Cell line frequently asked questions

1. Consider how much time, money and effort were spent in developing or obtaining your cell culture: Many widely used cell lines can be purchased for a few hundred dollars/pounds/euros per vial. In contrast, specialized and genetically engineered cultures used for the production of monoclonal antibodies, recombinant therapeutic proteins and vaccines come with higher significant costs. 
   
2. How difficult will it be to replace the culture or repeat the experiment or production run again?  Some cultures, such as hybridomas, may be truly irreplaceable. 
    
3. What are the consequences to your research, laboratory/facility and career if important cultures become contaminated with mycoplasmas or are lost due to other causes, such as an accident or cross contamination by another cell line?  Sometimes, researchers discover their cells lines are contaminated with mycoplasmas only after they send their cell lines to other laboratories who then discover the contamination with testing.   

Authentication is the verification of the identity and purity of human cell lines and is essential to the integrity of your research.

The human cell authentication assay identifies short tandem repeat (STR) markers—tiny repeating segments of DNA found between genes—at specific loci to establish a DNA fingerprint for every human cell line. Multiplex polymerase chain reaction (PCR) is used to simultaneously amplify the amelogenin gene and 17 of the most informative polymorphic markers in the human genome. The pattern of repeats results in a unique STR identity profile for each cell line analyzed.

There are many repositories that provide Human Cell Authentication services following ISO 9001 and ISO/IEC 17025 quality standards. Routine testing of cell cultures is critical as cell lines can undergo misidentification, cross-contamination, and genetic drift. Cell authentication is also now required by most funding bodies, e.g. the National Institutes of Health for funding and by a number of leading scientific journals.  

Aseptic technique is a set of principles and practices used by cell culture workers to reduce the potential of unwanted microorganisms or other cell lines from being introduced into the cell culture they are working with. 

Good aseptic technique is essential for successful long-term cell and tissue cultures. Strict adherence to these principles and practices provides benefits for your cultures. The aims for good aseptic techniques should include:

• protection of the cell line from microbial and cellular cross contamination;
• prevention of compromising the cell line by misidentification; and
• protecting the value of your cell line, experiments and cell culture processes.   

The degree of rigour used for good aseptic technique should always be of value to your cell cultures and their applications, whether for research or bioproduction. This is especially true when working with human cell lines known to contain oncogenic or infectious viruses or other harmful microorganisms. 

• All supplies and reagents that come into contact with the cultures must be sterile. 
• Wash hands before and after handling any cell culture. Hand washing stations should be readily accessible within the laboratory.  
• Wear appropriate personal protective equipment (PPE) 
• Handle only one cell line at a time. There are intrinsic risks of misidentification or cross contamination between cell cultures when more than one cell line is in use within the laboratory. 
• Use only one set of dedicated reagents for each cell line.  
• Handle continuous cell lines after the handling of short-term, finite cell cultures to prevent carry-over.  
• Quarantine and handle under strict precautions all incoming cell lines until testing determines the absence of mycoplasma. Alternatively, purchase cell lines from repositories which certify that materials are mycoplasma-free prior to distribution.  
• Avoid continuous long-term use of antibiotics within cell cultures. The overuse of antibiotics as prophylaxis may lead to cytotoxicity and pose an increased risk of covert mycoplasma contamination within the cell lines.  
• Cultures should be inspected daily for signs of contamination. In addition, testing at regular intervals for mycoplasma should be conducted to ensure the purity and integrity of the culture.  
• Promptly discard any contaminated cultures. Retention of these cultures poses a serious threat of cross contamination to other cultures in the laboratory. If clean-up/treatment of the contaminated culture is attempted, then any work with this culture should be performed in a separate laboratory for that purpose or reserved to the very end of the day to minimize transfer of the contamination. 

Cell or tissue culture refers to the growth and maintenance of cells derived from tissues of an animal, from insect to human, in an artificial environment often referred to as in vitro.

There are two ways to grow in vitro cell cultures;

1. Adherent cell culture – Cells are grown as monolayers in plastic dishes or flasks. Some cell types can be grown on gel-like substrates as multicellular aggregates to better mimic a physiological environmental stiffness.
2. Suspension cell culture – Cells are grown without adhesion contact to the substrates suspended in cell media. Generally, to ensure optimum cell growth, the suspension is agitated regularly.

Adherent cell culture is the most commonly used method of growing cells as it is able to be used across a wide variety of cell culture types. Suspension cell culture is useful as it can be easily scaled up for protein purification.

A primary cell culture refers to cells that have been removed from a live donor/specimen and have been established in an in vitro environment.

Secondary cell culture refers to cell lines that have been immortalized, usually by overexpressing an enzyme called human telomerase reverse transcriptase (hTERT), and can divide indefinitely. Many tissue types are not amenable to immortalisation, so primary cell culture may be the only option in some circumstances.

Both primary and secondary cell cultures have advantages and disadvantages:

1. Using primary cell cultures will provide biological results in specific cells of interest, that are more accurate to those seen in the host tissue. However, there will be a finite number of cells with a finite lifespan for you to use.
2. In contrast, secondary cell cultures will be easier to maintain and have a more uniform and long-lasting cell population, but as the cultures have grown and been passaged in artificial cell culture media, they will provide biological responses different from cells within the original host tissue. Additionally, these cells may have genetic drifting, changes or mutations acquired during repeated passaging and culture maintenance.

To overcome some of the limitations of secondary cell cultures, some researchers prefer to use human plasma-like cell culture media such as Plasmax^TM, rather than standard media.

Finite Cell Lines are cell cultures that will only survive a certain number of population doublings before senescence. All normal cells are finite. 

Passage:

The transfer or transplantation of cells, with or without dilution, from one culture vessel to another. It is understood that any time cells are transferred from one vessel to another, a certain portion of the cells may be lost and therefore, dilution of cells, whether deliberate or not, may occur. This term is synonymous with the term ‘subculture’. 

Passage Number:

The number of times the cells in the culture have been sub-cultured or passed. In descriptions of this process, the ratio or dilution of the cells should be stated so that the relative cultural age gap can be ascertained. This term is not synonymous with population doubling. 

Population Doubling Level:

The total number of population doublings of a cell line or strain since its initiation in vitro.

A formula to use for the calculation of ‘population doublings’ in a single passage is: number of population doublings = Log10(N/N0) X 3.33 where: N= number of cells in the growth vessel at the end of a period of growth N0= number of cells plated in the growth vessel. It is best to use the number of viable cells or number of attached cells for this determination. Population doublings level is synonymous with ‘cell generation time’. 

Seeding density refers to the number of cells either per cm2 (if an adherent cell line) or per ml (if a suspension cell line) that are resuspended in a fresh flask after subculture.

The recommended seeding density is usually provided in the relevant literature or website listing. The recommend numbers reflects the limits of a cell line to survive a split (ie. too low and there will not be a large enough population to successfully colonise the flask, too high and insufficient nutrients and space will be available for growth of any considerable length of time. If no recommendation values are available then aim to seed adherent cells at between 2-4 x 104 cells per cm2 and suspension cells at between 2-4 x 10 5 per ml. 

Procedure for Suspension Cell Lines 

• View cultures using an inverted phase contrast microscope. Cells growing in exponential growth phase should be bright, round and refractile.  
• Hybridomas may be very sticky and require a gentle knock to the flask to  detach the cells. EBV transformed cells can grow in very large clumps that are very difficult to count and the centre of the large clumps may be non-viable. 
• Do not centrifuge to subculture unless the pH of the medium is acidic (phenol red indicator = yellow) which indicates the cells have overgrown and may not recover.
• If this is so, centrifuge at 150 x g for 5 minutes, re-seed at a slightly  higher cell density and add 10-20% of conditioned medium (supernatant) to  the fresh media. 
• Take a small sample (100-200μl) of the cells from the cell suspension and count the cells.
• Calculate cells/ml and re-seed the desired number of cells into freshly prepared flasks, without centrifugation, just by diluting the cells. Refer to the data sheet supplied with the cell line for the recommended seeding density. 
• Repeat this every 2-3 days. 

 Procedure for Adherent Cell Lines 

• View cultures using an inverted microscope to assess the degree of confluency and confirm the absence of bacterial and fungal contaminants. 
• Remove spent medium. 
• Wash the cell monolayer with PBS without Ca2+/Mg2+ using a volume equivalent to half the volume of culture medium. Repeat this wash step if the cells are known to adhere strongly. 
• Pipette trypsin/EDTA onto the washed cell monolayer using 1ml per 25cm2  of surface area.
• Rotate flask to cover the monolayer with trypsin. Decant the excess trypsin. 
• Return flask to the incubator and leave for 2-10 minutes. 
• Examine the cells using an inverted microscope to ensure that all the cells are detached and floating. The side of the flasks may be gently tapped to release any remaining attached cells. 
• Resuspend the cells in a small volume of fresh serum-containing medium. 
• To inactivate the trypsin: Remove 100-200µl and perform a cell count. In the case of cells cultured in serum-free media, use a trypsin inhibitor e.g. soyabean trypsin inhibitor to inactivate the trypsin. 
• Transfer the required number of cells to a new labelled flask containing pre-warmed medium (refer to the appropriate Cell Line Data Sheet for the required seeding density). 
• Incubate as appropriate for the cell line. 
• Repeat this process as demanded by the growth characteristics of the cell line.

The Do’s 

• Use personal protective equipment (PPE), (laboratory coat/gown, gloves and  eye protection) at all times. In addition, thermally insulated gloves, full-face  visor and splash-proof apron should be worn when handling liquid nitrogen. 
• Use disposable head caps to cover hair. 
• Wear dedicated PPE for the tissue culture facility and keep separate from PPE worn in the general laboratory environment. The use of different coloured gowns or laboratory coats makes this easier to enforce. 
•  Keep all work surfaces free of clutter. 
• Correctly label reagents including flasks, medium and ampoules with contents and date of preparation. 
• Only handle one cell line at a time. This common-sense point will reduce the possibility of cross contamination by mislabelling etc. It will also reduce the spread of bacteria and mycoplasma by the generation of aerosols across numerous opened media bottles and flasks in the cabinet. 
• Clean the work surfaces with a suitable disinfectant (e.g. 70% isopropanol) between operations and allow a minimum of 15 minutes between handling different cell lines. 
• Maintain separate bottles of media for each cell line in culture. 
• Examine cultures and media daily for evidence of gross bacterial or fungal contamination. This includes medium that has been purchased commercially. 
• Check quality control information for all media and reagents and ideally testperformance prior to use. 
• Keep cardboard packaging to a minimum in all cell culture areas. 
• Ensure that incubators, cabinets, centrifuges and microscopes are cleaned and serviced at regular intervals. 
• Test cells for the presence of mycoplasma on a regular basis 

The Don’ts 

• Do not use antibiotics continuously in culture medium as this can lead to the appearance of antibiotic resistant strains and may mask underlying  contamination. 
• Do not allow waste to accumulate particularly within the microbiological safety cabinet or in the incubators. 
• Do not have too many people in the lab at any one time. 
• Do not handle cells from unauthenticated sources in the main cell culture suite. They should be handled in quarantine until quality control checks are complete. 
• Avoid keeping cell lines continually in culture without returning to frozen stock. 
• Avoid cell cultures from becoming fully confluent. Always sub-culture at 70- 80% confluency or as advised on the ECACC cell culture data sheet. 
• Do not allow media to go out of date. The shelf life is only 4 – 6 weeks at +4°C once glutamine and serum are added. 
• Avoid water baths from becoming dirty by regular cleaning. 
• Do not allow essential equipment to become out of calibration. Ensure microbiological safety cabinets are tested regularly. 

Typical quality control tests for a cell line would include:

• viability,
• cell number,
• growth,
• morphology,
• subculture following resuscitation

In addition, cell line identity (authentication or speciation), sterility (absence of bacterial and fungal contamination), absence of mycoplasma, absence of adventitious viruses is also performed.

Extra validation tests such as expression of a transgene or cellular behaviour can also be stated on the CoA as required. 

Quality Assurance (QA) is a programme or system that focuses on operational standards for all aspects of activities, to ensure that a procedure or products, e.g. cell lines, consistently meet quality standards. 

Quality Control (QC) Testing that is used to assess specific quality outcomes and is one component of a QA programme. Testing may extend to cell lines (referred to as validation), media and other reagents, culture vessels, equipment and facilities. 

A Certificate of Analysis (CofA) is a document describing the tests applied to a given batch of manufactured products (cell lines). For example, a frozen stock of a cell line will be tested upon resuscitation for viability, cell number, growth, morphology, subculture and 

Other tests will include: cell line identity (authentication or speciation), sterility (absence of bacterial and fungal contamination), absence of mycoplasma, absence of adventitious viruses. Other validation test such as expression of a transgene or cellular behaviour or expression of relevant serotype (hybridomas) can also be stated on the CoA as required.

Animal cell line identity is a crucial first step in cell line authentication, is frequently underappreciated and ignored by most research scientists. To address this need, a method of authenticating cell lines by species known as DNA barcoding was developed. In this method, species identification is achieved using a short section of DNA from a specific gene that is highly conserved among species. Cytochrome oxidase 1 (CO1) gene is the most commonly used gene for species identification. 

CO1 Barcoding: CO1 is a mitochondrial gene expressed in all animal species due to its integral role in energy production via the electron transport chain. Because inheritance of mitochondrial genes (MITO genes) is maternal, animals typically have only one variant of each MITO gene and these genes show more divergence compared to nuclear genes. CO1 DNA barcoding has been proven to be a highly effective tool for species identification due to the following reasons: the gene is relatively easy to amplify, only one variant of the gene exists. There is a high degree of evolutionary divergence among species-specific homologues of that gene.  Based on this species-to-species sequence variability of the CO1 gene, protocols have been developed for a PCR-based speciation assay using unique primer pairs that are species-specific and produce amplicons in a multiplex PCR reaction ≥ 20 base pairs apart.  

The CO1 assay is capable of distinguishing cell lines of pig, human, cat, Chinese hamster, Rhesus monkey, sheep, horse, African green monkey, rat, dog, mouse, rabbit, goat and cow origin. When the species of a cell line remains in question the ~650bp ‘barcode’ region of the CO1 gene can be sequenced for verification purposes. Most providers of this service perform CO1 analysis following ISO 9001:2015 and ISO/IEC 17025:2017 quality standards. 

Effective segregation of cell lines is essential to safeguard the credibility of your work especially in shared laboratories. In order to do this we would recommend the following steps: 

• Make sure the microbiological safety cabinet is clutter free and cleaned before you begin working on your cells. This can be done with 70% IPA on a daily basis. 
• Only work with one cell line at a time in the cabinet. Once you have finished working on the cell line spray down the cabinet with 70% IPA, allow it to settle and then set up the cabinet for the next cell line. This reduces the chances of cross contamination. 
• Keep separate and clearly labelled bottles of media and all other reagents for each cell line
• Aliquot media and reagents from stock bottles for use in the cabinet where possible.
• Discard any unused media and reagents; never return this to the stock bottles
• Practice good aseptic techniques 
• If possible, keep flasks segregated in the incubators 
• Keep accurate and detailed records 
• If you suspect that your cells may have become cross contaminated,  arrange for authentication testing of your cells.

Definition:

Cryopreservation refers to the ultra-low temperature storage of cells, tissues, embryos or seeds. This storage is usually carried out in the vapour or liquid phase of nitrogen. 

Cryopreservation:

Most cell cultures can be stored for many years, if not indefinitely, at temperatures below –130°C (cryopreservation). The advantages of cryopreservation include: generation of safety stocks to ensure against loss of the culture from equipment failures or contamination by microorganisms or other cell lines, elimination of the time, energy, and materials required to maintain cultures not in immediate use., preservation of cells with finite population doublings (that will ultimately senesce), insurance minimise or protect against phenotypic drift in the culture due to genetic instability and/or selective pressure and creating a standard reagent to be used for a series of experiments. 

Overview of cryopreservation procedure. 

As the cell suspension is cooled below the freezing point, ice crystals form and the concentration of the solutes in the suspension increases. The formation of intracellular ice can be minimized if water within the cell is allowed to escape by osmosis during the cooling process. A slow cooling rate, generally –1°C per minute, facilitates this process. However, as the cells lose water, they shrink in size and will quickly lose viability if they go beyond a minimum volume. The addition of cryoprotectant agents such as glycerol or dimethylsulfoxide (DMSO) will mitigate these effects. 

The standard procedure for cryopreservation required the freezing of cells slowly until they reach a temperature below –70°C in medium that includes a cryoprotectant like DMSO. Vials are transferred to a liquid-nitrogen freezer to maintain them at temperatures below –130°C. 

Composition of Freeze medium.

For cells grown in serum-supplemented medium:

Glycerol and DMSO are used at 5% to 10% respectively. They are the most common cryoprotectant agents. While DMSO can be toxic to cells, it penetrates them much faster than glycerol and yields more reproducible results. Unfortunately, DMSO can cause some cells to differentiate (eg, HL-60 promyeloblast cells) and may be too toxic for other cells (e.g, HBE4-E6/E7 lung epithelial cells). Glycerol should be used as alternative in these instances. Glycerol can be sterilized by autoclaving whereas DMSO must be sterilized by filtration.  

For cells grown in serum-free medium:

adding 50% conditioned medium (serum-free medium in which the cells were grown for 24 hours) to both the cell freezing and the recovery medium may improve recovery and survival. The addition of 10% to 20% cell culture-grade bovine serum albumin to serum-free freezing medium may also increase postfreeze survival. 

Overview:

The recovery of cryopreserved cells is straightforward: vials containing cells are thawed rapidly in a water bath at 37°C, removed from the freeze-medium by gentle centrifugation and/or diluted with growth medium, and seeded in a culture vessel in complete growth medium. 

Factors which affect the viability of recovered cells.

The resuscitation procedure may have to be modified in certain cases for each cell line to attain optimal cell viability upon recovery. Some of the critical parameters for optimization include the composition of the freeze medium, the growth phase of the culture, the stage of the cell in the cell cycle, and the number and concentration of cells within the freezing solution. 

What to expect after resuscitation?

Some cell lines, such as hybridoma cultures, take several days before they fully recover from cryopreservation. Some hybridomas show low viability on the first day in culture and will generate cellular debris. Viability for most cells declines and reaches a low point at 24 hours post-thaw. Most, if not all, of this decline appears to be due to apoptosis, rather than necrosis, induced by the stress of the cryopreservation process.  After this time point, cells begin to recover and enter exponential growth. 

A typical resuscitation procedure.

• Prepare a culture vessel (T-75 flask) so that it contains at least 10 mL of the appropriate culture medium equilibrated for temperature and pH.
• Remove the vial from the liquid nitrogen freezer and thaw by gentle agitation in a 37°C water bath (or a bath set at the normal growth temperature for that cell line).
• Thaw rapidly until ice crystals have melted (approximately 2 minutes).
• Remove the vial from the water bath and decontaminate it by dipping in or spraying with 70% ethanol.
• *Follow strict aseptic conditions in a laminar flow tissue culture hood for all further manipulations.
• Unscrew the top of the vial and transfer the contents to a sterile centrifuge tube containing 9 mL complete growth medium.
• Remove the cryoprotectant agent by gentle centrifugation (10 minutes at 125 × g).
• Discard the supernatant, taking care not to disturb the soft pellet, and resuspend the cells in 1 mL or 2 mL of complete growth medium.
• Pipette gently to loosen the pellet and break apart clumps. (If the cells normally grow as clusters, avoid over-pipetting during resuspension.)
• Transfer the cell suspension into the medium in the culture vessel and mix thoroughly.
• Examine the cultures after 24 hours and subculture as needed. 

For convenience, most laboratories will quote the speed of the centrifuge when describing a centrifugation step. This will only apply and remain constant if the same centrifuge and rotor are used. Centrifugation is better defined by the value of g, the radial acceleration relative to gravity. 

For example: g equals 1.18 x 10-5 multiplied by the radius of the rotor to the center of the tube multiplied by the rotations per minute2. In a centrifuge rotor rotating at 1500 rpm, with a of 18cms from the center of the spindle to the middle depth of the sample in the centrifuge tube, the g equals 1.18 x 10-5 multiplied by 18 and 15002 giving a value of 452.8 or approximately 450. Most benchtop centrifuges used in a cell culture lab will display the rotational speed in values for either g or rpm. Accompanying literature for the centrifuge will provide a conversion chart. 

If vials are immersed in liquid phase LN2 there is a risk of LN2 seeping into the vial; this could result in cross-contamination and an increased risk of the vial exploding when thawed. 

Although it can be tempting to source cell lines from colleagues they may inadvertently supply you with contaminated or misidentified cell lines. If they can provide you with confirmation that the cells are mycoplasma free and the DNA profile has been confirmed to be as expected you can have more confidence.  If not, is it worth taking the risk?