Prepare Supplemented Growth Medium
1. Remove 50 mL of volume from the bottle of MEM basal medium and discard.
2. Add 50 mL of fetal bovine serum (FBS) to the remaining volume of MEM.
3. Add 5 mL of 100X penicillin/streptomycin to the medium.
4. Add 5 mL of 100X MEM non-essential amino acids (NEAA) to the medium.
5. Add 5 mL of 100mM sodium pyruvate to the medium.
6. Mix thoroughly and label with the date and additives.
Collagen Coating of Tissue Culture Plates
IMR90 cells should be grown on dishes/plates/inserts that have been coated with 50 mg/mL bovine collagen I solution according to the separate method “Collagen Coating for Tissue Culture”.
Thawing Cryopreserved Cells
1. Warm medium and prepare materials prior to obtaining a vial of cells from cryostorage.
a. This reduces the amount of time cells are thawed in the presence of DMSO.
2. Remove vial from liquid nitrogen (wearing eye protection) and thaw in 37 °C water bath (1-2 min).
a. It is important that cells are thawed quickly to prevent damage and reduced viability.
3. Add 24 mL of growth medium to a 50 mL conical tube then transfer the thawed cells from the cryovial into the growth medium and gently mix by inversion.
4. Pellet cells via centrifugation at 1,000 x g for 4 minutes at room temperature.
5. Carefully aspirate the supernatant.
6. Gently resuspend the cells in 12 mL of pre-warmed growth medium, add 13 mL of additional growth medium, and transfer into a collagen-coated 15 cm dish.
7. Incubate in a humidified 37°C incubator with 5% CO2 and 20% O2 overnight.
8. Check cells the following day for attachment to the dish. Replace medium to remove dead cells (rounded, unattached cells). It is preferable to culture thawed cells for at least three passages before using in experiments.
a. NOTE: Cells may require being cultured until the fourth day after thawing prior to being ready to be passaged for the first time.
Cells should be split three days after being plated at 4.0 x 104 cells/mL
· Lower APD IMR90 cells grow very quickly and should be plated at 3.5 x 104 cells/mL to start and then adjusted up to 4.0 x 104 cells/mL accordingly as their growth rate slows down.
1. Pre-warm growth medium and PBS in a 37°C water bath and a trypsin aliquot at room temperature (~30 minutes).
2. Aspirate the cell culture medium.
3. Rinse cells with pre-warmed PBS and swirl gently.
a. 20 mL for 15 cm dishes
b. 10 mL for 10 cm dishes
4. Aspirate the PBS wash.
5. Add trypsin and rotate to spread evenly across the plate.
a. 1.5 mL for 15 cm dish
b. 750 μL for 10 cm dish
6. Place in 37°C incubator for 4 minutes, making sure not to stack dishes on top of each other. Following incubation, use the palm of your hand to hit the side of the dish to facilitate detachment. If cells are still attached, incubate for another 2 minutes. Incubate and tap dish until >95% of the cells are detached. Check detachment with microscope.
7. Add 12 mL medium and gently wash the surface of the dish with a pipette and triturate once to break up any large cell clumps. Transfer to a 50 mL conical tube.
8. Wash the dish with an additional 12 mL of medium, collect, triturate, and add to the cell suspension from Step #7.
9. Centrifuge at 1,000 x g for 4 minutes to pellet cells.
10. Aspirate the supernatant and resuspend in 12 mL of medium. Triturate at least four times to thoroughly break up the majority of clumped cells. Add additional growth medium to dilute the cell suspension if desired. Typically, medium is added such that there is a total volume of 12 mL per 2-3 15 cm plates collected.
11. Prepare a 1:1 dilution in Trypan Blue (50 μL Trypan Blue + 50 μL cell suspension) in an Eppendorf tube.
12. Count cells that exclude Trypan Blue on a hemocytometer (do not use an automated cell counter as these cells display variability in size and are not accurately quantified by automated methods).
a. NOTE: Blue staining of cells indicates a damaged membrane. Blue-stained cells are considered to be non-viable.
b. Record the number of cells obtained per plate.
c. Calculate the number of population doublings that occurred during the last passage (include the indicated values in the associated worksheet):
([(3.32(log(cells collected per plate)-log(cells plated per plate)) + previous APD])
13. Dilute cells to 4.0 x 104 cells/mL (or 5.0 x 104 cells/mL for IMR90-UNC cells) in growth medium in a 50 mL tube or larger sterile bottle to the total volume that will be used for plating (10 mL for each 10 cm plate and 25 mL for each 15 cm plate, making 2-5 mL extra).
14. Mix thoroughly by inversion and dispense the diluted cell suspension into each dish.
15. Incubate in a humidified incubator at 37°C with 5% CO2 and 20% O2. Cells will be ready to split three days after plating.
a. NOTE: Plating cells at a determined cell density will result in more predictable sub-culture and more reproducible performance in subsequent experimental assays.
16. Place bottles of medium and PBS in 5% CO2 incubator with lids partially unscrewed for 15 minutes to adjust pH before returning to 4 °C fridge to store.
Preparing Cells for Cryopreservation
1. Prepare freezing medium.
a. 50% FBS, 40% growth medium, 10% DMSO.
b. Filter through a 0.2 μm pore syringe filter.
2. Follow sub-culturing protocol through Step #10.
3. Determine the number of cells present and pellet cells by centrifugation at 1,000 x g for 4 minutes at room temperature.
4. Resuspend the cell pellet at a density of 1.25 x 106 cells/mL in freezing medium.
5. Aliquot into cryovials (1 mL per vial).
6. Put vials in a Mr. Frosty controlled freezing container (filled to indicated line with isopropyl alcohol) and place in a -80°C freezer overnight.
a. The chamber regulates cooling rate at about 1°C per minute.
b. NOTE: Isopropanol should be changed after every three freezing cycles.
c. NOTE: Do not leave vials at -80°C longer than overnight as it will impact viability after thawing.
7. Put the frozen vials in liquid nitrogen storage.