Isolation and Characterization of Adipose-Derived Stromal Cells

Abstract

Methods for adipose-derived stromal cell (ASC) isolation, characterization and the respective data generated differ between different research groups. Laboratories have developed in-house methods to isolate, expand and differentiate ASCs, which often makes data comparison difficult.

Methods for both research laboratory isolation and clinical applications are explored in this chapter. To use ASCs in the clinical setting, stem cell therapy products need to be prepared using good manufacturing practices (GMPs). This is achieved by eliminating animal-derived products from the manufacturing process. As a result, several human and chemically defined alternatives have been explored, especially during the isolation and expansion process.

ASC characterization criteria remain open ended, but should be compliant within the parameters of (1) adherence to plastic, (2) expression of a defined set of surface antigens and (3) the capacity to differentiate into various cell types.

Appendix: Brief Description of Current Methods

7.1.1 Manual Isolation Procedure

The isolation protocol is adapted from the procedures described by Zuk et al. (2001), Bunnell et al. (2008) and Estes et al. (2010) (Fig. A.1).

Fig. A.1

Outline of the procedure used f or isolating adipose-derived stromal cells. Lipoaspirate samples are processed by enzymatic digestion, and the stromal vascular fraction (SVF) is collected. Adipose-derived stromal cells from the SVF adhere to the plastic culture dish, and non-adherent cells are washed away after 24 h

The method requires the transfer of lipoaspirate material into 50 ml tubes (30 ml lipoaspirate/tube), followed by the addition of 20 ml phosphate buffer saline (PBS), supplemented with antibiotics (Fig. A.1). Depending on the future application of the adipose-derived stromal cell (ASC) isolates, culture medium may contain either FBS or human derivatives and antibiotics, usually penicillin and streptomycin (pen/strep). The adipose tissue is separated from peripheral blood contaminants by centrifugation at 1660 g for 3 min. The top oil layer is aspirated with a suction-assisted glass pipette system, and the compact lipoaspirate is carefully transferred to a sterile 50 ml tube. It is recommended that the washing steps are repeated until the compacted lipoaspirate material is golden yellow in colour without any visible evidence of peripheral blood contamination. The volume of the compacted lipoaspirate material should be recorded once most of the blood contamination has been removed.

To release resident ASCs in adipose tissue from the fibrous network, the lipoaspirate is enzymatically digested in culture plates. The most popular approach makes use of the enzyme, collagenase type I. Other enzymatic alternatives include dispase and trypsin. A filter-sterilized 0.1 % collagenase type I solution is prepared using PBS supplemented with 2 % antibiotics. The volume of collagen solution required is dependent on the volume of the compacted adipose tissue previously recorded. The ratio of compacted adipose tissue volume to the volume of collagen solution should be at least 2:1, meaning that the final volume of collagen digesting solution added to the adipose tissue in the culture plates should be half that of the washed compacted adipose tissue volume. The ratio of collagenase to adipose tissue should be optimized by each laboratory as well as for each isolation technique used. A sterile plastic pipette is used to mix the adipose tissue well, before incubation at 37 °C, 5 % CO₂ for 45 min. The sample may either be continuously agitated using an automated rotating system or agitated manually every 15 min with a plastic pipette to aid the mechanical breakdown of the fibrous tissue.

The collagenase-digested sample is then transferred to sterile 50 ml tubes. The tubes are shaken vigorously and centrifuged at 500 g for 5 min, resulting in the SVF pellet settling to the bottom of the tube. The compacted adipose tissue and collagenase solution supernatants are carefully aspirated, and the collagenase activity is neutralized by adding 2 ml of stromal medium to the AD-SVF pellet. Stromal medium may consist of either Dulbecco’s Modified Eagle Medium (DMEM) containing GlutaMax™, 4.5 g/L D-glucose and pyruvate or alpha-Modified Eagle Medium (α-MEM) containing GlutaMax™, supplemented with 10 % serum and 1 % antibiotics.

The AD-SVF pellets are pooled into a single tube (15 or 50 ml) and centrifuged at 265 g for 5 min, followed by aspiration of the supernatant. Red blood cells present in the AD-SVF pellet are lysed either by the addition of an ammonium chloride-based lysing solution or an enzymatic-based lysing solution like VersaLyse™ (Beckman Coulter, Miami, USA). After a 10–15 min incubation period at room temperature, the lysing reaction is stopped by filling the tube with PBS supplemented with antibiotics and centrifuged at 184 g for 5 min. The supernatant is removed and the pellet resuspended in stromal medium before filtering the cellular suspension through a 70 μm cell strainer to remove any larger, undigested fragments.

In order to seed the cells at the correct seeding density, an absolute cell count should be performed. Cell counts may be performed by using either a manual approach in which the cells are counted using a haemocytometer or an automated cell counting device such as a flow cytometer. Details are provided below.

The AD-SVF is seeded at an initial seeding density of 5 × 10⁵ cells per cm². In order to determine the volume of cell suspension required for initial seeding, the following formula should be used:

$$ \begin{array}{l}\mathrm{Volume}\ \left(\mu \mathrm{l}\right)=\left(\frac{\mathrm{initial}\ \mathrm{seeding}\ \mathrm{density}\times \mathrm{seeding}\ \mathrm{surface}\ \mathrm{area}\ \mathrm{of}\ \mathrm{tissue}\ \mathrm{culture}\ \mathrm{flask}\ \mathrm{or}\ \mathrm{well}}{\mathrm{total}\ \mathrm{number}\ \mathrm{of}\ \mathrm{viable}\ \mathrm{cells}}\right)\\ {}\kern8.25em \times \mathrm{cell}\ \mathrm{suspension}\ \mathrm{volume}\ \mathrm{in}\ \mu \mathrm{l}\end{array} $$

After the cells are seeded, the cultures flasks are swirled gently to ensure uniform distribution. The culture flasks are maintained in an incubator under standard cell culture conditions (humidity, 5 % CO₂ and 37 °C). The cultures are washed twice after 24 h with PBS supplemented with antibiotics to remove non-adherent cells, cellular debris and non-viable cells. Fresh stromal medium is added to the culture flasks and incubated under standard conditions until cells are 80–90 % confluent, implying that cells cover 80–90 % of the culture flask surface area. In order to determine the volume of cell suspension required for reseeding after thawing frozen samples, the following formula should be used:

$$ \begin{array}{l}\mathrm{Total}\ \mathrm{number}\ \mathrm{of}\ \mathrm{viable}\ \mathrm{cell}\mathrm{s}\ \mathrm{in}\ \mathrm{cell}\ \mathrm{s}\mathrm{uspension}\div \hfill \\ {}\kern1em \left(\mathrm{reseeding}\ \mathrm{density}\times \mathrm{reseeding}\ \mathrm{s}\mathrm{urface}\ \mathrm{area}\ \mathrm{of}\ \mathrm{tissue}\ \mathrm{culture}\ \mathrm{flask}\ \mathrm{or}\ \mathrm{well}\right)\hfill \end{array} $$

7.1.2 Methods Used to Obtain Absolute Cell Counts

7.1.2.1 Trypan Blue (0.4 %) Dye Exclusion Method

Trypan blue is a non-membrane permeable vital stain that is used to assess the viability of cells. Trypan blue is not able to cross the cell membrane of intact, viable cells and therefore only stains cells with compromised cell membranes. Dead cells display a distinct blue colour after staining, due to the accumulation of trypan blue in the cytoplasm of cells with compromised cell membranes The recommended procedure is as follows: (1) prepare the Neubauer counting chamber (haemocytometer) by carefully placing the cover slip on the counting grid; (2) carefully mix 80 μl of a 0.4 % trypan blue with 100 μl PBS and 20 μl of the suspension; (3) carefully load 10 μl of the solution onto both sides of the Neubauer counting chamber (haemocytometer); and (4) count the viable (unstained) cells as well as dead cells (stained) using a microscope (10 times objective lens). The following formula is used to determine the absolute cell concentration (Fig. A.1):

Absolute cell concentration (cells/ml) $=\left(\frac{Q1+Q2+Q3+\dots +Q8}{8}\right)\times$, where Q1–Q8 refer to eight quadrants on the haemocytometer. The factor 10 is to correct for the dilution of the sample with the 0.4 % trypan blue solution.

There are several commercial automated counting devices on the market that make use of trypan blue (0.4 %) dye exclusion assay principles. Examples of such devices are the Vi-Cell XR™ automated cell counter (Beckman Coulter, Miami, USA), Countess™ automated cell counter (Invitrogen, Carlsbad, USA) and TC20™ automated cell counter (Bio-Rad, Hercules, USA).

7.1.2.2 Absolute Cell Count Determination Using a Benchtop Flow Cytometer (Beckman Coulter Flow Cytometers)

An example of the strategy that is followed to obtain an absolute count on a benchtop flow cytometer is illustrated in Fig. A.2. The cell population of interest is identified by using a side scatter linear (SS lin) and a forward scatter linear (FS lin) histogram by placing a region around the cell population of interest only, excluding the counting beads and debris. The counting beads are enumerated in the CAL region. In order to obtain an accurate absolute count, it is important to count a sufficient number of cells as well as counting beads (>1000 events if possible) as well as to perform the count as soon as possible after the counting beads are mixed with the sample.

Fig. A.2

(a) A side scatter linear and forward scatter linear dot histogram displaying all the events measured by the flow cytometer. The flow beads (pink) and a gate were used to encircle the cell population that was counted until the CAL factor was reached. The gate-labelled cell population displays the cell population count that was expressed as the number of cells per μl cell suspension. (b) A histogram displaying the flow beads with a region of interest placed over the peak of the flow cytometry counting beads labelled as CAL. In this example, the specific calibration factor (assayed bead concentration) was 986

7.1.3 Induction of Adipogenesis In Vitro

ASCs are seeded at a density of 5 × 10³ cells/cm² in a six-well plate and maintained under standard culturing conditions (37 °C and 5 % CO₂) with complete stromal medium (α-MEM or DMEM, containing 10 % FBS and 1 % pen/strep). When 70–80 % confluency is achieved, the stromal medium is removed and replaced with adipogenic induction medium (Table A.1) in three of the wells of the 6-well plate. DMEM supplemented with serum and antibiotics is added to the remaining three wells. These wells serve as non-induced controls. The cultures are maintained for 21 days under standard culturing conditions of 37 °C, 5 % CO₂. During this period the induction and control media are replaced every second day.

Table A.1 Composition of induction media for adipogenic, osteogenic, chondrogenic and myogenic differentiation in vitro

7.1.3.1 Qualitative Assessment of Adipogenesis In Vitro Using Oil Red O

After 21 days of induction, the 6-well plates containing both induced and non-induced cultures are fixed by adding a 4 % formaldehyde solution for 1 h. A 0.5 % Oil Red O stock solution is prepared in isopropanol. An Oil Red O working solution is prepared from the stock solution by mixing three parts of the stock solution with two parts of double-distilled water (ddH₂O) (volume/volume). After removal (aspiration) of the fixative solution, the cultures are allowed to dry at room temperature before adding 1 ml of Oil Red O working solution to both the adipogenic induced and non-induced cultures, followed by a 20 min incubation at room temperature. The stain is removed, and the wells are thoroughly washed with ddH₂O until no pink discoloration of the freshly added ddH₂O is visible to the naked eye.

The cultures are then counterstained with Toluidine Blue O. A 0.01 % Toluidine Blue O counterstain solution is prepared (pH 11) by adding 0.005 g Toluidine Blue O and 0.005 g Na₂CO₃ to 50 ml ddH₂O. The Toluidine Blue O counterstain is added to the Oil Red O-stained cultures for 5 min at room temperature, after which the excess stain is washed away with ddH₂O. 1 ml ddH₂O is added to each well before microscopy analysis.

7.1.4 Induction of Osteogenesis In Vitro

Immunophenotyped ASCs are seeded at a density of 5 × 10³ cells/cm² in a six-well plate and maintained under standard culture conditions (37 °C and 5 % CO₂) with stromal medium (α-MEM, containing 10 % FBS and 1 % pen/strep) until about 60–70 % confluency. Osteogenic induction medium (Table A.1) is introduced to half the wells and DMEM supplemented with serum and antibiotics to the other half to serve as non-induced (control) cultures. The cultures are maintained for 21 days under standard culture conditions of 37 °C, 5 % CO_2. The induction and non-induction media are replaced every second day.

7.1.4.1 Qualitative Assessment of Osteogenesis In Vitro

Twenty-one days after induction of osteogenic diffe rentiation, the cells are fixed by addition of a 4 % formaldehyde fixative solution for 1 h. A 2 % Alizarin Red S classical stain is used to detect the calcium in the mineral matrix from mature osteocytes. An alizarin stock solution is prepared by adding 2 g Alizarin Red S powder to 100 ml of ddH₂O. The solution is mixed thoroughly using a magnetic stirrer until solutes are dissolved before filtering through filter paper.

The induced and non-induced cultures are pre-washed with 2 ml PBS at pH 4.2 for 5 min before introducing 2 ml of the 2 % Alizarin Red S stain and incubating the cultures for 10 min at room temperature. The cultures are washed thoroughly with ddH₂O to remove the excess stain. 1 ml ddH₂O is added to each well before microscopy analysis.

7.1.5 Induction of Chondrogenesis In Vitro

A suspension culture technique is usually used for the differentiation of ASCs into chondrocytes. ASCs are seeded in a T25 flask at a density of 5 × 10³ cells/cm² and maintained under standard culture conditions until about 60 % confluence. The cells are enzymatically removed from the flask (0.25 % trypsin-EDTA) followed by the neutralization of the enzymatic action with the addition of stromal medium (α-MEM, containing serum and antibiotics).

The cell suspension is transferred into a 15 ml tube, and the sample is centrifuged for 5 min at 400 g. The substrate is carefully aspirated until only the ASC pellet remains in the tube. The ASC pellet is suspended in chondrogenic induction medium (Table A.1), for the chondrogenic induced cultures or DMEM supplemented with serum only and for the non-induced cultures, and centrifuged at 400 g for 10 min. The tubes are carefully placed into the incubator without disrupting the pellet. The tube caps are slightly loosened to allow for gas exchange to occur. Cultures are incubated under standard conditions of 37 °C, 5 % CO₂ for 21 days. The induction and control media (0.5 ml) are replaced every second day. After 24 h, the ASC pellets contract into a sphere. The cells that have not been incorporated into the sphere after 48 h are removed from the suspension cultures during medium replacement.

7.1.5.1 Qualitative Assessment of Chondrogenesis In Vitro

Each induced and non-induced chondrocyte sphere is serially dehydrated in 30, 50, 70 and 90 % ethanol, followed by three changes of absolute ethanol for 15 min per dehydration step. The dehydrated chondrocyte spheres are infiltrated with 50 % LR White Resin in absolute ethanol for 1 h, followed by an infiltration in a 100 % LR White Resin overnight. To embed the tissue spheres, they are transferred into resin capsules with 100 % LR White Resin and polymerized for 24 h at 60 °C.

Ten to 15 serial transverse sections of between 0.5 and 1.0 μm (optimal 0.5 μm) are prepared using a microtome, and the sections are stained with 1 % Toluidine Blue O stain for 30 s. The glass slides containing stained sections are gently rinsed with ddH₂O. One drop of ddH₂O is added on the glass plate and covered using a cover slip before microscopy analysis is performed.