at RT for 10 min.
4. Rinse the slides 2x with distilled water, 5 min each.
5. 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 °C for 10 min.
6. Cool the slides in RT for 20 min.
7. Rinse the slides 2x with phosphate-buffered saline (PBS), 5 min each.
8. Add 100 µL of 10% fetal bovine serum onto the slides and incubate in a humidified chamber at RT for 1 h.
9. Incubate the sections with primary antibody solution (guinea pig anti-mouse Perilipin, 1:400) at 4 °C overnight.
10. Rinse the slides with PBS 3x, 5 min each.
11. Incubate the sections with secondary antibody solution (goat anti-guinea pig-488 IgG) for 2 h at RT.
12. Rinse the slides with PBS 3x, 5 min each.
13. Wipe off the water around the section with clean tissue paper.
14. Drop 4′,6-diamidino-2-phenylindole (DAPI) and Alexa Fluor 488-conjugated isolectin into the circle to cover the section on the slide.
15. Mount the coverslips and let them dry in the dark.
16. Observe the slides with a fluorescent microscope.

Wyniki

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 (

Dyskusje

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...

Materiały

Name	Company	Catalog Number	Comments
Alexa Fluor 488-conjugated isolectin GS-IB4	Molecular Probes	I21411
guinea pig anti-mouse perilipin	Progen	GP29
DAPI	Thermofisher	D1306
wide tip pipet	Celltreat	229211B
Confocal microscope	Leica	TCS SP2
nude nice	Southern Mdical University	/
light microscope	Olympus	/
50 mL tube	Cornig	430828
sterile bag	Laishi	/
microtome	Leica	CM1900
centrifuge	Heraus