Cell Biology Protocols

68 CELL CULTURE

Capillary action will draw the cell suspension under the coverslip, filling the area between the central and side gullies. Repeat this on the other side of the coverslip. (Note: If the coverslip is not positioned correctly both chambers and the gully either side of the chambers will flood with cell suspension and accurate cell counting will not be possible.)

•Place the chamber on the microscope platform and move it around until you have located the central grid of 25 small squares bounded by three parallel lines. Count the total number of cells (stained and unstained) in these 25 squares. (They are each subdivided into 16 smaller squares to facilitate counting.) In order to avoid counting the same cell twice, count the cells on the top line, the left-hand line and the centre of each small square bounded by triple lines. Use a tally counter if the numbers are large. (Note 1: The area counted

depends upon the cell density. If the cell number reaches 100 in the central square this is sufficient for accuracy. If the cell density is much lower, count the total number of cells in the four corners and take an average. For further accuracy it is advisable to repeat the total cell count and take an average of the two readings. Note 2: If you are using phase contrast on the inverted microscope, live and dead cells can be distinguished without staining. Live cells are brightly refractile with a distinct outline; dead cells are flat and grey with an uneven edge.)

•When you have counted the total number of cells, repeat the counts in the same areas, counting only the blue-

stained (dead) cells. Calculate the average number of dead cells and subtract from the total number of cells to obtain the number of live cells.

(Note: Accuracy can be confirmed by

repeating the count on unstained cells only.)

•The viability of the cell population is calculated using the formula:

number of live cells

Viability =

number of live + dead cells

×100%

•The concentration of cells in the cell suspension is calculated as follows:

Average number of cells in one large square (25 small squares) × 104 = no. of cells/ml

(Note: This is either the number of cells in the central large square or the mean of the counts in the four large corner squares.)

•The total number of cells in the cell suspension = no. of cells/ml × total volume.

•The total number of live cells in the suspension = total number of cells × % viability (Note: If you are calculating the number of cells from the trypan blue stained population, remember to allow for the twofold dilution factor introduced by diluting the cell suspension in the trypan blue solution.)

•Example: (see Figure3.1)

Volume of cell suspension = 20 ml

Total cell count of live cells in 4 corner squares = 160

Total cell count of live and dead cells in 4 corner squares = 200

Mean cell count of live cells in one large square = 40

Mean cell count of live and dead cells in one large square = 50

Viability of cell population

= 40/50 × 100%

= 80%

Concentration of viable cells

=50 × 104 × 80 cells/ml

100

=40 × 104 viable cells/ml

=4 × 105 viable cells/ml

Multiply this ×2 to allow for dilution factor = 8 × 105 viable cells/ml

Total number of cells in original cell suspension = 8 × 105 × 20

= 16 × 106cells

To adjust the cell number to 106/ml, centrifuge the cell suspension at 1000 rpm for 5 min. Pour off the supernatant and resuspend the cells in 16 ml of medium. Confirm the cell concentration by recounting the number of cells.

To adjust the cell number to 1.5 × 105 /ml:

Original cell count = 8 × 105/ml, dilution factor = 8/1.5 = 5.3

Therefore dilute 1 ml of 8 × 105/ml to 5.3 ml.

Use of cell cultures – Anne Wilson

Introduction

A wide range of cell lines representing diverse cell types and species are available commercially from national collections in Europe and America (ECACC, ATCC, DSmZ). In theory, cells grown in culture provide a continuous, replicate supply of material of a homogeneous characterized cell population. However, precautions need

70 CELL CULTURE

to be taken to ensure that the latter is true in practice. Cross-contamination of cell lines can occur during culture and the line may not therefore be as described [27, 28]. Particular attention needs to be paid to this possibility if lines are exchanged between laboratories rather than obtained from commercial sources. Isoenzymes, karyotypes and DNA fingerprinting are all appropriate tools for speciating cell lines [29–31]. A wide range of antibodies are also available for the immunocytochemical identification of phenotypic characteristics such as cytoskeletal proteins, surface markers and biosynthetic products.

Cell lines in long-term culture can show drifts in both phenotype and genotype. When using cell lines for a series of related experiments it is advisable to use cells from similar passage numbers. Generally these are within 10 passages and once the stock has passed this number fresh working stock should be thawed out for use. Guidelines for handling cancer cell lines have been published by UKCCR [32].

Continuous cultures of cell lines may become contaminated with mycoplasma. The presence of these micro-organisms affects a number of cell properties. Surface changes may alter the antigenicity of cells and their presence results in competition for a variety of metabolites needed for DNA synthesis, protein synthesis and other essential metabolic pathways. Signs of contamination include a decrease in growth rate, increased acidity of medium, deterioration of cells and the appearance of a grainy background. There are several different species found in culture and assay for the detection of mycoplasma should be a regular routine during maintenance of cell lines in long-term culture. A number of methods are available for detecting mycoplasma in culture, the simplest being the use of Hoechst 33258, a fluorescent stain which binds to DNA and reveals the presence of extracellular and extranuclear DNA [33]. A variety of commercially available kits can also be obtained [34].

Equipment

All equipment coming into direct contact with cells must be sterile.

Class II cabinet

Incubator set at 37 ◦C

Tissue culture flasks

Universal containers (conical-bottomed) with labels showing volume gradations

Pastettes (long-form and short-form)

Inverted phase contrast microscope with ×10 objective

Water bath set at 37 ◦C

Waste bottle for spent culture medium

Permanent black marker pen

Reagents

All reagents must be sterile.

Hanks balanced salt solution without calcium and magnesium (HBSS)

Culture medium appropriate to cell type

Trypsin/versene solution (1–2 ml/25 cm2)

(Note 1: If this has been purchased as a 10× strength solution, thaw out stock, dilute to 1× strength in HBSS and dispense 5 ml aliquots into sterile bijoux. Store at −20 ◦C.)

Procedure

•Switch on the Class II cabinet 10 min before starting and swab down its surfaces with 70% methanol/30% water.

Any equipment to be used in the cabinet should be placed towards the back so that air flow is not obstructed and sterility is maintained.

• Warm the reagents to be used to 37 ◦C.

•Remove the flasks for harvesting from the incubator and check them for health under the microscope.

•Decant the spent medium into a sterile

bottle for waste medium and rinse the monolayer gently with 3 × 5 ml washes of HBSS, pouring the wash medium from the flask into the waste bottle after each rinse. If the medium dribbles down the neck of the flask use a tissue soaked in 70% alcohol/30% water to wipe it off.

•Add 1–2 ml of trypsin-versene to the flask, re-cap it and tilt the flask horizontally in several directions to ensure the entire cell monolayer is bathed in enzyme solution.

•Place the flask horizontally into the incubator for 2–5 min.

•Remove it and check visually for cell detachment. If the cells have become rounded and are detaching from the cell surface this will be visible to the naked eye. If the cells are still attached return the flask to the incubator for several more minutes and check again for detaching cells. (Note: Cell lines vary in the speed with which they detach; times may vary from 1 to 20 min. Harvesting procedure is usually specified in details accompanying purchased cell lines.)

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•Once the cells have begun to detach add5–10 ml of culture medium containing serum to the flask. Re-cap it and tap the side of the flask against the palm of your hand several times; the shearing forces cause further detachment of the cells. Use the microscope to check for complete detachment of cells.

•Use a pastette or pipette to dispense the detached cells into a universal. Rinse the surface of the flask with another 5 ml of medium and transfer the remainder of the detached cells into the universal container. Check the flask under the microscope to make sure that all the cells have detached and been recovered.

and viability determination (see Protocol 3.8).

Notes

1 Familiarize yourself with the normal appearance of the cells in the different stages of culture so that you notice abnormalities. These may be due to a number of factors including: pH too acid or too alkaline, nutrient deficiency due to infrequent media changes, cells seeded at too low a density in the starting culture, cells left in stationary phase for too long, contamination by micro-organisms (bacteria, yeast, fungi, mycoplasma), crosscontamination with another cell line, or natural drift in phenotype and/or genotype.

2 Cells should be harvested at the same time point in the life of the culture, ideally when they are reaching the end of the exponential growth phase and before the culture has become stationary, though this may be

dependent upon the state of the cells required for subfractionation.

3 Take the following precautions to avoid cross-contamination between

solutions, pipettes and waste medium disposal bottles for each cell line.

(c) Never put a pipette that has been in contact with cells back into a medium or reagent bottle. (d) Clean the cabinet before handling the next cell line. (e) Handle fast-growing cell lines (e.g. HeLa) that grow from a small inoculum of cells last.)

Cryopreservation – Anne Wilson

Stocks of cells can be kept for long periods by freezing them down and storing them in liquid nitrogen.

Critical factors in the successful freezing of cells include the rate of freezing, the type and concentration of cryoprotectant and the cell concentration [35–39]. Although optimal results are obtained using a programmable cell freezer it is possible to control the rate of freezing in other ways. These include the use of a thickwalled polystyrene box stored in a −80 ◦C freezer or a special device designed to fit into the neck of a liquid nitrogen cell storage bank, thus holding the cells in vapour phase and allowing them to cool down slowly. These methods do not give such a high viability on recovery of cells from frozen stock as does the programmable freezer. Avoidance of ice crystal formation within the cell is critical and this is minimized in several ways, including incorporation of a cryoprotectant such as dimethylsulfoxide (DMSO) or glycerol, an increased serum concentration, a high cell number and a controlled slow rate of freezing. For most mammalian cells, the optimum rate is −1 ◦C/min. DMSO at 10% is the most commonly used cryoprotectant

and the protocol outlined here describes its use. However, it may be toxic to some cells. (Note: DMSO is a polar solvent and can penetrate the human skin, together with any substances dissolved in it. Take care when handling it and wear protective gloves.)

The method of recovery of cells is also critical and should aim to avoid osmotic shock. Frozen cells need to be thawed quickly to 37 ◦C and then diluted very slowly in culture medium.

PROTOCOL 3.10

Freezing cells

Equipment

All equipment coming into direct contact with cells must be sterile.

Plastic freezing vials (2 ml)

Pastettes (long-form and short-form)

Universal containers (conical-bottomed) with labels showing volume gradations

Black permanent marker pen

Waste bottle for medium

Improved Neubauer haemocytometer chamber with coverslip

Bench centrifuge

Inverted phase contrast microscope with ×10 objective

Programmable cell freezer or Thick-walled polystyrene box or

Device for neck of liquid nitrogen cell bank or storage vessel

Dewar flask with vented lid suitable for the storage of liquid nitrogen

Liquid nitrogen cell bank

Storage vessel for liquid nitrogen

Trays or straws for storing ampoules in cell bank

Face protection

Insulated protective gloves

Long forceps

Micropipette (1–5 ml)

Tips for micropipette

Reagents

All reagents must be sterile.

Hanks balanced salt solution without calcium and magnesium (HBSS)

Dimethylsulfoxide (DMSO)

Culture medium containing 10–20% serum

1% Trypan blue

Liquid nitrogen in Dewar flask

Procedure

•Determine the viability and cell count of the population for freezing using

Protocol 3.8.

•Calculate the volume of diluent needed to resuspend the cells at 5 × 106/ml.

•Prepare this volume of a 10% solution of DMSO in culture medium and mix well.

•Label the appropriate number of freezing vials clearly with the date, pass number and cell type using the black marker pen.

•Centrifuge the cell suspension at 1000 rpm for 5 min, discard the supernatant and resuspend the cells in the required volume of 10% DMSO. Mix well.

•Dispense the cell suspension as 1.8 ml aliquots into the freezer tubes using a micropipette and cap them securely.

•For programmable freezing the vials can be transferred directly to the freezer and the freezing cycle started (see manufacturer’s instructions).

•Transfer the vials to the long-term storage facility when freezing is complete.

•For freezing with a device for the neck of the cell bank or storage vessel, try the

following as a starting point: Keep the filled freezing vials at 4 ◦C for 60 min, transfer to the freezing device and keep in vapour phase overnight, then transfer them to the long-term storage facility.

•For freezing with a polystyrene box, try the following as a starting point:

Keep the filled freezing vials at 4 ◦C for 60 min, transfer to the box and hold the box at −20 ◦C for 60 min, transfer the box to −80 ◦C overnight, then transfer them to the long-term storage facility.

(Note: Frozen cells can be stored at −80 ◦C for several months.)

PROTOCOL 3.11

Thawing cells

Equipment

All equipment coming into direct contact with cells must be sterile.

Water bath set at 37 ◦C

Protective face mask

Insulated protective gloves

Long forceps

Universal containers (conical-bottomed) with labels showing volume gradations

Watch with second hand

Pastettes (long-form and short-form)

Bench centrifuge

Dewar flask for storage of liquid nitrogen

Reagents

All reagents must be sterile.

Culture medium warmed to 37 ◦C

Liquid nitrogen in Dewar flask

70% methanol/30% water

Procedure

•Half-fill the Dewar flask with liquid nitrogen.

•Remove vials from the nitrogen cell bank and transfer to the Dewar flask containing liquid nitrogen for transport to the lab. (Note: There is a risk of vials exploding when removed from the nitrogen bank. Protective face mask and gloves must be worn until the vial has been completely thawed.)

•Remove one freezing vial from the flask using forceps.

•Holding the vial at arm’s length, plunge it into the water bath and agitate the vial

in the water to thaw it quickly and bring it up to 37 ◦C.

•Immerse the thawed vial in 70% methanol/30% water to sterilize it and wipe dry with a tissue. (Note: Make sure you have previously noted the details written on the side of the vial in case they get washed off.)

•Remove the lid of the vial in the Class II cabinet and transfer the thawed contents of the vial into a universal container, using a pastette.

•Take up some warmed culture medium

into the pastette and add one drop ( 100 µl) to the cell suspension. Mix it gently and add another drop 15 s later.

Continue until the volume has been doubled. Then add 500 µl every 15 s with gentle mixing until the volume has doubled again. Complete the dilution

by adding 1000 µl every 15 s up to

16 ml. (Note: The rate of addition of medium is gradually increased as the volume increases because the risk of osmotic shock decreases.)

•When dilution is complete, centrifuge

medium. Mix gently and centrifuge again at 1000 rpm for 5 min. Repeat this step to wash off residual DMSO.