Culture of Mouse Keratinocytes




1. Place a small amount of dry ice in the base of the animal anaesthetizing chamber. Stuff the hole and cover the base with gauze and lay the newborn mice on top. Cover with the lid and leave for 30mins.


2. Assemble the following in the cell culture hood:

35 ml Betadine diluted 1:10 in sterile dH20

2x 35mls sterile dH20

2x 35mls 70% ethanol

3 pairs forceps, scissors, scalpel handles and blades in a beaker or tube of 70% ethanol

6 well plates for skin

24 well plate for the tails

sterile square culture dishes for the dissection

Thaw Dispase (use Becton Dickinson from fisher diluted 1:4 in Hanks; need 2mls/skin)


Day 1


1. Distribute 1 ml of Dispase (25 U/ml) in each well of a 6-well (35 mm) plate

2. Cut the heads off the mice and place them in Betadine solution (6 max) using long forceps. Wash for 2 min. Maintain sterility! Do not touch the mice, tubes or caps after this point.

3. Sequentially transfer the mice into the first and second tubes of sterile dH20, then the first and second tubes of 70% EthOH washing for 2min. at each step.

4. Transfer the mice into a square culture dish. Process each individual as follows: cut off the tail and place in the 24 well plate for genotyping, cut off the arms and legs. Open the skin along the length of the back. Hold the skin along its length with the grip forceps and roll the body away from it with the scalpel. Do not puncture the body wall and avoid contact with the intestine which could bring infections. Flatten the skin dermis side down on the lid of the culture dish.

5. Float the skin dermis down in Dispase. Incubate 0/N (minimum 2-3 hr) at 4°C

6. Immediately perform genotyping on the tails.

Culture of Mouse Keratinocytes


Day 2

1. Assemble the following in the cell culture hood:

One 50ml and one 15ml labeled tube/skin

Thaw 10X Trypsin and dilute with Hanks

Thaw DNAse and sterilize

Gauze pads

Forceps and scalpels in 70% ethanol

DMEM and Keratinocyte medium

60 mm cell culture dishes

1, 5, 10 ml sterile pipettes

Sterile stuffed Pasteur pipettes

2. Separate epidermis from dermis using watchmaker’s forceps and place in the lid of the 6 well plate.

3. Add 0.5 ml of 0.05% Trypsin to the epidermis and mince using sterile surgical blades

4. Add 1.5 ml of Trypsin to the minced epidermis and transfer into a 15 ml Falcon tube and incubate at 37°C (water bath) for 10-15 min.

5. Add one drop DNAse (stock 25 mg/ml in TE filtered through 2 Ám filter) and mechanically release the epidermal cells by nine vigorous pipettings using a Pasteur pipette. Avoid bubbles.

6. Filter the solution into a 50 ml Falcon tube through a sterile gauze and immediately wash through with 10ml of complete DMEM to inactivate the Trypsin.

7. Centrifuge at 1000 rpm for 5-10 min

8. Remove the DMEM and resuspend the pellet of cells by gentle pipetting with 3 ml of low calcium KGM medium (mix KGM: No Ca KGM 1:2 and add BPE) and plate out in a 60 mm culture dish



Day 3

Observe the culture but do nothing; they always look terrible and the cells need 24 hrs undisturbed to attach and spread.


Day 4

Rinse with Hanks, add fresh low calcium KGM and assess the degree of confluency. When cells reach 50 % confluency you may want to switch to DMEM to permit junction formation.