4.2 Cell culture procedures

The following important conditions must be satisfied to achieve successful cell culture:

·         Incubation temperature should be 36°C.

·         The pH for growth should be between 7.2 and 7.4.

The levels of glucose and L-glutamine can influence cell growth, and correct levels for each cell line should be checked before attempting to put it into culture (typical levels for glucose and L-glutamine are 1-4 mM and 2 mM, respectively). A range of inorganic ions, amino acids and vitamins are essential for cell survival and will usually be included in basal growth media from proprietary sources. Both oxygen and carbon dioxide are essential and are provided either as a mixture of CO₂ and air supplied to the culture vessel or by sealing the vessel tightly to retain the CO₂ produced by cell metabolism.

4.2.1    Aseptic technique

Skill in aseptic technique is important to maintain sterility during media preparation and cell cultivation procedures. Furthermore, it is a vital component in ensuring operator protection from infectious agents that may be present in culture materials. Some important elements in aseptic technique are:

·         Sterilize all glassware for handling cell cultures and media (see below).

·         Avoid splashes, spills and aerosols.

·         Avoid liquid transfer by pouring.

·         When adding (or replacing) medium, never touch the neck of the culture flasks with the bottle containing the medium or use the same pipette to transfer medium to more than one bottle. Ideally, aliquot the total amount of medium required for each batch of culture bottles being handled and store the remainder at 4–8°C. Dedicate separate medium for each cell line.

·         Separate clean and contaminated materials in the BSC II.

·         Minimize exposure of sterile media and cell cultures to open air (even within the BSC II).

·         Perform any final preparation of sterile media (i.e. addition of serum or other additives) before dealing with cell cultures.

Because of the risks of contamination and cross-infection, cell culture in the virus diagnostic laboratory is best carried out in closed vessels, usually screw-capped tubes and flat-sided bottles. WHO does not recommend the use of 24-well plates for the isolation of polioviruses from stool specimens as this method is inappropriate to conditions encountered in many laboratories of the Global Polio Laboratory Network. Cultures are initially set up in growth medium supplemented with 10% serum. Once the cells have formed a confluent monolayer, cultures are changed to maintenance medium which is designed to maintain cultures in a healthy state for as long as possible without stimulating growth; this is achieved by reducing the serum content, usually to 2%.

4.2.2    Preparation of glassware

Due to the difficulty of cleaning and recycling glassware to culture quality, many laboratories have resorted to using disposable cell culture plasticware. If a laboratory chooses to use glassware, however, it must ensure that all glassware is meticulously cleaned and sterilized so that cell cultures will not be affected by traces of proteinaceous material, detergent, pyrogens, water deposits and other residual materials which may get deposited on the glassware.

Glassware cleaning protocols should be developed along the lines of the following procedures:

·         Use care in handling glassware as most breakages occur during the cleaning process.

·         Before cleaning, decontaminate glassware by autoclaving or soaking overnight in chlorine solution (0.5%).

·         Decontaminate pipettes in a container containing chlorine.

·         Rinse all glassware as soon as possible after use.

·         Store soiled items in water containing a disinfectant or cleanser to avoid drying and making items harder to clean.

·         Use 7-X, DECON or similar detergent for thorough cleaning of all laboratory glassware. These detergents are easily rinsed from glassware without leaving residues. (DO NOT use domestic dishwashing liquid detergent under any circumstances.)

·         Clean glass by scrubbing with a brush. Periodically inspect brushes for wear to avoid scratching glass.

·         Thoroughly rinse items in tap water, followed by at least 5–7 changes of distilled or deionized water. Even the smallest residual amounts of cleansers, disinfectants or acids can affect the growth of cell cultures.

·         Dry glassware on racks or peg boards and inspect after drying. If glassware is hazy, has a film or blotches are evident, then additional cleaning is required before use.

·         Sterilize cell culture glassware using a hot air oven at 180°C for three hours to destroy pyrogens. Non-glass components which may not withstand 180°C should be sterilized by alternate methods such as autoclaving, and re-assembled aseptically.

Chromic acid wash: Some heavily soiled glassware may require vigorous methods to clean and traditionally this has required the use of chromic acid (10% potassium dichromate in 25% sulfuric acid). Chromic acid, however, is a hazardous substance, with safety and environmental concerns. There are effective commercially available substitutes to chromic acid which include: Fisher product, Contrad 70 or VWR Scientific products, Chem-Solv, phosphate-free formulations of RBS-35, PCC-54 and Nochromix (also supplied by Fisher).

If chromic acid must be used, follow all normal safety precautions for using concentrated acids and acid solutions. As with any other cleaning process, all cleaning solutions must be completely rinsed from the glassware through copious changes of tap water followed by several changes of distilled water.

4.2.3    Selection of cell culture systems

Many cell culture systems support the growth of polioviruses and other enteroviruses.

Regional reference laboratories (RRL) are advised to obtain cell cultures from the official collections. Requests for these cell lines should be submitted to IVB/VAM, WHO, Geneva.

National poliomyelitis laboratories can in turn apply to their designated RRL for supplies of these cell lines. As soon as possible after the receipt of cell cultures, a cell bank should be established in liquid nitrogen, or if this is not available, in a mechanical freezer at -70°C or lower. Cells stored at -70°C will not remain viable for very long periods and aliquots should be resuscitated every 4–6 months, passaged to build up numbers, and stored again at -70°C.

4.2.4    Preparation of cell culture systems

Cells should be received with documented evidence for the key characteristics relating to the quality of cell cultures as described above. In handling cell cultures, laboratory personnel must be concerned not only with preventing microbial contamination of the cultures, but also with avoiding contamination of the working environment with cell culture materials. All cultures must be considered potentially hazardous, whether inoculated or uninoculated. After use all cultures and their fluids should be decontaminated by autoclaving. Cross-contamination between different cell types, especially continuous cell lines, is an ever-present hazard. To avoid this, different cell lines should never be processed at the same time. All working areas should be thoroughly cleaned between the preparation of different cell types.

Cell culture media employed in virology can be divided into two main categories, growth media and maintenance media.

Growth media (GM), high in serum content (usually 10%), promote rapid cell growth. After a monolayer has formed and prior to inoculation with virus, the growth medium is removed and replaced with maintenance medium.

Maintenance media (MM), low in serum content (usually 2%), are intended to keep the cell cultures in a steady state of slow cell replication whilst maintaining cell metabolism during the period of viral replication.

Fetal calf serum is the serum of choice: it is good for promoting cell growth and it lacks viral inhibitors. If serum from other sources is used, it must be pre-tested for the presence of inhibitors to the viruses being studied. All sera for cell culture use must be inactivated at 56°C for 30 minutes.

4.2.5    Passaging a cell culture

(i) Maintenance of L20B and RD cell cultures

See Section 4.3 for details on preparation of key reagents. Have available the following items:

·         Culture flasks with confluent monolayers of L20B or RD cells (N.B. cultures that have been confluent for longer than two weeks should not be used);

·         Phosphate buffered saline (PBS) without calcium and magnesium;

·         Trypsin (or trypsin/Versene);

·         Growth medium;

·         Trypan blue (if cells are counted);

·         Cell counting chamber;

·         Cell culture tubes and flasks;

·         Pipettes.

(ii) Procedure

·         Examine the cells for quality (i.e. an entire monolayer of healthy cells) and absence of contamination as determined by visual examination.

·         Decant growth medium from the cell culture flask and gently wash the confluent cell layer twice with Ca and Mg free PBS.

·         Add 0.25% trypsin solution (or equal parts of 0.25% trypsin and 1:5000 Versene solution) in PBS to the monolayer, dispersing it evenly. (A volume of 0.5 ml is adequate for a 25 cm2 flask.)

·         Place the flask in a 36°C incubator until the cells detach from the surface: this may be assisted by tapping the side of the flask a few times. Check for complete detachment of cells by examining under an inverted microscope.

·         Re-suspend the cells in growth medium (4.5 ml to a 25 cm2 flask), which halts the action of the trypsin. Gently aspirate the suspension a few times through a fine Pasteur pipette to break up cell clumps.

·         Dilute with growth medium to the desired concentration based either upon counting the cells (see below) or upon a pre-determined split ratio (usually 1:3 or greater). The optimum split ratio (determined by cell counting) required to obtain confluent monolayers of cells in appropriate time must be determined for each new batch of cells received in the laboratory and whenever there are changes to major media components (e.g. fetal calf serum, MEM). The split ratio will quickly become apparent as experience is gained with each culture.

·         Seed fresh culture flasks or tubes, cap tightly, and place in a 36°C incubator.

·         Change tubes to maintenance medium when the monolayer is nearly confluent (2–3 days). Flasks are usually subcultured every 5–7 days, at a split ratio determined by experience.

Table 4.2: Approximate volumes and seeding levels for L20B and RD cell culture

(iii)   Alternative procedure

·         Decant growth medium (GM) from the cell culture flask and gently wash the confluent cell layer with PBS (without the calcium and magnesium components).

·         Add 0.25% trypsin solution (or equal parts of 0.25% trypsin and 1:5000 Versene solution) in PBS, sufficient to cover the cell monolayer.

·         Incubate at 36°C until all the cells detach from the flask (check with inverted microscope).

·         Centrifuge the cell suspension at 100 g for 10 minutes and remove the supernatant.

·         Resuspend cell pellet in GM to desired concentration based either upon counting the cells (use Trypan blue to determine ratio of viable to non-viable cells) or use a 1: 2 to 1: 8 “split” and seed fresh culture vessels/tubes. Change tubes to maintenance medium (MM) when nearly confluent (2-3 days). Flasks are usually subcultured every 5–7 days, a 1: 6 to 1: 8 “split” being typical.

(iv) Cell counting

Accurate numbers in a cell suspension can be calculated by counting the cells in a haemocytometer (e.g. improved Neubauer); it is important to disperse the cells thoroughly by pipetting up and down. A typical method for enumerating cell concentration using “improved Neubauer” haemocytometers is given below.

1)      Dilute 0.2 ml of the cell suspension in 0.2 ml of trypan blue (N.B. use 0.1% w/v trypan blue in PBS solution); non-viable cells are stained blue.

2)      Immediately mix well with a fine Pasteur pipette and aspirate sufficient volume to fill both sides of the haemocytometer chamber.

3)      Count viable cells in each of the four corner squares bordered by triple lines, omitting cells lying on these lines (see Figure 4.2). This is repeated for the second side of the chamber. N.B. cell counts of less than fifty cells are unlikely to be reliable.

4)      If a marked degree of cell “clumping” (aggregation) is observed, discard and re-suspend the original cell suspension.

5)      Calculate the mean count of the total viable cells per four corner squares (N.B. viable cells are not stained by Trypan blue).

6)      Count and calculate the mean count of the other half of the counting chamber. For a valid test, the results of the two counts should be within 20% of the mean value.

7)      Calculate the viable cell concentration per ml using the following formula:

C₁ = t x tb x 1/4 x 10⁴

t = total viable cell count of four corner squares

tb = correction for the trypan blue dilution (counting dilution was 1/tb)

1/4 = correction to give mean cells per corner square

10⁴ = conversion factor for counting chamber

C₁ = initial cell concentration per ml

Example: t = 480; tb = 2; C₁ = 480 x 2 x 1/4 x 10⁴ = 2.4 x 10⁶ cells per ml

8)       Calculate the dilution factor (d) to obtain the working cell concentration per ml (C₂).

d = C₂ (working cell concentration) / C₁ (initial cell concentration)

Example:  C₁ = 2.4 x 10⁶

                  C₂ = 2 x 10⁵

Then: d = C₂ /C₁ =  (2 x 10⁵) / (2.4 x 10⁶) = 2/24 = 1/12
The working concentration can be obtained by mixing 1 volume of the original cell suspension with 11 volumes of the growth medium.

9)       Dispense the cells in growth medium, seed into flasks/tubes and incubate at 36°C. Most continuous cell lines should form confluent monolayers within a few days.

Figure 4.2: Cell counting using a haemocytometer (based on Freshney)

Important variables in these counting chambers are:

a)       the depth of the chamber. In the example above this is 0.1 mm; in some counting chamber types, however, this is 0.2 mm.

b)      the number of smallest squares per cm². In the example above, there are 25 squares per cm², each 0.2 mm long and 0.2 mm wide; in some counting chambers, however, there are 16 squares per cm², each 0.25 mm long and 0.25 mm wide.

When counting chambers with different specifications are used, different algorithms have to be followed for the correct calculation of the number of cells per ml. It is important to check the specifications of the counting chamber in use and follow the calculation instructions that go with individual counting chambers.

(v)       Preservation of cell cultures

It is possible to maintain stocks of cells in a viable state for long periods at low temperatures by the addition of a cryoprotectant such as dimethyl sulfoxide (DMSO) to the cell growth medium. The essential features of the method are to freeze the cells slowly (i.e. at approximately -1°C/min), keep them at a temperature below -70°C while frozen and to thaw them rapidly ready for the preparation of fresh cell culture stocks. Long-term storage can only be achieved reliably when cells are stored at or below -135°C.

4.2.6    Freezing cells

1)      Use only cultures of cells that are in a healthy state (i.e. rapidly growing but not completely confluent).

2)      Detach cells with Trypsin (or Trypsin/Versene — see Section 4.2.4). Use sufficient flasks to yield a minimum of 4 x 106 cells/ml in the final cryoprotectant solution.

3)      Re-suspend cells in growth medium; centrifuge at 100 x g for 10 minutes.

4)      Discard the supernatant and re-suspend thoroughly the cell pellet in pre-chilled growth medium containing 20% fetal calf serum and 10% (v/v) dimethyl sulfoxide.

5)      Dilute 0.1 ml cell suspension in trypan blue and count cells in a haemocytometer as described in Section 4.2.5.

6)      Adjust cell concentration to 4–8 x 106 cells/ml (if large flasks will be used for cell revival) or 2 x 106 cells/ml (if small flasks will be used for cell revival) in growth medium containing DMSO.

7)      Dispense in 1 ml or 2 ml volumes in clearly labelled (cell name, laboratory of origin, passage number and date of freezing) screw-capped, external thread vials (caps should be tightly closed), or polypropylene-sealed/glass-sealed ampoules. The former are suitable for storage in gaseous nitrogen, the latter for storage in liquid nitrogen.

8)      Freeze vials/ampoules slowly. Ideally the temperature should drop at 1°C/minute. Place vials/ampoules in the special container that holds them in the gaseous phase of the liquid nitrogen vessel. Commercial devices are available for which a formula is supplied by the manufacturer for the level vials/ampoules are held, number to be stored and length of time required to achieve this temperature drop (see Figure 4.3). Alternatively, place vials/ampoules wrapped in paper towels or cotton wool in a polystyrene container with a wall thickness of ~25 mm and place this in the -70°C freezer overnight.

9)      Transfer the vials to the gaseous phase (-150°C to -180°C) and polypropylene or glass-sealed ampoules to the liquid nitrogen (-196°C) storage containers (see Figure 4.4). For long-term storage of cells (i.e. a period of years) liquid nitrogen storage is more reliable.

Figure 4.3: Apparatus for controlled cooling of cells (after Freshney)

4.2.7    Thawing cells

Remove vial/ampoule from gaseous/liquid nitrogen and transfer immediately to a water bath or preferably a beaker of sterile water at 36°C.

When contents are completely thawed, wipe outside of vial/ampoule with alcohol to reduce bacterial contamination, transfer cell suspension to culture flask. Add, drop-wise, sufficient growth medium for the production of a cell monolayer (N.B.  If storage vials contain cells at a concentration of 4 x 10⁶ cells/ml, then 1 ml cell suspension should be sufficient for one or two 75 cm² flasks). The viability of the thawed cells may be significantly reduced if growth medium is added rapidly at this delicate stage.

Incubate flask until cells are adherent (6–8 hours) or overnight at 36°C. Carefully decant medium (to get rid of DMSO present) and add fresh growth medium.

As an alternative to the above procedure spin thawed cell suspension (made up to 10 ml slowly with growth medium) at 80 x g for 10 minutes; discard supernatant and re-suspend cell pellet in sufficient growth medium for production of a cell monolayer and incubate at 36°C.

Figure 4.4: Liquid and gaseous nitrogen storage containers (after Freshney)

4.2.8    Establishment of cell banks

The RD and L20B continuous cell lines recommended for use by all virus laboratories studying poliomyelitis have been well characterized as regards species identity and lack of contamination by bacteria, mycoplasma and infectious virus.

Regional reference laboratories (RRL) are advised to obtain cell cultures from the official collections. Requests for these cell lines should be submitted to IVB/VAM, WHO, Geneva. A number of RRLs have been requested to establish a master cell bank (MCB) for each cell line. Using scrupulous laboratory techniques of cell culture passage and storage, these RRLs can supply cell cultures directly to the National Laboratories that come within their responsibility. National Laboratories are strongly encouraged to store supplies of these cells, in gaseous or liquid nitrogen, as the working cell bank (WCB) for their own use. Figure 4.5 outlines the formation of the MCB and WCB.

Figure 4.5: Establishment of a cell bank

The advantages of initiating a cell bank are:

·         The expense of transport of original cell lines from official sources is minimized.

·         The RRL acts as the MCB repository of well-controlled cell culture stocks.

·         The National Laboratory is nearer a supply of good viable cell cultures held at the MCB, but is also encouraged to be self-supporting by producing and storing cells in its own bank (WCB).