Dr. Joel Aronowitz — A Method for Isolation of Stromal Vascular Fraction Cells in a Clinically Relevant Time Frame

Dr. Joel Aronowitz

Dr. Joel Aronowitz

Ryan A. Lockhart

Cloe S. Hakakian

Abstract

There is increasing interest in the clinical applications of adipose-derived stem cells (ASCs) and the stromal vascular fraction (SVF) based on promising preclinical data. As adipose-derived therapeutics begin to translate into the clinical setting, it is important to maintain patient safety as well as uniformity in technique. Here, we describe a method for isolation of stromal vascular fraction cells in a clinically relevant time frame. Analytical laboratory techniques are mentioned, but respective protocols are not provided here.

Introduction

Adipose-derived stem cells (ASCs) have drawn significant attention in the past decade for potential clinical applications. The combined paracrine effects and multipotent differentiation capacity of ASCs [1, 2, 3, 4, 5, 6, 7] has been examined for a variety of therapeutic clinical applications including breast reconstruction/augmentation, facilitation of diabetic ulcer healing, facial rejuvenation, various orthopedic conditions, repair of cardiac muscle tissue after myocardial infarction, and many more [8, 9, 10, 11, 12]. Since many of these applications have shown promise for clinical application based on findings in the laboratory setting, both in vitro and in animal models, groups around the world are beginning to translate these technologies into the clinical setting. Therapies based on adipose-derived stem cells begin with the isolation of the stromal vascular fraction (SVF) cells. Stromal vascular fraction cells can be isolated in two different ways: with and without the use of tissue dissociation enzymes. Nonenzymatic methods tend to be relatively inexpensive compared to their enzymatic counterparts, but yield fewer cells per milliliter of lipoaspirate processed. Additionally, the resulting composition of the cell populations is different, with mechanical methods yielding a lower frequency of progenitor cells and a higher frequency of CD45 cells [13, 14, 15, 16]. Overall, the proteolytic disruption of the extracellular matrix allows for greater cell recovery. While seemingly the preferred method, enzymatic isolation adds additional factors to consider in regards to patient safety in the clinical setting, including residual enzyme activity and allergic reaction to tissue dissociation enzymes. After SVF isolation, therapies can use the SVF cells for treatment or cultural expanded cells. Cultural expansion offers the advantage of a more homogeneous population and a greater number of ASCs, but requires a minimum of two procedures to complete treatment: one for liposuction/harvest and a second for treatment after the cells have been culturally expanded. Using SVF has the advantage of being able to be completed in a single procedure, including liposuction, isolation and treatment. Here, we describe a method for isolation of SVF cells in a clinically relevant time frame of 60–90 min. This method is able to isolate an average of 2.0 × 10 nucleated cells per milliliter of lipoaspirate with average viability >90%.

Materials 2.1. Supply List for Isolation

1. Package of sterile, 25 count 50 mL disposable flip-top centrifuge tubes. AQ2
2. 1000 cc bags of sterile lactated Ringer’s solution (130 mmol/L Na , 109 mmol/L Cl , 28 mmol/L lactate, 1.5 mmol/L Ca , 4 mmol/L K ).
3. 500 mL glass beakers (for waste and buffer).
4. 500 mL separatory funnel with ring stand.
5. Disposable, individually packaged, sterile latex gloves (as needed).
6. Sterile, individually packaged 3 cc syringe with 18 gauge needle.
7. Sterile, individually packaged 1 cc syringe with 18 gauge needle.
8. 12 mL sterile water.
9. Four vials Vitacyte CIzyme AS (35 Wünsch units collagenase activity per vial).
10. 100 μm cell strainer, sterile, individually packaged.

Enzyme Preparation

1. Using a sterile 3 cc syringe, transfer 3 cc of sterile water at room temperature into each of the four vials of lyophilized tissue dissociation enzymes in order to resuspend (see Note 1).
2. Once the enzyme has fully dissolved, remove the enzyme solution from vials using the 3 cc syringe and transfer the contents of each vial into 50 mL centrifuge tubes. Raise the volume of each of the four tubes to a final volume of 50 cc using warm lactated Ringer’s solution.
3. Place tubes into the heated shaker for a minimum of 30 min at 37 °C and 200 rpm. Be sure to keep warm and shaking until use.

Tumescent Solution Preparation Tumescent solution is prepared by a trained surgical technician in the operating room.

1. Fifty cubic centimeters of lidocaine plain is injected into a sterile 1000 cc bag of saline solution.
2. Two milliliters of AMP epinephrine 1:1000 solution is injected into the saline/lidocaine solution.

Methods

All preparation and isolation occurs inside of a P&C Multi-Station (Fig. 1). The Multi-Station contains a centrifuge (400 cc capacity) and a heated shaking unit contained within the confines of a laminar flow hood. All procedures and preparations should be conducted inside of the laminar flow hood. All glassware is autoclaved prior to use. The laminar flow hood should be fully sterilized before and after isolation. Sterile gloves should be worn at all times to avoid contamination of samples.

Fig. 1 Inside the P&C Multi-Station. The left side contains a centrifuge with a 400 cc capacity. The right side contains a heated shaking unit

Dr. Joel Aronowitz — Inside the P&C Multi-Station. The left side contains a centrifuge with a 400 cc capacity. The right side
contains a heated shaking unit

Lipoaspiration

All equipment/supplies required for liposuction are not included in this protocol as they can vary from facility to facility. Suction-assisted liposuction was conducted using a “super-wet” tumescent solution (see Subheading 2, item 3) and a 2.7 mm blunt tipped cannula with a vacuum pressure of −25 mmHg.

1. Harvest site varies from patient to patient based on donor site availability and aesthetic desires of the patient. Donor site selection is left to the discretion of the patient and surgeon.
2. Prior to harvest, the patient is infiltrated with 750 cc–1000 cc of tumescent solution (see Subheading 2, item 3 for recipe) to limit blood loss, pain and bruising due to harvest. Tumescent is allowed to infiltrate for 10–15 min prior to harvest.
3. Using a 2.7 mm blunt tipped cannula, fat tissue is aspirated into sterile canisters (see Note 2).
4. Lipoaspirate is then transferred into a sterile 500 mL separatory funnel for isolation. Isolation is carried out by a technician in a sterile laminar flow biohood (see Note 3).

Isolation of the Stromal Vascular Fraction

1. Wash lipoaspirate with an equal volume of warm (37 °C) lactated Ringer’s (LR) solution. After addition of LR, gently mix and allowed the mixture to separate. The mixture will separate into two distinct portions, a lower aqueous layer (Red/clear) and the upper fatty layer (yellow/pink). The lower, aqueous layer is discarded. Wash three times or until fat layer is yellow (no longer pink) (Fig. 2).
2. Aliquot 25 cc washed lipoaspirate into sterile 50 cc centrifuge tubes (see Note 4).
3. Add 25 cc of warm (37 °C) tissue dissociation enzyme solution to each 25 cc aliquot of washed lipoaspirate. Invert to mix.
4. Incubate lipoaspirate/enzyme solution in a heated shaker at 37 °C, 200 rpm for 20–30 min (see Note 5).
5. Following incubation, centrifuge the tubes at 2000 rpm (700 × g) for 10 min to separate components.
6. After centrifugation, three layers are distinctly visible in the tubes. The uppermost layer will contain fat and oil. The middle layer will be the aqueous layer, which will appear red/clear. At the bottom of the tubes there is a pellet. The pellet contains the stromal vascular fraction cells (Fig. 3).
7. Remove the aqueous and fatty/oil layers. Dispose of all biohazard materials properly in biohazard waste containers.
8. Combine the SVF pellets into two sterile 50 cc tubes (four pellets into each tube). Pellets are combined by suspending them in a small amount of warm LR (see Note 6).
9. Bring each tube containing pellets to a volume of 50 cc using warm LR (see Note 7).
10. Washing: Centrifuge at 700 × g for 5 min. Save the pellet and pour off the aqueous layer. Repeat two more times.
11. After the third and final wash, combine the two remaining pellets into a single tube. Adding a small amount of LR may make the transfer easier.
12. Filter through a 100 μm cell strainer in order to remove tissue fragments which remain from the digestion of the adipose tissue. Use LR to wash the cells through the strainer (Fig. 4) (see Note 8).
13. Bring final volume to 15 cc using warm LR (see Note 9).
14. The following volumes are taken for analysis (see Note 10): (a) 0.1–0.2 cc for cell count and viability using Chemometec NC-200 (see Note 11) (Fig. 5) (b) 2.1 cc for sterility testing Gram stain: <0.1 cc SVF needed 2 week aerobic culture: 1 cc SVF in 9 cc Tryptic Soy Broth (TSB). 2 week anaerobic culture: 1 cc SVF in 9 cc Thioglycollate broth. © 2 cc for flow cytometry Markers tested: FITC-CD31, APC-CD34, PE-CD45.
15. The remaining volume is returned to the surgeon.

Fig. 2 Lipoaspirate separated in the separatory funnel. The upper, yellow layer is lipoaspirate. The lower pink layer is a mixture of blood and tumescent

Dr. Joel Aronowitz — Lipoaspirate separated in the separatory funnel. The upper, yellow layer is lipoaspirate. The lower pink layer
is a mixture of blood and tumescent

Fig. 3 The result of centrifuging the lipoaspirate after enzymatic digestion

Dr. Joel Aronowitz — The result of centrifuging the lipoaspirate after enzymatic digestion

Fig. 4 The left tube contains the detritus and tissue fragments which have been removed via the 100 μm cell strainer. The right tube contains the strained SVF

Dr. Joel Aronowitz — The left tube contains the detritus and tissue fragments which have been removed via the 100 μm cell strainer. The right tube contains the strained SVF

Fig. 5 A representative readout from the nucleocounter NC-200. The sample tested was a 10% dilution of the final SVF output. The final volume of SVF was 10 cc total. 400 cc of lipoaspirate was processed. The total nucleated cell count for the whole SVF sample was 128.4 million nucleated cells. 321,000 nucleated cells were isolated per cc of washed lipoaspirate processed

Dr. Joel Aronowitz — A representative readout from the nucleocounter NC-200. The sample tested was a 10% dilution of the final SVF output. The final volume of SVF was 10 cc total. 400 cc of lipoaspirate was processed. The total nucleated cell count for the whole SVF sample was 128.4 million nucleated cells. 321,000 nucleated cells were isolated per cc of washed lipoaspirate processed

Notes

1. The tissue dissociation enzyme mixture used here contains collagenase and neutral protease (Dispase). The level of collagenase activity is 35 Wünsch units [17] per vial, but the total collagen degrading activity is reported to be 330,000 units [18] per vial. Make sure that enzymes are of GMP grade. Be aware that the level of activity and the units used to express the amount of activity varies between manufacturers and activity may vary from batch to batch from the same manufacturer. The certificate of analysis received with enzymes will give the exact level of activity. Proper enzyme preparation may vary between products and should be done according to the recommendations by the manufacturer.
2. Volume of lipoaspirate is measured as volume after washing. Larger volumes are usually received from the surgeon, but this often contains a significant amount of tumescent solution which is removed during washing. Using this method, we have processed between 50 cc and 800 cc of washed lipoaspirate.
3. A sterile funnel may make the transfer easier for larger volumes of lipoaspirate.
4. Transfer the washed lipoaspirate from the separatory funnel to a 500 mL beaker. It is easier to aliquot washed lipoaspirate from a beaker as opposed to a separatory funnel.
5. Tubes may need to be manually inverted and mixed every 5–10 min, as the lipoaspirate may collect at one end of the tube. Mixing will provide more even digestion of tissue.
6. The number of pellets varies based on the amount of fat tissue processed. Every 25 cc of fat = 1 pellet. 200 cc of lipoaspirate = 8 pellets.
7. In a previous study we were able to demonstrate negligible collagenase levels after three washing steps [19]. If desired, collagenase can be neutralized using autologous serum. This is done by washing in 20% serum which can be made using 10 cc serum and 40 cc LR for one of the wash steps. Serum can be generated by centrifuging heparinized blood at 800 × g for 10 min.
8. The amount of detritus varies depending on the volume of lipoaspirate processed. If sample contains a large amount of tissue fragments, the strainer may become clogged and the SVF will not flow through. If this happens an additional strainer may be used to filter the remaining portion. Not all cells are recovered after straining, as some remain trapped within the pellet of tissue fragments. An additional washing and straining of the detritus can be conducted to increase cell yield, but this will extend the isolation time.
9. A final volume of 15 cc is arbitrary and can vary based on personal preference. When mixing SVF with fat for Cell-assisted Lipotransfer (CAL), final volume is not as important because there is usually a larger volume of fat, but when directly injecting SVF, a smaller volume (5 cc–10 cc) may be desired.
10. Samples with <80% viability will not be approved for clinical use. Samples which detect any contamination during the initial gram stain will not be approved for clinical use. A patient whose sample has a positive aerobic/anaerobic culture will be closely monitored for infection. A positive culture does not mean that a patient will develop an infection.
11. A dilution of the final SVF product may be needed to achieve a cell concentration in the readable range for the cell counting method used. Typically a 1:10 dilution in LR was sufficient, but in a few rare cases a 1:100 dilution may be required.

None of the authors have a financial interest in any of the products, devices, or drugs mentioned in this chapter.

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