Basic Tissue Culture Technique

Boyd Lab

Preface

All human tissue culture work should be carried out wearing gloves and long-sleeved gown in a biosafety cabinet contained in an appropriate area. Gloves and consumable tissue culture products must be disposed in a biohazard bin.

All solutions below are assumed to be sterile unless stated otherwise.

Media

The priniciple media type used in the Boyd Laboratory is RMPI-1640, purchased from CSL in powder form and made up in 20-40 L batches to pH7.0, stored at 4•C up to 2 years. Most major suppliers of culture reagents and media also stock RPMI. Other media types are sometimes used for specific applications for example SFM (Life Tech) for growing hybridomas for antibody purification; DMEM for some mouse lines; Iscoves for chicken cell culture.

RPMI is a useful universal media type since it can be used for human cellular work and is easily adjusted for mouse work by the addition of 2g NaCl/10L (290 mOsm to 325 mOsm)

Media Supplements

• Medium is routinely supplemented before use in culture by adding 25 or 50 mL of FBS (fetal bovine serum, CSL) ie 5% or 10% depending on culture needs (can be kept as a frozen stock in aliquots).
• Glutamine is added to 2mM since this essential amino acid is not stable in aqueous solutions. It is usually added as 5mL of a 200mM solution (5g in 170mL water). Glutamax™ is now used which does not break down (both can be kept as a frozen stock in aliquots)
• Other supplements are used for specific culture situations such as for transfected cell lines or for Fetal Thymic Organ Culture (FTOC) See methods

Thawing cell lines from liquid nitrogen stocks

• Aspirate 8-9mL of media into a 10 mL tube and place this on ice or in a refrigerator for 15 min before thawing cells -the preservative DMSO (dimethyl sulfoxide, BDH/Merck) if allowed to warm around cells may enter them and become toxic.
• Bring vial(s) from nitrogen store immediately to a 37°C water bath. If thawing a number of vials store them in dry ice prior to thawing and thaw only one or two vials at a time.
• Swirl a vial or two in the bath taking care not to immerse completely, to prevent any bath water from entering the sealed cap.
• When there is only a small amount of ice left in the tube ie still cold, take the vial(s) to a sterile cabinet/hood and coat with 70% ethanol to wash off bath water and kill any bacteria on the outside surfaces.
• Aspirate the contents of the vial(s) (usually 1-2 mL) into the 8-9 mL cold media and centrifuge immediately 400g, 5 min, 4°C.
• Wash cells in fresh media by aspirating the supernatant, resuspending the cells in fresh media and centrifuging again.
• Finally resuspend the cells in 1 mL and count then plate.

Counting cells

We use ethidium bromide/acridine orange (EtBr/AcOr) inclusion/exclusion as our principle method (see here for recipe) but others may also be valid such as the use of a Coulter™ counter or trypan blue.

• The cells suspension may need to be diluted to count a reasonable number -1/10 to 1/50 is often sufficient.
• Take 10 µL of the diluted cells and add this to 10µL of EtBr/AcOr and mix gently. Add 10µL to the edge of a haemocytometer coverslip mounted on the haemocytometer chamber. Repeat on the other side.
• Count at least 100 cells for accuracy and calculate the cell number / mL.

The main centre grid is 1mm x 1mm in area and 0.1 mm deep i.e. 10 ^-4 mL

eg Cell count x 2 x dilution (ie x 10 or x 50) x 10 ⁴ = cells / mL

Plating cells

This term refers to decanting an appropriate cell suspension into tissue culture vessels which may be of a tray type of various well numbers, flasks of varying volumes or even petri dishes (where the term orginates)

Freezing cells down

Cells to be frozen are harvested from the culture vessel and centrifuged as before, then resupsended in cold 10% DMSO in FBS(can be kept as a frozen stock in aliquots)

• For quantity, as a rule of thumb:
  • 1 vial (1 mL of 10% DMSO in FBS) per small flask of cells -15 cm²
  • 3 vials for a medium flask -75 cm²
  • 6 vials for a large flask -150 cm²
• The vials are quickly transferred to a 'Mr Frosty™' or if one is not available, a wadd of cotton wool and placed at -70°C until frozen (wait at least 1-2 hours to O/N)
• Then transfer the vials to liquid nitrogen storage containers.

Growth techniques

You must be taught to recognise healthy from non healthly cells by someone experienced and learn when cells should be transferred/passaged.

Adherent vs Suspension culture

Cells that grow in suspension vs those that adhere to the tissue culture vessel are handled differently.

• Suspension cultures may be divided by simple pipetting/resuspension.
  
  
• Adherent cells actively bind to the plastic ware, usually in a reversible fashion. They may be split/divided/transferred by the use of EDTA alone or solution containing EDTA and trypsin

Method

• Remove/aspirate the media from the cell culture vessel
• Wash the vessel/cells with serum free media or PBS to remove traces of FBS containing media
• Add enough trypsin/EDTA (Life Tech pre-made stock or 0.1% trypsin / 2mM EDTA in balanced salt solution, no Mg or Ca) solution to cover the base surface area and transfer the vessel to 37°C incubator for 5 minutes.
• Using the trypsin/EDTA solution, resuspend and wash the vessel floor the aspirate the cells to a tube containing media for washing by centrifugation.
• After washing the cells may be counted or divided appropriately for further culture, application ie staining procedure, lysis etc, or resuspended in 10% DMSO/FBS for freezing (see above)

Expansion

Cells resuspend as above may be divided in two for expansion, usually when 70-90% confluency (=100% surface area of vessel). They may be able to take a higher dilution ie 1/10 or more if hardy and in exponential phase. Generally this is done by taking one well of a given size and dividing the cells between 2 wells of the same size and keeping the volume the same eg:

• Resuspend a 96 well containing 200 µL cells and transfer 100 µL of cells each into two new wells, then add 100 µL fresh media to make a final volume of 200 µL each.
• Depending on cell growth, 2 to 4 wells from a 24 well tray may be transferred to a small flask (15 cm²)
• 2-3 small flasks to a medium flask (75 cm²)
• 1-2 medium flasks to a large flask (150 cm²)

Maintenance Culture

To keep stable cells in culture a convenient method is to take the cultures to a 24 well tray and serially dilute the cells across the row ie 1 mL + 1 mL dilution then add 1 mL back to the finished dilutions to make 2 mL final volume in each well. One row is usually enough with most cells to last a maximum of 1 week before repeating the process. Assay the cells regularly to maintain their specificity and do not do this routinely, rather if there is a specific reason.

Use of antibiotics

Antibiotics may be necessary for some cell lines to propagate under selective pressure, however for routine culture of most cell types, the use of antibiotics to prevent infection must be discouraged.

Reasons for not using antibiotics

• Antibiotics may cause the delay of onset of infection until it is too late.
• An infection in the presence of anitbiotics is not uncommon and is obviously then of resistant microorganism strain. This has the risk of entering the environment.
• The use of antibiotics supports poor sterile technique

Disposal of infected media/cells

• If a media bottle/supply has become contaminated without antibiotics present it may be disposed of in the water supply with large volume dilution.
• If a media bottle/supply has become contaminated with antibiotics present it must be disposed of by first killing the organisms with sodium hypochlorite (BDH) at 40 ppm from concentrated stock for at least 1 hour. The bottle may then be emptied into the water supply with large volume dilution with tap water.
• If a culture container i.e. a flask or plate etc has become contaminated with or without antibiotics for human or mouse work present, it should be disposed of in a biohazard bin.