Mammalian cell tissue culture techniques protocol

General details of cell culturing and sub-culturing

The following is a general guideline for culturing of cell lines. All cell culture must be undertaken in microbiological safety cabinet using aseptic technique to ensure sterility.

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Contents

1. Preparing an aseptic environment
2. Preparation of cell growth medium
3. Creating the correct cell culture environment
4. Checking cells
5. Sub-culturing
6. Adherent subculture protocol 
7. Sub-culturing loosely attached cell lines requiring cell scraping for sub-culture
8. Sub-culturing attached cell lines requiring trypsin
9. Sub-culturing of suspension cell lines
10. Changing media
11. Passage number

​1) Preparing an aseptic environment

Hood regulations

• (a) Close hood sash to proper position to maintain laminar air flow
• (b) Avoid cluttering

Autoclaving

• (a) Pipette tips (or can be purchases pre-autoclaved, DNAse/RNAse free)
• (b) Glass 9” Pasteur pipettes
• (c) 70% ethanol  (Be sure to spray all surface areas)

​All media, supplement, and reagents must be sterile to prevent microbial growth in the cell culture. Some reagents and supplements will require filter sterilization if they are not provided sterile.

Watch our aseptic technique video protocol​ for detailed guidelines on avoiding contamination.

2) Preparation of cell growth medium

Before starting work check the information given with the cell line to identify what media type, additives, and recommendations should be used.

Most cell lines can be grown using DMEM culture media or RPMI culture media with 10% Foetal Bovine Serum (FBS), 2 mM glutamine and antibiotics can be added if required (see table below).

Check which culture media and culture supplements the cell line you are using requires before starting cultures. Culture media and supplements should be sterile. Purchase sterile reagents when possible, only use unders aseptic conditions in a culture hood to ensure they remain sterile.

​​General example using DMEM media

3) Creating the correct culturing environment

Most cell lines will grow on culture flasks without the need for special matrixes etc. However, some cells, particularly primary cells, will require growth on special matrixes such as collagen to promote cell attachment, differentiation, or cell growth. We recommend reviewing the relevant literature for further information on the cells you are culturing.

The following is an example for endothelial and epithelial cells:

For human cells, coat flasks with 1% gelatin.  Alternatively, for other cell types such as BAEC, flasks can be coated with 1% fibronectin. 

1. Prepare 10mL of coating solution composed of 1% gelatin or 1% fibronectin by diluting with distilled water, followed by filtration.  This is efficient to coat about 5 flasks.
2. Pipette coating solution into flask.  Rock back and forth to evenly distribute the bottom of the flask.  Let sit in an incubator for 15-30 minutes.
3. Aspirate coating solution and wash with sterile dH₂O before seeing cells. 
   
   

4) Checking cells

Cells should be checked microscopically daily to monitor health, grow rates and confluency (% surface area covered with cell monolayer).

Adherent cells should be mainly attached to the bottom of the flask, show an adherent morphology (cell line dependant) and refract light around their membrane (refer to Abcam cell line data sheet images).

Suspension cells should show a circular morphology and refract light around their membrane. Some suspension cells may clump (dissociation reagents such as Pluronic PF68 could be added to promote clump removal).

Media that includes phenol red should be pink/orange in color (media color may change depending on CO₂ environment). For imaging application media without phenol red can be used and will avoid interference with imaging acquisition. A pale yellow colour of media would indicate a acidity and decrease of pH which is often associated with contamination or unhealthy cells.

Discard cells if:

• They are detaching in large numbers (attached lines) and/or look shrivelled and grainy/dark in color.
• They are in quiescence (do not appear to be growing at all).

5) Sub-culturing

Also referred to as cell splitting and cell passaging.

Split ratios or seeding densities can be used to ensure cells are ready for an experiment on a particular day or maintain cell cultures for future use or as a backup. Suspension cell lines are seeded based on volume so seeding densities will be calculated as cells/mL, whereas adherent cell lines are seeded based on flask surface area so will be calculated as cells/cm². Cell lines often require specific seeding densities so always check the guidelines for the cell line in use. Slow growing cells may not grow if a high split ratio is used. Fast growing cells may require a high split ratio to make sure they do not overgrow.

Adherent cell lines can be split using cell line specific split ratios or seeding densities (cells/cm²):

• 1:2 split should be 70-80% confluent and ready for an experiment in 1 to 2 days
• 1:5 split should be 70-80% confluent and ready for an experiment in 2 to 4 days
• 1:10 split should be 70-80% confluent and ready for sub-culturing or plating in 4 to 6 days. 

Split ratios are based on flask surface area, e.g.:

1 x 25 cm² flask Split 1:3 would yield 3 x 25 cm² flasks or 1 x 75 cm²

Suspension cell lines should be maintained using cell line specific seeding densities (cells/mL):

• 2e5 should be ready for an experiment in 3-4 days
• 1e6 should be ready for an experiment in 1-2 days

If cells are to be left unattended for longer periods (i.e. bank holiday weekends) it is recommended to use a lower than normal seeding density/split ratio.

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6) Adherent subculture protocol (using dissociation reagent) 

When the cells are approximately 80% confluent (80% of the flask surface is covered by cell monolayer), cells should still be in their log phase of growth and will require subculturing. It is not recommended to allow cells to become over confluent as this may negatively affect gene expression and cell viability.
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1. Remove cell culture media and dissociation reagent from the fridge and place in a 37^oC incubator and allow to come to temperate 37^oC.
   - Do not leave media in the incubator for longer than is necessary as the media components will degrade over time.
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2. Switch on and perform a basic clean for your biological safety cabinet.
   - Spray all media bottles, pipettes and centrifuge tubes with ethanol before placing in the biological safety cabinet.
3. Under the biological safety cabinet, remove the conditioned media and gently wash the cell monolayer with room temperature DPBS.
   - Carefully add DPBS to side of flask so not to forcefully dislodge adherent cells.
4. Remove the DPBS using a sterile serological pipette and add pre-warmed dissociation reagent (Trypsin-EDTA) to the flask and place in an incubator for ~2 mins (dissociation times can vary between cell lines). Check flask frequently to ensure all cells have dissociated from flask surface.
   - Not all cells will require trypsinization, and to some cells it can be toxic. Trypsin can also induce temporary internalization of some membrane proteins, which should be taken into consideration when planning experiments. Other methods such as gentle cell scraping or using a very mild dissociation reagent (Versene) can often be used as a substitute in these circumstances.
5. When all cells are detached, neutralise the dissociation reagent with serum containing growth medium appropriate to the cell line in culture.
6. Transfer cell suspension to a centrifuge tube. Using sterile media, wash flask and transfer to centrifuge tube, ensuring all cells have been harvested from flask.
7. Centrifuge the cell suspension for 5min @ 1000rpm, room temperature.
8. Discard the supernatant and gently flick the cell pellet (to break up pellet), then resuspend cells in sterile media to a suitable volume for counting.
9. Consult Abcam counting cell using a hemocytometer protocol.
10. Based on count and viability data, seed cell suspension for an appropriate flask and density, e.g. T175, 30mL at 2e4 cells/cm².
    - Label culture flask with all necessary info e.g. Cell Line, passage number, etc.
11. Immediately incubate the newly seeded cultures in a 37^oC/5% CO₂ air humidified incubator.

7) Sub-culturing loosely attached cell lines requiring cell scraping for sub-culture

1. When ready, carefully pour off media from flask of the required cells into waste pot (containing approximately 100 ml of 10% sodium hypochlorite) taking care not to increase contamination risk with any drips.
2. Replace this immediately by carefully pouring an equal volume of pre-warmed fresh culture media into the flask.
3. Using cell scraper, gently scrape the cells off the bottom of the flask into the media. Check all the cells have come off by inspecting the base of the flask before moving on.
4. Take out required amount of cell suspension for required split ratio using a serological pipette.
   e.g. for 1:2 split from 100 ml take 50 ml into a new flask
   1:5 split from 100 ml take 20 ml into a new flask
   1:10 split from 100 ml take 10 ml into a new flask
5. Top the new flasks up to required volume (taking into account split ratio) with pre-warmed fresh culture media
   eg. in 25 cm² flask approx 5-10 ml
   75 cm² flask approx 10-30 ml
   175 cm² flask approx 40-150 ml
   
   

8) Sub-culturing attached cell lines requiring trypsin

Note – not all cells will require trypsinization, and to some cells it can be toxic. It can also induce temporary internalization of some membrane proteins, which should be taken into consideration when planning experiments. Other methods such as gentle cell scraping, or using very mild detergent can often be used as a substitute in these circumstances.

1. When ready, carefully pour off media from flask of the required cells into waste pot (containing approximately 100 ml 10% sodium hypochlorite) taking care not to increase contamination risk with any drips.
2. Using aseptic technique, pour/pipette enough sterile PBS into the flask to give cells a wash and get rid of any FBS in the residual culture media. Tip flask gently a few times to rinse the cells and carefully pour/pipette the PBS back out into waste pot.

   This may be repeated another one or two times if necessary (some cell lines take a long time to trypsinize and these will need more washes to get rid of any residual FBS to help trypsinization)

3. Using pipette, add enough trypsin EDTA to cover the cells at the bottom of the flask.
   e.g. in 25 cm² flask approx 1 ml
   75 cm² flask approx 5 ml
   175 cm² flask approx 10 ml
4. Roll flask gently to ensure trypsin contact with all cells. Place flask in 37°C incubator. Different cell lines require different trypsinization times. To avoid over-trypsinization which can severely damage the cells, it is essential to check them every few minutes.
5. As soon as cells have detached (the flask may require a few gentle taps) add some culture media to the flask (the FBS in this will inactivate the trypsin)
6. Using this cell suspension, pipette required volume of cells into new flasks at required split ratio. These flasks should then be topped up with culture media to required volume
   e.g. in 25 cm² flask approx 5-10 ml
   75 cm² flask approx 10-30 ml
   175 cm² flask approx 40-150 ml
   Leave cells overnight to recover and settle. Change media to get rid of any residual trypsin.

9) ​Sub-culturing of suspension cell lines​

1. Check guidelines for the cell line for recommended split ratio or subculturing cell densities.
2. Take out required amount of cell suspension from the flask using pipette and place into new flask. 
   ​e.g.For 1:2 split from 100 ml of cell suspension take out 50 ml
   ​For 1:5 split from 100 ml of cell suspension take out 20 ml
3. Add required amount of pre-warmed cell culture media to fresh flask.
   e.g. For 1:2 split from 100 ml add 50 mls fresh media to 50 ml cell suspension
   ​For 1:5 split from 100 ml add 80mls fresh media to 20 ml cell suspension

10) Changing media

If cells have been growing well for a few days but are not yet confluent, then they will require a media changing to replenish nutrients and keep correct pH. Cells produce positive growth promoting factors which are secreted into their media so it can be beneficial to perform a half media change replenish nutrients provided by the media and also maintain these positive growth factors.

To change media, warm up culture media at 37°C using a water bath or incubator for at least 30 min. Aspirate old media from the flask and replace the media with the necessary volume of fresh pre-warmed culture media and return to incubator.

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11) Passage number

The passage number is the number of sub-cultures the cells have gone through. Passage number should be recorded and not get too high. This is to prevent use of cells undergoing genetic drift and other variations.

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