General mammalian cell culture

This work is licensed under CC BY-SA 4.0

This SOP contains multiple parts: frozen cell recovery, passaging, growing on glass coverslips, and transfection with Lipofectamine 2000.

Procedures must be performed in sterile environment (laminar flow hood). All reagents and plastics, except single cell plates, must be sterilized before bringing into the flow hood by spraying with 70% ethanol and wiping dry. Always attach sterile pipette tips to the aspirator to prevent cross-contamination.

Reagent

Culture media: DMEM with 10% FBS, 1% Pen-Strep, 1% Glutamax. Once made stable at 4C for at least one month.

Thawing frozen cells

Adapted from general cell culture protocol from Fisher Scientific.

1. Remove the cryovial containing the frozen cells (500uL) from liquid nitrogen storage and immediately place it into a 37°C water bath. You may put cells with dry ice for short term transport before thaw.
2. Aliquot 1.5mL of culture media into each well of a 6-well plate³. Prewarm the plate in incubator.
3. Quickly thaw the cells (< 1 minute) by gently swirling the vial in the 37°C water bath until there is still some ice in the vial.
4. Transfer the vial it into a laminar flow hood. Before opening, wipe the outside of the vial with 70% ethanol.
5. Add the thawed cells directly into wells with prewarmed media.
6. Put the plate back into incubator. Next day exchange into new media to remove dead cells.
7. Grow until cells reaches near confluency. Then passage into 10cm plate for maintenance (use all cells, split into one 10cm plate, see below).

Passaging/splitting cells

HEK293 splitting follows 1:10 ratio (one confluent dish split into 10 new dishes of the same size). Use 1:3 ratio for U2OS and SH-SY5Y cells as their doubling time is longer. Volumes below are for 10cm plates.

1. Prewarm whole DPBS bottle and an aliquot of trypsin-EDTA (1mL per plate) to 37C.
2. Add (10-X)mL of media to each new 10cm plate. Mark accordingly. Put plates in 37C incubator until use. X depends on the split ratio (3 for U2OS/SH-SY5Y and 1 for HEK293).
3. Aliquot media into a conical tube, 9mL per plate. Prewarm the media to 37C.
4. Take out confluent plate from incubator. Aspirate media from the plate to split. Keep covered.
5. Add 10mL of DPBS to the plate. Add from the side (not directly onto cells).
6. Swirl gently and aspirate all DPBS.
7. Use P1000 to add 1mL of trypsin-EDTA. Add directly onto cells.
8. Swirl plate to make sure even distribution of trypsin.
9. Put plate in 37C incubator. Dissociate for 3min (U2OS/SH-SY5Y) or 1min (HEK293)⁴.
10. Add 9mL of prewarmed media directly onto plate. Pipette to triturate. Make sure that cells are dissociated properly under light (no apparent clusters)⁵.
11. Transfer XmL of cells to the new, prewarmed 10cm plates with media. X depends on the split ratio (see Step#2)⁶.
12. Place new plates back into 37C incbator. Put reagents back.

Growing cells on glass coverslips

Typical usage: immunostaining, in situ hybridization, etc.

Use cell-culture grade water (autoclaved milli-Q or ultrapure) for preparing stock and working solution.

Preparing poly-D-lysine solution and sterile coverslips

1. Prepare stocks as 1mg/mL in water. Store as 500uL aliquots and store in -20C.
2. Prepare 70% ethanol and pour into a 10cm dish. Incubate coverslips in ethanol for initial sterilization wash (room temperature, 5mins for each side up).
3. Take out coverslips from ethanol and tap dry with paper towel.
4. Lift one side of coverslip to allow air dry. Turn on UV light of the flow hood for 15min.
5. Put sterile coverslips into a 10cm dish for later use.

Coating coverslips

1. Take out 1mg/mL PDL stock solution and dilute to 50ug/mL with water to get working solution.
2. Incubate coverslips with working solution in the flow hood for at least one hour (overnight incubation for best results).
3. Remove the working solution. Wash the coverslip with water thoroughly⁷.
4. Tap dry with paper towel and then air dry. Coated coverslips are ready to use or store at 4C sterile for up to two weeks.

Transfection with Lipofectamine 2000

Refer to Fisher complete protocol for more details. Volumes below are for 24-well plates. DNA must be endotoxin free.

1. Cells should be ~80% confluent at transfection. Swap into new media (0.5mL) for each well.
2. Warm optiMEM medium to room temperature (or 37C).
3. Dilute 1ug of plasmid DNA in 50uL of optiMEM medium for each well.
4. Dilute 1uL of lipofectamine 2000 in 50uL of optiMEM medium for each well. Pipette to mix and incubate in hood for 5min.
5. Add diluted plasmid to diluted lipofectamine. Pipette rigorously to introduce bubbles. Incubate in hood for 20min.
6. Add 100uL of mix to each well. Rock for even distribution.
7. Incubate overnight for transfection. Next morning swap into fresh media.
8. Expression should be visible typically 16-24hr after adding transfection mix.

Per well of 24-well plate: 100uL OM/LF mix with 1ug plasmid, add into cell with 0.5mL media.