Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

Samples (n = 3) were obtained from the consenting, healthy donors through the Beaumont Hospital BioBank, Royal Oak, MI and the Providence Hospital, Southfield, MI under HIC- (HIC #2012-101) and IRB- (IRB# 820662-3), respectively, approved protocols and processed as approved by the IBC (#2858) of Oakland University, Rochester, MI.

1. Processing Human Umbilical Cord, Cord-placenta Junction, and Fetal Placenta and Isolating Cells

1. Aseptically collect samples of UC approximately 10 cm in length, including 5 – 6 cm connected to the CPJ and 3 – 4 cm of FP. Immediately place each sample in a collection cup containing medium (DMEM with 4,500 mg/mL glucose and antibiotic solution (0.1% gentamicin, 0.2% streptomycin, and 0.12% penicillin)) and store at 4 °C until it is transported to the lab by placing it on ice in a Styrofoam box. Process the sample at 4 °C within 2 – 4 h of collection.
2. Place the sample in a 150-mm Petri dish kept on ice in a biosafety cabinet. Using a needle and syringe, rinse it multiple times with ice-cold PBS to remove blood clots. Make sure the sample is kept wet with PBS during processing and do not let it dry. To maintain sterility, handle the sample aseptically using autoclaved PBS and surgical tools throughout sample processing.
3. Carefully examine the sample to identify different anatomical regions: UC, CPJ, and FP. First dissect the UC. Hold the fetal end of the UC with forceps and carefully make the first cut just at the top of the CPJ using a pair of scissors. Make the second cut below the CPJ to separate the CPJ (1.5 – 2.0 cm) from the FP. Place the separated UC, CPJ, and FP in separate Petri dishes for further processing.
4. Cut the UC longitudinally with the help of scissors and forceps, completely exposing the blood vessels and surrounding WJ without disturbing the epithelium.
5. Scrape the WJ away from the blood vessels and inner epithelium of the subamnion using a scalpel, and then remove the blood vessels. Be sure to collect any remaining perivascular WJ under and around the blood vessels, and place the collected WJ in a separate Petri dish.
6. Collect the remaining tissue, cord lining (CL), in a separate Petri dish.
7. After separation of the tissues representing CL, WJ, CPJ, and FP, replace the PBS with 3 – 5 mL of commercial trypsin solution and separately cut each of the tissues into 1- to 2-mm pieces using scissors.
8. Incubate the tissue pieces in commercial trypsin solution at 37 °C for 30 min in a 5% CO₂ incubator for partial digestion of the samples. Observe the partial digestion by visualizing release of cells using a phase contrast microscope.
9. To the partially-digested samples, add an equal volume of culture medium (CM; DMEM with 4,500 mg/mL glucose and 2 mM L-glutamine, supplemented with 10% human serum/FBS and antibiotic solution (0.1% gentamicin, 0.2% streptomycin, and 0.12% penicillin)) to neutralize the commercial trypsin solution. Transfer the content into 50-mL centrifuge tubes and let the partially-digested tissue pieces settle for 3 min.
10. Carefully aspirate the supernatant, including single cells (they do not expand efficiently), and plate 15 – 20 partially-digested tissue pieces on a 75-cm² tissue culture flask and add 9 mL of CM. Incubate at 37 °C and do not disturb the culture flasks for 2 – 3 days to allow for the adherence of tissue pieces.
    NOTE: The remaining portion of the partially-digested tissue samples can be cryopreserved for the future isolation of cells.
11. After 3 days of incubation, change the CM and look for the appearance of outgrowth of cells from the explant on a daily basis; cell growth should be evident from the adherent explants after 3 – 4 days, 7 – 10 days, and 11 – 14 days of incubation for CPJ, CL/WJ, and FP, respectively. From this point onwards, change the CM after every 3 days or as needed.
    NOTE: Cell growth reaches 70% confluence 7 – 10 days after the initial cell outgrowth from the explant. Note that non-adherent tissue pieces will not give rise to any cell outgrowth until they adhere to the plastic. Care must be taken so that the flask is not disturbed in order to avoid the detachment of the adherent tissue pieces. If there are any floating pieces after the first 3 days of culturing, they could be rescued by transferring them to a new flask for attachment.
    NOTE: When cell growth from the tissue explants reaches 70% confluence, they should be subcultured. As stated above, MSCs from CL/WJ, FP, CPJ and will reach confluence in 10 – 14 days, 14 – 20 days, and 17 – 24 days, respectively.
12. To subculture outgrown cells from the explants, dissociate the cells by using 1 – 2 mL of commercial trypsin solution/75-cm² culture flask and incubate at 37 °C for 3 min. Neutralize with 1 – 2 mL of CM and centrifuge 1,500 x g for 3 min. Make sure to rotate the flask while adding trypsin for even spreading throughout.
13. NOTE: The pelleted cells are considered passage 0 (P0) and will be suspended in CM, counted, and subcultured at a seeding density of 1 x 10⁴/cm² for amplification, characterization, and cryopreservation. The excess cells can be cryopreserved at P0.

2. Characterizing the Isolated Cells

1. Colony forming efficiency (CFE) assay
   1. Coat the 100-mm Petri dishes (surface area of 60 cm²) with 0.1% gelatin for 30 min and place them in the CO₂ incubator at 37 °C.
      NOTE: Although not necessary, gelatin-coating improves cell adherence.
   2. Remove the excess gelatin and seed the coated plate with P0 cells at a concentration of 1.63 cells/cm². Add 3 mL of CM and incubate in the CO₂ incubator at 37 °C.
   3. Monitor the seeded plates for clonal growth by light microscopy (LM) and change the medium every 3 days.
   4. After 10 – 14 days of cell growth, wash the Petri dishes twice with PBS and fix the cells in 4% paraformaldehyde for 30 min at room temperature.
      NOTE: Paraformaldehyde is toxic. Please practice caution by wearing gloves and eyewear during use.
   5. Stain the fixed cells with 0.1% crystal violet for 1 h at room temperature and rinse the plate with tap water until the unbound blue stain is gone.
   6. Count the number of colonies comprised of at least 50 cells using LM.
2. Expression of cell surface markers
   1. Culture the isolated cells to 70% confluence, dissociate with the commercial trypsin solution, and wash and pellet by centrifugation at 1,500 x g for 3 min. Suspend the cells in FACS buffer (1x PBS with 2% FBS and 0.1% sodium azide).
      NOTE: Sodium azide is toxic. Please practice caution by wearing gloves and eyewear during use.
   2. Stain the cells (~ 10⁶) with FITC- or APC-labeled MSC-specific antibodies (FITC-conjugated antibodies against: CD44 and CD90 or APC-conjugated antibodies against: CD29, CD73, and CD105) in FACS buffer for 30 min at 4 °C in dark.
   3. Centrifuge the stained cells at 670 x g for 5 min, aspirate the supernatant, and resuspend the cells in 500 µL of FACS buffer.
   4. Analyze the antibody-stained cells using FACS according to the manufacturer's protocol.

3. Differentiation Potential

1. Adipogenic differentiation
   1. Culture the isolated cells to 70% confluence, dissociate with the commercial trypsin solution, and wash and pellet by centrifugation at 1,500 x g for 3 min.
   2. In a CO₂ incubator at 37 °C, culture the cells at a concentration of 100,000 cells/well of a 6-well plate containing 2 mL of CM.
   3. After 24 h, replace the CM with adipogenic differentiation medium (0.5 µM isobutyl-methylxanthine, 1 µM dexamethasone, 10 µM insulin, and 200 µM indomethacin) and incubate. Change the differentiation medium every 3 days or as often as needed.
   4. After 3 weeks of incubation, fix the cells with 2 mL of 4% paraformaldehyde per well for 30 min at room temperature and stain with Oil Red O to detect the production of lipid droplets in order to analyze for adipogenic differentiation.
      NOTE: Paraformaldehyde is toxic. Please practice caution by wearing gloves and eyewear during use.
   5. Treat the fixed cells with 60% isopropanol (2 mL/well of 6-well plate) for 5 min.
   6. Replace the isopropanol with 2 mL of Oil Red O stain (filter 3 parts Oil Red O stain and 2 parts distilled water), incubate for 15-30 min at room temperature, and wash 3 – 4 times with tap water to remove any unbound stain.
   7. Visualize the red-stained lipid droplets, indicative of adipogenic cell lineage, using LM.
2. Chondrogenic differentiation
   1. Culture isolated cells to 70% confluence, dissociate with the commercial trypsin solution, and wash with PBS.
   2. To pellet cells for chondrogenic induction, centrifuge 250,000 cells in a 15-mL centrifuge tube with 2 mL of CM at 11,000 x g for 8 min. Incubate the pellets overnight at 37 °C.
   3. After 24 h, gently replace the CM with chondrogenic differentiation medium (20 ng TGFβ1, 10 ng insulin, 100 nM dexamethasone, and 100 µM ascorbic acid) without disturbing the pellet and continue the incubation. Change the differentiation medium every 3 days or as often as needed.
   4. After 3 weeks of incubation, fix the cells with 2 mL of 4% paraformaldehyde per well for 30 min at room temperature. Cryosection and stain with 1% toluidine blue to examine the presence or production of extracellular matrix in order to determine chondrogenic differentiation.
      NOTE: Paraformaldehyde is toxic. Please practice caution by wearing gloves and eyewear during use.
   5. For cryosectioning, gently wash the harvested cell pellets twice with PBS and fix with 1 mL of 4% paraformaldehyde for 30 min at room temperature.
      NOTE: Paraformaldehyde is toxic. Please practice caution by wearing gloves and eye wear during use.
   6. Wash the pellets twice with PBS to remove the paraformaldehyde and transfer into a sucrose/OCT solution (1:2, 20% Sucrose:OCT). Incubate at room temperature for 24 h to allow the solution to completely seep into the cell pellets; this will allow smooth sectioning.
   7. Transfer the pellets into the OCT molds. Freeze the molds at -20 °C for 4 – 6 h and cryosection about 10 – 12 micron-thick sections of the cell pellets using a cryostat for toluidine blue staining.
   8. Wash the cryosections gently with PBS; add few drops of 1% toluidine blue stain, covering the pellet sections on the slide completely; and incubate for 1 h at room temperature.
   9. Wash the slides with PBS multiple times to destain. Remove the excess stain by dipping the slide in HCl-based alcohol solution (95% alcohol + 0.5 mL of HCl) multiple times until the unbound blue color is gone. Mount the stained sections using 100 µL of mounting solution for the long-term preservation of the slides.
      NOTE: The blue staining of the sections indicates the presence of glycosaminoglycans and glycoproteins and will be observable using LM.
3. Osteogenic differentiation
   1. Again, culture the isolated cells to 70% confluence, dissociate with the commercial trypsin solution, and wash and pellet by centrifugation at 1,500 x g for 3 min.
   2. Subculture the cells at a concentration of 100,000 cells/well of 6-well plates containing 2 mL of CM in a CO₂ incubator at 37 °C.
   3. After 24 h, replace the CM with osteogenic differentiation medium (0.1 µM dexamethasone, 10 µM β-glycerophosphate, and 50 µM ascorbate phosphate) and continue the incubation. Change the differentiation medium every 3 days or as often as needed.
   4. After 3 weeks of incubation, fix the cells with 2 mL of 4% paraformaldehyde per well for 30 min at room temperature and stain with alizarin red to detect the presence of a calcium deposition in order to determine osteogenic differentiation.
      NOTE: Paraformaldehyde is toxic. Please practice caution by wearing gloves and eyewear during use.
   5. Gently wash the fixed cells twice with distilled water and treat with 2% alizarin red stain (2 mL/ well in a 6-well plate) for 1 h at room temperature in the dark.
   6. Remove the excess stain by gently washing with tap water so that the calcium deposits do not dislodge.
   7. Visualize the red-stained calcium deposits using LM.

4. Quantitative Reverse Transcriptase Polymerase Chain Reaction (qRT-PCR) Analysis

1. Culture the isolated cells to 70% confluence, dissociate with the commercial trypsin solution, wash, and pellet by centrifugation at 1,500 x g for 3 min to isolate the total cellular RNA.
2. Wash the cells with PBS to remove traces of FBS, and then proceed to the RNA isolation using a commercial RNA purification kit, following the manufacturer's instructions.
3. Purify the isolated total RNA by treating with DNase at 37 °C for 30 min using a thermocycler. Synthesize cDNA using commercial kit following the manufacturer's instructions.
4. Prepare triplicate 10-µL PCR reactions in a 96-well plate, with each reaction containing 5 µL of SYBR-green PCR supermix; 3 µL of double-distilled water; 0.5 µL of each primer, forward and reverse; and 1 µL of the diluted (1:10) cDNA.
5. Perform PCR under the following parameters: 10 min at 98 °C, followed by 44 cycles of 30 s each at 98 °C, 20 s at 60 °C, and 30 s at 72 °C.
6. Normalize the amplification of the target genes to the reference genes, GAPDH and β-ACTIN; the primer sequences are listed in Table 1.

5. Statistical Analysis

1. Perform all analyses using statistical software. Present the data as the mean ± SEM. Results with a p-value (**p ≤ 0.01 and *p ≤ 0.05) were considered statistically significant.