Primary keratinocyte cell culture

Stucky lab Keratinocyte Primary Cell Culture as of Sept 2, 2024

Gather Tools and Reagents:

1. Equipment – all tools/equipment should be sprayed with 70% EtOH before placing in hood. 5-10min UV sterilization will also help reduce chance of contamination.
   1. 2x Forceps – moving coverslips and separating dermis/epidermis after dispase digestion.
   2. 1 mL pipette & pipette tips
   3. 200uL pipette & pipette tips
   4. 2x small petri dishes – labeled HBSS, Trypsin
   5. 1x 5mL centrifuge tube - labeled Dispase 
   6. Tube racks for 15/50mL conical tubes
   7. 4x 15mL conical tubes – labeled HBSS, Trypsin, Keratinocyte, Waste
   8. 1x 50mL conical – labeled EL+ w/ today’s date and your initials.
   9. 1x hemocytometer
   10. 1x 12-well plate
   11. Glass coverslips
2. Reagents
   1. EpiLife (supplemented with HKGS) (4C)
   2. HBSS- (4C)
   3. PBS- (1x) (4C)
   4. Versene – need 750uL (4C)
   5. Fungizone – 100uL aliquot (-20C)
   6. Pen/Strep – 200uL aliquot (you only need 125uL) (-20C)
   7. Dispase – 20mg/mL in petri dish w/ 2mL HBSS (4C)
   8. Trypsin – 2x 100uL aliquots, need 150uL (-20C)
   9. FBS – need 1x 750uL aliquot (-20C)
   10. Trypan Blue

Prepare media and coverslips:

1. Clean hood (70% EtOH) move all equipment into hood (no reagents at this time) and UV sterilize hood+equipment for 5-10mins. Dissection area can be set up at this time.
2. After UV sterilization, obtain reagents – Laminin should be thawed in fridge (~5mins), dispase can be weighed out with the scale, the remaining reagents can be placed in the hood after being sprayed with 70% EtOH.
3. Incubate coverslips with laminin – 2hrs.
   1. Place a single coverslip in each well – make sure you only place 1 in each well.
   2. Dilute laminin aliquot with 500uL PBS-
   3. Pipette 35-50uL onto the center of each coverslip – for your first culture start with 45uL. Adjusting droplet volume can affect cell density.
   4. Place plate in 37C incubator for ~2 hours
4. Prepare EpiLife medium with supplements (EL+) – keep warm in bead bath or 37C incubator.
   1. 50mL of EpiLife medium
   2. 100uL Fungizone
   3. 125uL Pen/Strep
5. Mix up trypsin-EDTA solution in the appropriated labeled 15mL conical: make at least 0.75 mL per sample. 
   1. For 1.5 mL solution: 
      1. 150uL trypsin stock (need 2 aliquots)
      2. 750uL versine
      3. 600uL HbSS-
6. Prepare dispase at a concentration of 20 mg/mL in HBSS-, 2mL per sample (so 40mg in 2mL HBSS-), place this in the 5mL centrifuge tube labeled Dispase.
7. Place 2mL each of PBS and HBSS in the appropriate petri dishes, hold off on adding Trypsin to the labeled petri dish.
8. Place 4mL HBSS in the appropriately labeled 15mL conical.

Culture Protocol:

1. Excise skin sample from 1 paw.
2. Place dissected skin sample in HBSS to wash off procedural chemicals.
3. Once washed, transfer skin sample to dispase solution. Digest at RT for 45mins on shaker plate or in 4C fridge for up to 24 hours.
4. Gently separate the epidermis from the dermis with the 2 pairs of forceps (epidermis is thinner and clear, papery; dermis is thicker, more of an opaque white/pink and gloppy). Discard dermis.
5. Wash epidermis in HBSS (previously filled petri dish) and then place in currently empty trypsin dish.
6. Pipette 750uL trypsin solution on top of epidermis, making sure to completely cover the sample. Incubate for 27 mins at RT (25-30min). 
7. After incubation, add FBS aliquot to sample (neutralize trypsin), and rub the epidermis on the base of the petri dish to separate cells from the epidermal sheet. This can be done with the tip of a 1mL pipette tip. 5-10 mins of rubbing should produce enough cells. Watch for clouding of media.
8. Add 2mL HBSS to petri dish, continue to rub epidermis if more cells are needed. Pipette media into 15mL conical (labeled keratinocyte) and then wash petri dish with an additional 2mL HBSS to collect additional cells.
9. Centrifuge collected cells for 6min @ 200g.
10. During spin down step remove 12-well plate from incubator and remove laminin from coverslips. Try to avoid touching the coverslip with pipette tip. 
11. Wash laminin droplet with 35uL EL+. Remove EL+ and replace plate cover. Try to avoid letting the laminin dry on the coverslips. It may be best to reserve this step for after the cells have been counted.
12. Observe pellet and discard supernatant. Resuspend the pellet in .5 – 1mL EL+ complete media.
    1. Gently titrate to break up larger clumps – 20x times with 1mL pipette, 10x times with 200uL pipette
13. Count cells w/ hemocytometer (desired concentration is ~78*10^4 cells/mL)
    1. Mix 20uL of cell suspension with 20uL trypan blue. Add 10uL to one side of hemocytometer. Count healthy cells (clear, circular) and average 4 corner quadrants.
    2. Calculations: Average cell count * 2 (dilution factor w/ trypan)/78 (desired conc) * initial volume of EL+ added during resuspension = final volume needed to reach desired concentration
    3. Example (concentration is x*10^4):
       1. Average – 322/4= 80.5
       2. Account for dilution – 80.5*2= 161
       3. Divide by desired conc – 161/78= 2.06
       4. Multiple by initial media volume – 2.06*500uL= 1032uL
       5. You would need to add 532uL of EL+ to reach the desired cell density of 78*10^4
14. Plate cells on laminin. Make sure to resuspend cells if they appear to have settled in solution. 45uL on the center of each laminin droplet should be enough.
15. Incubate cells in 37C incubator for 2hrs. Feed w/ 1mL of EL+ by gently pipetting onto the side of the well.
16. Allow cells to plate down in incubator for 48 hours. Change media – Remove 700uL of media from each well, feed with an additional 1mL EL+.
17. Cells can be used at 72hrs (3 days post culture)