Primary mouse keratinocyte isolation and culture

Keratinocyte Isolation and Culture Protocol

        Modified from LifeTechnologies and CellNTech protocols with alterations (contributions from 

trial-and-error and Zela Lazarova)

Updated: 1/20/2015 - FM

Phase 1: 

1. Prepare EpiLife medium with supplements (EL+) in sterile hood
   1. To make 50 mL of EL+ with 60 microMolar calcium:
      1. 50 mL of EpiLife medium already supplemented with HKGS (Human Keratinocyte Growth Medium – both from LifeTechnologies) can just pour it
         1. Already contains EGF, L-glutamine, pituitary extract
      2. 100 uL Fungizone (Gibco, 250 ug/mL amphotericin B)
      3. 125 uL Pen/Strep (Gibco, same aliquots used for culture)
2. Take out EL+ so it can warm up to room temp.
3. Sterilize dissection tools, clean scope and area, place down sterile gauze. 
4. Get 2 sterile 35mm culture dishes per sample. Add 2 mL sterile PBS to one.
5. Prepare dispase at a concentration of 10 mg/mL in the EL+ media (can also use HbSS- if desired). This concentration is sufficient, but others have been used successfully also. 
   1. For 2 samples, about 2 mL per sample – make 4 mL (40 mg dispase) – mix with HBSS-
   2. Add ~2 mL dispase solution to the remaining culture dishes.
6. Sacrifice mouse, spray paw in ethanol, wipe with clean Kim wipe. Repeat if desired.
7. Dissect (glabrous) skin from any paws as desired. Place tissue in the PBS. Wash the tissue by agitating with forceps in PBS. Then transfer to new culture dishes and add complete media. Can again all be in the same dish if they’ll be separated at the dispase step.
8. Move samples to new culture dishes with dispase solution and label dishes. Shake samples in dispase at room temperature for ~45 minutes (more if thick or old or Nair’d), and protect from light with paper towel.
   1. Place skin in dispase solution epidermis side up 
   2. Alternative: Move samples to 4C refrigerator to incubate for multiple hours 10-15 recommended, but can be as short as 5-6 hrs. 
9. Obtain laminin aliquots to thaw. Sterilize glass coverslips or 12-well plate. In hood, dilute with 500 uL PBS. Plate 42-50 uL of laminin where desired, or 300 uL for entire well bottom. Incubate for 2 hours.

Phase 2:

1. Clean laminar flow hood and bring tissue samples over with 2 sterile forceps. Transfer the samples to a new dish with 2 mL sterile PBS or HBSS-. (wash)
2. Gently separate the epidermis from the dermis (epidermis is thinner and clear, papery; dermis is thicker, more of an opaque white/pink and gloppy). Discard dermis. 
3. Mix up trypsin-EDTA solution: make at least 0.75 mL (minimum) per sample. 
   1. Obtain trypsin (stock aliquots are 0.5%), EDTA (versene, 1x, 50% of total volume) and HbSS-. We want 50% EDTA. 
   2. For 1.5 mL solution: 150 uL trypsin stock (need 2 aliquots), 750 uL versene, and 600 uL HbSS-. Mix.
   3. When added to dish keep in “bubble” do not spread solution out!
4.  In a sterile 35 mm culture dish, place 750 uL of the trypsin-EDTA solution. Place one sample’s epidermis portion in the trypsin, floating with the basal keratinocytes facing the solution. – Turn hood light off but NO UV
5. Incubate in trypsin for 27 min at room temperature (25-30 min).
6. Once laminin has incubated for 2 hrs, bring to hood, suck off excess, and let dry in the back (open). 
7. After incubation, add 1 mL HbSS to the trypsin and epidermis. Rub the epidermis on the base of the petri dish to separate cells from the epidermal sheets, watching as the medium becomes more turbid. 
8. Transfer the cells and liquid to a 15 mL centrifuge tube. Wash the original petri dish with 1-2 mL FBS (fetal bovine serum) to neutralize trypsin. Transfer to the centrifuge tube. Add another 1-2 mL of HbSS to the epidermal sheets and continue rubbing the epidermis on the base of the plate. Transfer these also.  à Typical total volume for one paw – 6mL
9. Centrifuge the collected cells for 6 min at 200g (~setting 4, slightly higher).
10. Aspirate and discard supernatant.
    1. Wash with HBSS- as needed (if you still see FBS yellowish color wash again)
    2.  Resuspend the pellet in 1-2 mL of the EL complete medium.  With two paws and good dissociation, 2 mL will work best.
    3. Pipet up and down to break up the pellet
11. Count the cells 
    1. Mix 20 uL of cells + 20 uL Trypan Blue. Add 10 uL to one side of a hemacytometer. Count healthy cells using standard procedure (average number of cells per large corner square * 2(dilution factor with TB) * 10⁴ = number of cells per mL).
12. Plate the cells on laminin. We optimally want 3.5 x10^4 per cm2 (protocol recommendations vary, however). 
    1. Plate sizes: 
       1. A 35x10 mm dish = 8.5 cm2. Want ~30x10⁴ viable cells per dish.
       2. A droplet is about 1 cm2. Want 3.5x10⁴ cells per dish. This is a high estimate.
       3. A 12 well plate is 22.5 mm dish = 3.8 cm2. Want 13.3x10⁴ cells per dish.
    2. Recommended plating volume.
       1. 35x10mm dish = 1 mL
       2. Droplet or cover slip = 45 uL (can be more)
       3. 12 well plate = 172 uL
    3. Dilute the cells to ~78 x10⁴ cells per mL. 
    4. Plate the cells.

Phase 3:

1. Feed the cells after 15 min to 2 hr (1 mL for 12 well plates, 1.5 mL for 35 mm dishes). If doing so earlier, pipette very carefully. Keratinocytes won’t be fully stuck down for quite some time. 
2. Leave the cells alone for 48+ hours before changing media.
   1. When changing the media, pull off 700 uL off and then add 1 mL EL+

Category