Culturing mammalian cells

This was written about HEK293T cells.

Materials

Reagents

• Complete media: DMEM (Dulbecco’s Modified Eagle Medium) with 10% v/v fetal bovine serum (FBS)
• Optional: 1x penicillin/streptomycin (buy as 100x stock; prevents bacterial contamination)
• OptiMEM (for transfections; can use serum-free DMEM as a cheaper alternative)
• Dulbecco’s PBS without calcium or magnesium
• Versene (PBS with EDTA; use to detach cells without proteolyzing cell surface receptors)
• Semi-optional: Trypsin/EDTA solution (can buy 0.05% or 0.25% trypsin, any will work for HEK cells and make detaching cells faster than versene)
• Tissue-culture grade DMSO (for freezing cells)
• Trypan blue
• 70% ethanol, bleach

Equipment

• TC Hood, incubator
• Cell counter – e.g. hemocytometer
• Microscope
• Flasks: T75 flasks are most common, similar flasks range from T25-T225 (number indicates culture area). TC-treated petri dishes are cheaper but prone to accidents
• Plates: 96-, 48-, 24-, 12-, and 6-well plates depending on assays
• Falcon tubes
• Cell freezing container: I like the styrofoam kind, can also use an isopropanol Mr. Frosty
• -80 °C freezer, liquid nitrogen storage tank
• Water bath (37-40 °C)
• Pipettes for aspiration (sterilized glass or disposable plastic)
• Vacuum trap
• Filter pipette tips

Some transfection reagents

• Polyethylene imine (cheapest, dirtiest)
• Lipofectamine (relatively cheap, occasionally toxic)
• Fugene (slightly more expensive but non-toxic)

General Tissue Culture Practices

1. Keep hood tidy and free of waste to deter contamination. Do not block vents or air flow will be disrupted.
2. Spray and wipe hood down with 70% ethanol before and after each use
3. Everything that enters or leaves hood should be sprayed with 70% ethanol including pipettes, tubes, flasks, and your gloves
4. Use a dedicated lab coat for tissue culture separate from E. coli work
5. Never open flasks outside of the hood
6. Human cell lines can propagate human pathogens
7. Check cells regularly for signs of contamination (yellow and/or foul-smelling media; rounded, slow-growing or otherwise unhealthy cells)
8. Some journals will ask for checks for mycoplasma, which commonly infect cultures (even when you use pen/strep)  

Thawing Cells

Mammalian cells are not healthy below 37 °C or with DMSO so it’s best to thaw them quickly

1. Before starting: warm media to ~37 °C in water bath (takes around 1 hr)
2. Remove vial of cells from liquid nitrogen
3. Thaw quickly in 37 °C water bath (not in your hand)
4. Dilute cells in ~5 ml fresh media in a falcon tube
5. Centrifuge at 400 x g for 3 minutes
6. Aspirate media carefully without disturbing cell pellet
7. Resuspend gently in ~5 ml fresh media
8. Add to desired flask along with more media to correct volume (see below). Typically cells are frozen at 3-5 million per ml, to be plated in a T75 with ~13 ml media.
9. Gently rock flask to distribute cells evenly across surface
10. Record passage number and place in incubator overnight
11. Check morphology the next day: cells should have attached to plate overnight and spread out (not rounded). If dead cells remain in solution you can gently aspirate media and add fresh  

Splitting cells

HEK293T usually double in slightly under 24 hours and must be split every 2-3 days

1. Check confluence of flask under the microscope (occupancy of cells in area of field of view)
2. Also check media color (red = healthy, orange = ready to split, yellow = unhealthy or contaminated)
3. Should split before cells reach 100% confluency
4. Before starting: warm media, versene or trypsin, PBS to 37 °C
5. Aspirate media using a sterile pipette
6. Gently wash adherent cells with PBS to rinse media away (FBS contains trypsin inhibitors) a. Do not pipette directly on cell layer; pipette on side/back of flask
7. Gently rock plate, aspirate PBS using a sterile pipette
8. Add versene or trypsin according to chart below, gently rock plate to distribute liquid
9. Place in incubator for ~3 minutes
10. Check cell morphology under microscope a. detaching adherent cells should round up and float off.
11. Add an equal volume of fresh media. Rinse the bottom of plate to completely detach cells
12. Add detached cells to a falcon tube and centrifuge at 400 x g for 3 minutes
13. Gently resuspend cells in fresh media
14. Split cells as desired. Typical HEK293T splits are ~1:6 every 2-3 days. For example, resuspend cells from a confluent T75 in 6 ml of fresh media, add 1 ml of suspension to a new T75 with 12 ml fresh media. Adjust volumes and math for scaling up or down in flask size
15. Gently rock the flask to distribute cells across surface
16. Record passage number and date
17. Place in incubator. Check morphology and confluence daily

These routine splits are for maintaining cells. For experiments, cells should be counted and viability checked with Trypan blue for reproducibility.  

Cell Transfection

Using PEI

Materials

1. PEI solution
2. 150 mM NaCl solution
3. Seeded cells in a plate or flask at <80% confluency
4. plasmids to be transfected

Procedure

1. On the bench, prepare the DNA in an Eppendorf tube for each transfection. Aim for a total of 750-800 ng DNA for each well in a 24-well plate, but adjust for different surface areas for other size wells/plates. Mastermixes are fine if one transfection corresponds to multiple wells. Keep an extra tube to mix PEI and NaCl solution later.
2. In the TC room, turn on the biohood, spray down the inside with ETOH, and then spray down your tube rack, pipette tip boxes, and Falcon tubes with the PEI and 150 mM NaCl solution and put those inside also.
3. Adjust the volume of all the transfection tubes with NaCl solution such that you have 13 ul per well (for a 24-well plate - it will be 33 ul per well for a 6-well plate).
4. In the extra tube, mix 1:12 PEI/NaCl for a total volume equal to the number of wells * 12 ul (for a 24-well plate).
5. Vortex tubes and wait 10 minutes, leaving tubes at RT.
6. Add a volume of the PEI/NaCl solution to each tube equal to its DNA solution volume.
7. Vortex tubes, spray down, and reinsert in the hood.
8. Open up plate or flask to expose the cells and media. For each transfection, pipette the DNA/PEI mixture, dispense it dropwise around the well (20-25 ul per well for a 24 well plate, 65 ul for a 6 well plate).
9. After dispensing all the DNA, shake the plate/flask on the biohood internal surface to stir the mixture into the media and place it back in the incubator.
10. Spray down the hood surface with ETOH, put all the materials back, and turn off the hood light and vent.

Freezing cells

Cells should not be maintained indefinitely (usually less than 20 passages or so). Store low-passage stocks in liquid nitrogen and thaw new vials routinely. Cells usually should be frozen slowly in 5-10% DMSO in complete media at ~3-5 million cells/ml (see ATCC page for cell-specific instructions)

1. Detach cells with versene or trypsin as above. Pellet in centrifuge
2. A T75 usually has enough cells for two vials and a split, so resuspend in ~2.5 ml fresh media
3. Count cells and note viability using hemocytometer and trypan blue stain
4. Dilute or re-pellet as needed to get 3-5 million cells per ml
5. If also splitting, take cells as needed
6. Add DMSO to remaining cells for a final concentration of 10% v/v
7. Gently mix and add to cryo-vials. Record passage number and cell count on the vial.
8. Cells should be frozen slowly with DMSO to prevent ice from swelling and bursting cells
9. Put vials in room temperature cell freezer and seal
10. Place cell freezer in -80°C freezer for several hours to overnight to allow vials to freeze.
11. Transfer to liquid nitrogen for long-term storage.  

Useful information for various sizes of cell culture dishes and flasks

There are various sizes of dishes and flasks used for cell culture. Some useful numbers such as surface area and volumes of dissociation solutions are given below for various size culture vessels.