Keratinocyte Colony Formation Assay: An In Vitro Assay to Assess the Ability of Keratinocytes to Grow into Colonies

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Primary Keratinocyte Harvesting and Seeding

1. Euthanize four to five adult female mice via CO₂ inhalation according to approved animal facility/IACUC standards. However, this protocol may also be used for single female/male mice.
   1. Clip approximately 9 cm² of the dorsal fur with an electric animal clipper and place all clipped mice into a 500 mL jar with enough povidone-iodine antiseptic solution (not scrub) to cover them for 1-2 min. Shake the jar well to coat the antiseptic solution over the mice evenly.
   2. Pour off the solution and rinse with clean water until the liquid runs clear. Repeat the same antiseptic wash followed by a water rinse. After the antiseptic wash with iodine solution, soak all the mice in 70% ethanol for 5-10 min.
      NOTE: Mice with light fur (e.g., BALB/c) will retain a slight yellowish color from the antiseptic iodine washes, while darker mice (e.g., C57BL/6) will not. Notably, cell viability or culture growth is not affected due to the yellow color.
2. Carefully remove the clipped dorsal skin using thumb forceps and scissors in a laminar flow hood. Place the excised dorsal skin in a conical tube filled with PBS/2x gentamycin. Avoid using ventral side skin to prevent mammary cell contamination in culture.
3. Using autoclaved forceps and a scalpel, place one dorsal skin at a time hairy-side down onto a thin Petri dish. Start scraping all subcutaneous tissue, including fat from the ventral side of the skin tissue, until it is semi-translucent. Try to remove maximum traces of subcutaneous tissue without tearing the skin (hair follicle part). Also, do not scrape for a long time and avoid skin drying.
   1. Keep scraped skin in PBS solution until all other remaining skins are processed. Then, using a scalpel, slice the skins into 0.5 cm x 1-1.5 cm strips and place the hairy side up onto a sterile Petri dish.
4. Carefully pipette 20 mL of PBS/2x Gentamicin solution mixed with trypsin (0.25%) into a 100 mm x 20 mm plastic Petri dish. Using sterile forceps, transfer the skins and float them hairy-side up on the surface of trypsin solution for 2 h in a 32 °C incubator.
   NOTE: Trypsinization time and temperature are crucial for a good yield of highly culturable primary keratinocytes. Although other methods may provide good yields of viable cells, the culturability of the keratinocytes has been less satisfactory.
   1. During this 2 h incubation time, coat dishes with collagen coating and place them at 37 °C for 1 h (see step 1.1.2). For clonogenic assays, the 60 mm Petri dishes are coated 24 h prior to keratinocyte harvesting to allow the irradiated 3T3 feeder layer to attach to the bottom and spread evenly.
      NOTE: Coating of culture dishes for clonogenic culture is very important for proper attachment, colony formation, and ultimate growth/proliferation of mice epidermal keratinocytes.
5. Epidermal scraping step: Arrange a sterile plastic square Petri dish and create a surface at a 30° incline for proper epidermal scraping with 15 mL of harvesting medium (see Table 1). Carefully remove a floating skin strip from the trypsin solution with curved forceps. Then, with a new scalpel blade, scrape off the epidermis and hairs into the medium.
   1. Do not use excessive force, but use sufficient force to scrape the epidermis or affect cell viability.
   2. During epidermis scraping, keep the blade perpendicular to the skin piece. If the blade is angled toward the motion of the blade, there are more chances of skin tearing. If the blade is angled away from the blade movement, there are more chances of incomplete epidermis removal.
6. Carefully pour the harvesting medium containing scraped epidermal cells into a sterile 60 mL jar (autoclavable and reusable) with a 1.5-inch magnetic stir bar. Rinse the Petri dish with an additional harvesting medium to collect the remaining epidermal cells and bring the final volume to 30 mL. Use a magnetic stirrer and stir at 100 rpm for 20 min at room temperature. This process will remove the trapped epidermal cells from hairs.
7. Place a sterile 70 µm cell strainer on the top of a 50 mL conical tube. The cell strainer fits on top of a 50 mL conical tube.
   1. Take the jar inside a biosafety hood. Remove the stir bar and pour the contents into the strainer filter attached to a 50 mL conical tube. Strain out undesired hair and sheets of stratum corneum.
   2. Press the hairs along with stratum corneum materials within the strainer and manipulate them to release the trapped hair cells. Use 5 mL of harvesting medium to allow remaining trapped cells to release and flow into the tube. Bring the total volume up to 50 mL in a conical tube.
8. Cap the 50 mL conical tube with cell filtrate and centrifuge at 160 x g for 7 min at 4 °C. Next, aspirate the supernatant and add 5 mL of harvesting medium. Finally, resuspend the cell pellet by triturating gently ~20x with a 5 mL pipet.
   1. At this step, keep the cells in an icebox to avoid any aggregation. Next, add 25 mL of harvesting medium and triturate again (20-25x).
   2. To ensure an accurate keratinocyte count at this step, take 1 mL of cell suspension and add 19 mL harvesting medium. Now, the cell dilution is 1:20 in a 50 mL conical tube. If keratinocytes are harvested from a single mouse, adjust the volume of the cell suspension. For example, instead of 30 mL, use 15 mL and make a 1:10 dilution for counting the cells.
9. Pipet 0.5 mL of diluted cells from the 50 mL conical tube (1:20 dilution) and transfer to a 1.5 mL autoclaved microcentrifuge tube. Remove 0.2 mL (200 µL) of cell mix to another 1.5 mL microcentrifuge tube and add an equal volume (200 µL) of 0.4% Trypan blue solution. Gently mix this solution 3x and transfer cells to a hemocytometer for counting nucleated keratinocytes.
   1. Score all dark blue cells positive for Trypan blue as non-viable dead cells, and score small gold and pinkish cells as viable cells. Thus, the final keratinocyte yield per mouse should be around 30 million viable cells.
10. Centrifuge the original 50 mL conical tube (from step 1.8) for 7 min at 160 x g at 4 °C. At this step, resuspend the cells in desired medium and make the required dilution for seeding cells.
    1. For mass culture, seed 2 to 4 million viable gold cells in each 35 mm Petri dish in cell culture medium (KGM-type medium + supplements for monolayer cultures) (see Table 1). For a clonogenic (colony formation) assay, seed 1,000 cells per 60 mm Petri dish on X-ray irradiated Swiss mouse 3T3 feeder layers with collagen-coating and modified Willia's E medium with supplements and serum.
    2. Use a total of 2 to 4 mL medium for 35 mm and 60 mm Petri dishes, respectively. Also, formulate DMEM and Williams E medium procured from ATCC with reduced levels of sodium bicarbonate for use at 5% CO₂.
11. Grow the culture cells (mass or clonogenic) at 32 °C, 95% humidity with 5% CO₂. Change the media one day after initial seeding for mass culture, and then 3x weekly thereafter. However, for clonal assays, the first medium change is 2 days after cell seeding and 3x weekly thereafter.
12. For clonal culture, cultivate cells at two- and four-week intervals. After completion of clonal cultures (2 or 4 weeks), aspirate the medium. Fix the colonies in 10% buffered formalin overnight at room temperature and stain with 0.5% rhodamine B in autoclave water for 1 h. Then, remove the rhodamine B stain with a pipette from the edge of the dishes.
    1. Rinse the dishes in cold autoclaved water until the dishes run clear. Put the dishes inclined on the lid of the dishes and let them dry for final colony counting. Notably, when performing clonogenic culture assay, never pipet <1 mL of cells. If cells are concentrated, serially dilute the cell concentration.

Table 1: Solution and media recipes.