- Western blotting guide
- Western blotting buffers & recipes
- Western blot sample protocol
- Western blotting dictionary
- Co-Immunoprecipitation protocol
- Western blot troubleshooting tips
- Mammalian cell & tissue culture guide
- How to use a haemocytometer
- RIPA Buffer – Cell lysis buffer
- Subcellular fractionation protocol
Cell Culture Guide – Techniques and Protocols
Below is a guide for mammalian cell tissue culture, dealing with maintenance, splitting and cyropreservation of mammalian cell lines. Please note that all cell culture must be undertaken in a laminar flow hood using aseptic technique.
Cell culture is a key tool used in cellular and molecular biology, enabling the modelling of the physiology and biochemistry of cells. Additionally, cell culture enables researchers to determine the effects of drugs and toxic compounds on cellular responses. Cell culture remains one of the most widely used techniques in research today due to the consistency and reproducibility of results that can be obtained from using a batch of clonal cells. The growth of cells from an animal or plant in a favorable environment is known as cell culture. Different cells have different culturing requirements necessary for optimum growth, which can be catered for via medium selection and supplementation. The medium used acts to supply the essential nutrients (amino acids, carbohydrates, vitamins, minerals), growth factors, hormones, gases (O2, CO2) and regulates the physicochemical milieu (pH, osmotic pressure, temperature).
A primary culture refers to cells which have been cultured directly after tissue isolation. Once these cells become confluent i.e. occupy all available space in the flask they need to be passaged or split by transferring them to a new growth vessel which will provide the cells with more room for continued growth and expansion. Following the first passage the primary culture now becomes known as a cell line. Cell lines derived from primary cultures have a limited life span, meaning that they can only divide a limited number of times before becoming senescent. As the cells are passaged cells with the highest growth capacity predominate, resulting in genotypic and phenotypic uniformity in the population. Immortal cell lines are widely used in cellular and molecular biology in order to bypass problems such as senescence. Cells can become transformed and thereby immortalized through a number of different means; spontaneously, chemically or virally. Once transformed these cells acquire the ability to divide indefinitely and become a continuous cell line.
Cell lines can either be:
- Adherent (Anchorage dependent)- Adherent cells must be cultured while attached to a solid or semi-solid substrate and will often require trypsinization for subculturing (see 2.6 for more details)
- Suspension (Anchorage Independent) – Suspension cells do not require a solid growth substrate and can be grown floating in culture medium.
Below is a general guideline for the culturing of cell lines. All cell culture must be undertaken in microbiological safety cabinet using aseptic technique to ensure sterility, protective clothing must be worn throughout.
Good aseptic technique is designed to create a sterile environment and act as a barrier between microorganisms and the sterile cell culture hood. Key aspects of this technique include the use of sterile reagents, media. Sterile handling ensures that nothing is entered into the hood which hasn’t been sprayed with 70% Ethanol.
Laminar Flow Hood Preparation
The cell culture hood provides as aseptic work area while ensuring the containment of any infectious splashes or aerosols generated by microbiological procedures. There are 3 kinds of cell culture hoods, designated as Class I, II and III, these have been developed to meet varying research and biosafety needs. Choose the right one for your experimental needs.
Using the Laminar Flow Hood
- Remove the cover and turn on the hood.
- Close the hood sash to the proper position to maintain laminar air flow.
- Spray down all surface areas of the hood with 70% ethanol.
- Avoid cluttering.
- Ensure all equipment used (ie Pipette tips, pipette, racks, eppis, flasks, reagents) is sterile before putting them in the hood. This can be done by autoclaving or through the use of purchased pre-autoclaved, DNAse/RNAse free equipment. Spraying down pipettes and racks with 70% Ethanol.
- 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.
- Avoid pouring media and reagents directly from bottles or flasks.
- Use each pipette only once to avoid cross contamination.
- Do not unwrap sterile pipettes until they are ready to be used and do this inside the hood.
- Never uncover a sterile flask, bottle, petri dish, etc. until the instant you are ready to use it and never leave it open to the environment. Return the cover as soon as you are finished.
- If you remove a cap or cover, and have to put it down on the work surface, place the cap with opening facing up.
Before starting ensure that you are working with the right media type and culture supplements for your cell line of interest. These must always be sterile and only opened inside the hood. Most cell lines can be grown using Dulbecco’s Modified Eagle Medium (DMEM) culture media or Roswell Park Memorial Institute (RPMI) culture media with 10% Foetal Bovine Serum (FBS), 2 mM glutamine and antibiotics can be added if required.
Example- How to prepare DMEM Media
- Remove 50 ml of media from a 500ml bottle.
- Volume is now 450 ml.
- Add 10% FBS = 50 ml.
- Add supplements 2 mM glutamine = 5 ml, 100 U penicillin / 0.1 mg/ml streptomycin = 5 ml.
Certain endothelial cell lines require special collagen matrices to grow. These matrices ensure attachment, differentiation and growth of the cell line. Before starting your cell culture experiment it is important to carry out a literature review of your cell line of interest to ensure any additional growth requirements such as matrices are meet. Be aware of the type of flask you are using i.e. vented or non-vented. If using a non-vented/plug flask be sure to loosen the cap to allow gaseous exchange. If using a vented flask ensure that the filter does not get wet as this will affect the efficacy of gaseous exchange.
Example- How to prepare a 1% Gelatin matrix for endothelial and epithelial cells.
- Prepare 10mL of coating solution composed of 1% gelatin/1% fibronectin by diluting with distilled water followed by filtration.
- This will coat about 5 flasks.
- Pipette the coating solution into flask.
- Rock back and forth to evenly distribute the matrix on the flask.
- Leave to sit in incubator for 15-30 minutes
- Completely remove the coating solution by aspirating before seeding.
Cells should be checked daily under the microscope to ensure they are healthy and growing as expected. It is important to become familiar with what healthy cells look under the microscope as this will make it easier to detect quiescent or infected cells through changes in morphology.
Discard cells if
- The media changes from a pinky/red colour to a murky yellow. This is a sign of infection and pour aseptic technique.
- If adherent cells are detaching in large numbers, indicative of cell death.
- If the appearance of the cells has changed dramatically – looking disheveled and grainy.
- If they become quiescent.
Mammalian cells should be split/passaged when they reach confluence, which occurs when 70-80% of the flask is covered with cells. In adherent cells confluence can be noted under the microscope when there is no visible space available for cellular growth. For suspension cells confluence can be noted when the cells clump together and the medium appears turbid when the flask is swirled. It is important not to let the cells become over confluent, with 70-80% confluency being the optimum amount, after such a point contact inhibition comes into effect and the cells require a longer recovery period following re-seeding. The speed at which your cell line grows will determine the split ratio used. For instance slow growing cell lines will not do well if split at a high ratio. Fast growing cell lines may require a high split ratio as it will take them a shorter amount of time to reach confluency in comparison to slower growing cell lines. As a general rule cells should not be split more than 1:10 as this is too low for the cells to survive. Varying the seeding density of your cultures will ensure that your cells are ready for an experiment on a particular day. As a general guide, from a confluent flask of cells: 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. However this may vary depending on the growth rates of your cell line.
- Ensure cells are least 80% confluent.
- Warm fresh cell culture media at 37°C in a water bath or incubator for at least 30 minutes.
- Ensure your flasks are labelled correctly with the passage number, cell line, split ratio and date.
- Prepare a waste container with bleach for waste medium containing approximately 100ml of 10% sodium hypochlorite.
- Carefully pour off the media into the waste container.
- Add an equal volume of pre-warmed fresh culture media into the flask.
- Gently scrape cells off the bottom of the flask into the media using a cell scraper.
- Ensure all cells have been scraped off the flask.
- Take out the required amount of cell suspension 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.
- Add the required volume (taking into account split ratio) to pre-warmed fresh culture media. e.g. in 25 cm2 flask approx 5-10 ml 75 cm2 flask approx 10-30 ml 175 cm2 flask approx 40-150 ml.
- Carefully pour off the media from the flask into the waste container.
- Using aseptic technique, pipette pre-warmed PBS into the flask to wash the cells and remove any residual media and FBS.
- Gently rock the flask back and forth to rinse the cells with PBS and pour into the waste container. Repeat this 3 times. It is important to remove any residual FBS for efficient trypsinization.
- Once the washes are complete add trypsin EDTA (also pre-warmed). Enough trypsin should be used so that the cells are covered. For 25 cm2 flask approx 1 ml, 75 cm2 flask approx 5 ml, 175 cm2 flask approx 10 ml.
- Like with PBS gently rock the flask back and forth to ensure trypsin contact with all cells.
- Place the flask in a 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 under the microscope whilst gently tapping the flask for detachment.
- Once the cells have become detached add some culture media to the flask. The FBS in the media will inactivate the trypsin.
- Pipette gently up and down the wall of the flask to remove any additional adherent cells.
- Count the cells and pipette the required volume of cells into new flasks. 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.
* Trypsinization can be toxic to some cells. It can also induce temporary internalization of some membrane proteins, this should be taken into consideration when designed experiments. Other methods such as gentle cell scraping, or use of detergent can often be used as a substitute in these circumstances.
- Some suspension cell lines will have recommended split ratio or sub-culturing cell densities. Check this before you begin.
- Take out required amount of cell suspension from the flask using a pipette and place into a 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 20ml
- Add the required amount of pre-warmed cell culture media to a fresh flask. e.g. For 1:2 split from 100 ml add 50mls fresh media to 50 ml cell suspension. For 1:5 split from 100 ml add 80mls fresh media to 20 ml cell suspension.
If your cells have growing for a few days but are not confluent enough to be split the media will have to be changed to replenish nutrients and maintain pH.
For adherent cells
- Pre-warm the media for at least 30 minutes before use in a waterbath or incubator.
- Pour off the old media into a waste container.
- Replace with the same volume of prewarmed new media and return to incubator.
For suspension cells
- Pre-warm the media for at least 30 minutes before use in a waterbath or incubator.
- Pour the media containing the cells into a 15 ml or 50 ml falcon tube and centrifuge. The rpm and centrifugation time may vary between cell lines.
- Following centrifugation your cells should be pelleted at the bottom of the falcon, pour the old media into waste and resuspend the pellet in the same volume of media as previously used.
- Place in a fresh flask and return to incubator.
The passage number is the number of sub-cultures the cells have gone through. Passage number should be recorded and not get too high. Cell lines with passage numbers of greater than 30 are more likely to acquire genetic abnormalities compared to lower passage cells (Esquenet et al., 1997; Lin et al., 2003; O’Driscoll et al., 2006). P30 is generally accepted as the upper limit of passage number for certain cells in culture, as further culturing may result in substantial variation of molecular profiles, limiting their in vivo applications and reproducibility (Calles et al., 2006; O’Driscoll et al., 2006; Wenger et al., 2004).
Freezing down cell lines is essential component of cell culture as replacing cell lines is expensive and time consuming, it also ensures that if your cells become infected you have back up stocks and can start again. As soon as surplus cells become available, preferably at a low passage number to limit genetic drift, they should be frozen as seed stocks. Working stocks can then be prepared from the seed stock. The best and most widely used method for cryopreserving cells is storing them in liquid nitrogen in media with a cryoprotective agent such as dimethylsulfoxide (DMSO). Cyroprotective agents such as DMSO reduce the freezing point of the medium and enable a slower cooling rate, which greatly reduces the risk of ice crystal formation, known to cause cell death and damage.
Protocol for freezing cells
- Determine the total number of viable cells via a cell count and the required volume of freezing medium.
- Centrifuge the cell suspension. Centrifugation speed and duration can vary depending on cell type.
- Decant the supernatant into waste without disturbing the pellet.
- Resuspend the pellet in cold freezing medium at the recommended viable cell density for the specific cell type.
- Aliquot the cell suspension into cryogenic storage vials, label clearly the date, cell type and passage number.
- Freeze the cells in a controlled rate freezing apparatus, decreasing the temperature approximately 1°C per minute. Alternatively, place the cyrovials containing the cells in an isopropanol chamber and store them at –80°C overnight.
- Transfer frozen cells to liquid nitrogen, and store them in the gas phase above the liquid nitrogen.
*Freeze your cultured cells at as a high concentration and as low a passage number as possible. Ensure that at least 90% of the cells are viable before freezing them down. Always use sterile cryovials for storing frozen cells. Always use proper sterile technique and work in a laminar flow hood.
Safety Notes Caution should be exerted when using DMSO as it is known to facilitate the entry of organic molecules into tissues. Use gloves and wear appropriate protective clothing when handling this reagent. Dispose of the reagents in compliance with local regulations.
Thawing cells is extremely stressful to the frozen cells and therefore unlike freezing down cells which should be done slowly, thawing cells should be done as rapidly as possible, to ensure a high survival rate of cells.
Protocol for thawing cells
- Remove cells from liquid nitrogen using caution.
- Immediately thaw the frozen cells rapidly (<1 minute) in a water bath at 37°C.
- When the ice has thawed transfer into a laminar flow hood.
- Transfer the now thawed cells drop by drop into a centrifuge tube containing the desired amount of pre-warmed media.
- Centrifuge the cell suspension. Centrifugation speed and duration can vary depending on cell type.
- Following centrifugation remove the supernatant with disturbing the pellet.
- Carefully resuspend in fresh medium and transfer into an appropriate flask size and incubate.
One of the most common contaminants in cell culture laboratories is mycoplasma. Mycoplasma is a basic bacterium lacking a cell wall, considered to be the smallest self-replicating organism. Mycoplasma are very difficult to detect as a result of their small size approximately > 1 micrometer, it is not until they achieve extremely high densities and cause the cell culture to deteriorate that their presence becomes known. Many slow growing mycoplasma can persist in culture undetected without causing cell death however, they can alter the behaviour and metabolism of the host cell in culture. Chronic mycoplasma infections may also lead to a decreased rate of cell proliferation, reduced saturation density, and agglutination in suspension cultures. The only way to detect such mycoplasma infections is by periodically testing the cell cultures using; fluorescent staining (e.g., Hoechst), ELISA, PCR, immunostaining, autoradiography, or microbiological assays.
Antibiotics in cell culture should be used sparingly. Continuous use of antibiotics encourages antibiotic resistance, allows low-level contamination to exist and can mask a variety of contaminants. Some antibiotics can also cross react with the cells and interfere with the cellular processes under investigation.
Cell culture laboratories can vary greatly depending on research institute and area of investigation. All cell culture laboratories do however have some essential common equipment and requirements, the most important of which being the maintenance of an aseptic environment free from pathogens. Below is a list of the common equipement and materials you can expect in more cell culture laboratories. This list is not all inclusive, deviations from which can be noted depending on research area.
- Cell culture hood (i.e., laminar-flow hood or biosafety cabinet)
- Incubator (humid CO2 incubator recommended)
- Water bath
- Refrigerator and freezer (–20°C)
- Cell counter (Automated Cell Counter or hemocytometer)
- Inverted microscope
- Liquid nitrogen (N2) freezer or cryostorage container
- Cell culture vessels (e.g., flasks, Petri dishes, roller bottles, multiwell plates)
- Pipettes and pipettors
- Syringes and needles
- Waste containers
- Media, sera, and reagents