Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
Applied Veterinary Virology:
The isolation and identification of viruses using cell
cultures
Authors: Prof Estelle Venter
Licensed under a Creative Commons Attribution license.
CELL CULTURE PROCEDURES
Cell culture media
Water: Water represents more than 99% of the material that comes into contact with cells whether as
a medium ingredient, in the processing operation or as the final rinse of containers in which the
cultures are grown.
Purification methods that can be used are:
Ion exchange: This involves the use of ion exchange resins for the removal of ionic impurities from
water.
Reverse osmosis: If a semi-permeable membrane, permeable to the solvent but not to the solute, is
placed at an interface between two solutions of different solute concentrations, then solute transport is
prohibited and a free energy difference due to the concentration gradient develops across the
membrane. Under these circumstances solvent moves through the membrane from the low- to the
high-concentration compartment, either until the concentration difference is zero, or until the
hydrostatic pressure difference reaches equilibrium with the free energy driving force. The process is
known as “osmosis” and the hydrostatic pressure difference which stops the net transfer of solvent is
the osmotic pressure. Because it is in a thermodynamic equilibrium, if the pressure applied to the
high-concentration side of the membrane is increased above the osmotic pressure, solvent molecules
are driven against the concentration gradient. This process is termed “reverse osmosis”.
Theoretically, reverse osmosis allows only water molecules to traverse the membranes, although in
practice, polar ions up to a molecular weight of 100 can transfer. Molecular weights of 1000 or more
are totally excluded, meaning that “reverse osmosis” water is invariably pyrogen-free.
“Reverse osmosis” is the most suitable and economic primary system for the preparation of media
and cleaning of equipment and glassware. Water purified by “reverse osmosis” has a very low organic
content and a small ionic content. Normally a “reverse osmosis“ unit is linked to a water mains by a
system incorporating a sealed storage tank (to which is attached a hydrophobic vent filter), a
circulating pump, an ion exchange column, an in-line 0,45 μm filter cartridge (to protect an in-line UV
system) and a 0,2 μm filter cartridge upstream of the purified water outlets.
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
Distillation: This is the only method for producing water for critical pharmaceutical purposes.
Distillation, although producing the highest quality water, is the most expensive method of treatment
because of high capital and operational costs, and for that reason it is only appropriate for purifying
water for the most critical applications.
Basic salt media
The culture medium is by far the most important single factor in culturing animal cells. The function of
this medium is to provide an environment for survival and also to provide substances required by the
cells which they cannot directly synthesize. The composition of early tissue culture media was based
on biological fluids such as plasma, lymph and serum, and tissue extracts especially of embryonic
origin. Basal tissue culture media were developed to include only the minimal components which were
essential for growth.
Types of basal media:
There are four main categories of basal media for mammalian cells and several categories for
insect cells. These are:

Eagle’s medium and derivatives, e.g. BME (basal medium Eagle’s), EMEM (minimum
essential medium with Earle’s salts), AMEM (alpha modification of MEM), DMEM
(Dulbecco’s modified Eagle’s medium), GMEM (Glasgow modification of Earle’s medium),
and JMEM (Joklik’s modification of MEM)

Media designed at Roswell Park Memorial Institute (RPMI), e.g. RPMI 1629, RPMI 1630,
and RPMI 1640

Basal media designed for use after serum supplementation, e.g. Fischer’s, Liebovitz,
Trowell, and Williams’

Basal medium designed for serum-free formulations, e.g. CMRL 1060, Ham’s F10 and
derivatives, TC 199 and derivatives, MCDB and derivatives, NCTC and Waymouth
For insect cell cultures, the basal media are:

Grace’s medium

Schneider’s medium

Mitsuhashi and Maramorosch medium

IPL-41 medium


Chiu and Black medium
D-22 medium
Constituents of basal media
Allowance is usually made for differences in salt concentration to obtain the optimum
osmolality. Different incubation temperatures may also require adjustment to be made to the
composition of buffering components, since pH may change with temperature due to alterations
in the solubility of CO2 and in ionization and pKa of buffers. The following may be considered:

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Balanced salt solution
Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
Most of these solutions do not include nutrients required for most cells for long-term
maintenance or growth although glucose may be included

Earle’s balanced salt solution

Dulbecco’s phosphate-buffered saline


Hank’s balanced salt solution
Eagle’s spinner salt solution
Dulbecco’s phosphate-buffered saline and Hank’s balanced salt solution are intended for use
equilibrated with air, while Earle’s balanced salt solution and Eagle’s spinner salt solution
require equilibration with a gas phase containing 5% CO 2 in order to maintain the correct pH.

Buffering systems: Phosphate buffering systems are normally used – see discussion
below

Energy source: Carbohydrates are a major energy source for cultured cells. Glucose is the
most frequent used sugar. Other sugars, e.g. maltose, sucrose, fructose, galactose, and
mannose, may also be included. Glutamine can also supply a major proportion of the
required energy in some cells

Amino acids: Most animal cells have a high requirement for glutamine. Glutamine acts
both as an energy source and as a carbon source in the synthesis of nucleic acids. Other
amino acids are often added to compensate either for a particular cell type’s incapacity to
make them or because they are made but lost in the medium

Vitamins: Several vitamins of the B group are necessary for cell growth and multiplication.
Many vitamins are precursors for cofactors and must be added

Hormones and growth factors: These are included in some media (especially serum free
media) at relatively low concentrations

Proteins and peptides. Although an absolute requirement for proteins and/or peptides by
cells in culture has not been established, relatively few media have been formulated in
which cells grow rapidly in the total absence of proteins or peptides

Fatty acids and lipids: There is no consensus regarding an essential role for lipids in cell
culture

Accessory factors: Amongst these are the ‘trace’ elements, especially iron, zinc, copper,

and selenium
Antibiotics
Buffering systems
A phosphate buffer system is normally used provided by sodium and potassium phosphates,
sodium bicarbonate, other natural ingredients such as sera, proteins, amino acids, as well as
synthetic, organic molecules, such as HEPES (N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic
acid),
TRIS
(tris-hydroxymethyl-aminomethane),
TRICINE
(N-tris-(hydroxymethyl)-
methylglycine), BES, (N,N’-bis-(2-hydroxy-ethyl)-2-aminoethanesulfonic acid) and others. Three
or more of these are simultaneously used to control the pH in most cell culture media.
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
The use of sodium bicarbonate, with or without CO 2 added to the gas phase of the atmosphere
in which the cells are grown, significantly improves the buffering situation. The use of sodium
bicarbonate, with or without CO2 added to the gas phase of the atmosphere in which the cells
are grown, significantly improves the buffering situation. The interrelationship between the
bicarbonate ion concentration in the medium, the pH and the partial pressure of carbon dioxide
in the gas phase is important. Carbon dioxide gas is very soluble in culture media. A small
proportion (less than 1%) combines chemically with water to form the weak carbonic acid which
dissociates to yield hydrogen and bicarbonate ions.
H2O + CO2  H2CO3  H+ + HCO3Thus the acidity will gradually increase as the partial pressure of carbon dioxide is increased.
Since sodium bicarbonate is a salt, it will dissociate virtually completely. The concentration of
bicarbonate ion will be very close to that of the salt added. Additions from dissociation of
carbonic acid or losses due to dissociation of bicarbonate to carbonate are negligible. Thus for
a given temperature, one can predict the initial pH if the bicarbonate ion concentration and
partial pressure of carbon dioxide are fixed.
The levels of CO2 in the atmosphere in which cell culture systems are grown vary from that of
ambient air (i.e. about 0,03%) up to 40%. Generally, either no CO2 is added or, alternatively,
5% to 10% CO2 is added to the atmosphere.
Prophylactic use of antibiotics
The prophylactic use of antibiotics in cells and tissues with a high risk of microbial
contamination and in conjunction with aseptic techniques can considerably reduce the
overgrowth of bacteria (including mycoplasmas) and fungi.
The majority of antibiotics used for controlling microbial contamination of cell or tissue cultures
are effective against both Gram-positive and Gram-negative bacteria. Gentamycin and
ciprofloxacin exhibit activity against mycoplasmas, and the polyenes, amphotericin B and
nystatin are useful in preventing contamination with yeasts and fungi (Wiedemann and Atkinson
1991).
The combination of penicillin G at a concentration of 105 U/ℓ, streptomycin sulphate (100 mg/ℓ)
has found wide application in preventing bacterial and fungal contamination in cell and tissue
cultures. More recently gentamicin sulphate at a concentration of 50 mg/ℓ has been used as an
alternative to streptomycin sulphate because of its activity against Pseudomonas species. This
combination has been particularly useful in reducing the incidence of bacterial contamination in
cell cultures used for isolating viruses from clinical samples. However, the use of gentamicin is
not to be encouraged.
Several factors should be considered when determining the most appropriate antibiotic or
combination of antibiotics for use in cell or tissue culture. The antibiotic’s cytotoxicity and
stability in cell culture media and its spectrum of antimicrobial activity should be determined.
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
Also, the types of contaminants that may be encountered and interactions with other antibiotics
should be considered.
A list of antibiotics and their use can be found in Paul (1975), Schaffner (1979) and Gray &
Brenwald (1991).
One should always test the toxicity of the antibiotic to cell cultures before using the antibiotic.
Serum
Foetal bovine serum (FBS) is now most frequently used at 10% concentration although this
may be changed for specific applications. Advantages of serum use include the following:
•
Serum represents a cocktail of most of the factors required for cell proliferation and
maintenance
•
Serum is an almost universal growth supplement which is effective for most cells. Using
serum-supplemented medium therefore reduces the need to spend time developing a
specific, optimized medium formulation for every cell type under investigation
•
Serum buffers the cell culture system against a variety of perturbations and toxic effects,
such as those due to pH change, or presence of ions of heavy metals, proteolytic activity,
or endotoxin
The use of serum also imposes a number of difficulties which impact on the safety,
reproducibility, and cost of biopharmaceuticals produced in animal cells. These difficulties can
be minimized by careful selection and validation of serum sources. Although almost all new
manufacturing processes using animal cells are designed for serum-free media in order to
avoid these difficulties, many existing processes still use FBS-supplemented medium. This
situation is unlikely to change fundamentally in the near future since regulatory constraints
generally make it impractical and uneconomical to alter existing processes.
Potential problems with the use of serum
There are a number of serious disadvantages incurred when serum is used to supplement a
culture medium:
•
Lack of reproducibility. Serum batches vary considerably depending on such factors as the
characteristic of the source animals used, the feed stuffs employed, and on the time of
year. Different batches contain different absolute and relative levels of growth factors.
Certain factors may be deficient in some batches, while others may be present at
excessive, inhibitory levels for some cell types
•
Risk of contamination. Serum can represent a major route for the introduction into cultures
of adventitious agents including bacteria, fungi, mycoplasmas, and viruses. Health checks
on donor animals should be applied and the facilities used for the collection and
processing of serum must be checked. Serum, after filtration, is checked for the presence
of viruses. A standard quality control procedure should be applied
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
•
Influence on downstream processing. This is a problem in protein studies when proteins
produced by the cell culture need to be harvested and purified. Serum proteins added may
inhibit or influence this process because of their high concentration compared to the
proteins of interest
•
Availability and cost. Correctly collected, processed, and validated FBS will always
contribute greatly to the cost of a tissue culture laboratory
Serum storage and use
Serum is rapidly frozen by the supplier immediately after bottling and is thereafter held at –
20°C. Few data are available on the shelf life of serum held in this way, but 2 years has
become accepted as a rule of thumb. However, one study indicated that 2 years at –20°C is a
conservative estimate of shelf-life and that this could be extended to 5 years. When required,
serum should be thawed rapidly during which process it is gently mixed to minimize protein
denaturing due to salt concentration effects. A water bath at 37°C is best although the serum
should be removed as soon as fully thawed and not allowed to warm up. Thawed serum should
be clear and there should be no significant precipitation. Once thawed, it can be held at 4°C for
a maximum of 2 - 3 weeks. Serum should not be re-frozen.
Sterilization of cell culture media
Sterilization of media is general done by filtration.
Tissue culture technique
Prevention of contamination by any extraneous organisms (bacteria, yeast, fungi, mycoplasmas,
viruses) is a prerequisite in tissue culture. The basic principles of good technique are essentially
common sense, but it is worth restating the obvious to ensure that the first principles are correct.
Blanket antibiotic cover is to be avoided at all costs. Antibiotics have their place in case of
“emergency” or in the early stages of primary isolation of cells from contaminated tissues. However,
routine use of them in tissue cultures cannot be justified and should not be a “cover” for poor
laboratory practices. Indiscriminate antibiotic use will inevitably lead to problems. Hidden infections
can occur, which only become apparent during experimental or production procedures, and may
render the cell line unusable for the particular purpose envisaged.
Recommended procedures for handling cell lines (valid for all types of facility)

Avoid multiple-use facilities, if possible

Only handle one cell line at a time. Such a common-sense approach should cut to the
minimum the possibility of miss-labelling, mix-ups, and cross-contamination between cell
lines

Examine cultures daily for evidence of cross-contamination

Maintain separate bottles of media (and preferably other reagents) for each cell line

Institute quality control procedures for all the reagents used in tissue culture even if
purchased from a commercial supplier
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures

Keep the working areas uncluttered (consumable supplies should be kept in racks below
the work area), swab with a suitable disinfectant between handling cell lines, and allow a
minimum of time (15 min) between the handling of different cell lines

Dispose of all used materials safely, efficiently and routinely

If separate work areas are not available, at least designate separate incubators for
different types of materials and carry out procedures done on a daily basis with quality
controlled cultures before others

Routine quality control procedures should be carried out more frequently in multi-use
facilities

Ensure that laboratory coats are regularly changed

Wash hands thoroughly before and after commencing laboratory work. If possible, use
sterile gloves

Avoid the generation of aerosols (e.g. by too vigorous pipetting) in the work area

Inside the safety cabinet liberal use of a disinfectant e.g. 70% ethanol) to swab work
surfaces/bottles/flasks is recommended

Train and monitor junior staff in tissue culture techniques before allowing them to handle
particularly valuable material

Maintain facilities efficiently

Ensure that all incubators, microscopes, centrifuges and other equipment are clean and
serviced regularly

Discard unused cultures from incubators as soon as they are no longer required

If laboratory topography and finance permit, install facilities which can be routinely
fumigated
Information used to describe a cell line or strain for publication

History

Population doubling number (subculture number)

Culture medium used

Growth characteristics

Absolute plating efficiency

Morphology

Frequency of cells with various chromosome numbers in a culture

Karyotype characteristic of the stem line

Whether sterility tests for mycoplasmas, bacteria and fungi have been done

Whether the species of origin of the culture has been confirmed

A description of a cell strain should also include the procedure of isolation, the number of
population doublings, and the length of time since isolation.
Strategies for the derivation of primary cultures and establishment of cell lines
Primary cultures are generally derived directly from a particular organ or tissue, although the
use of whole embryos (e.g. chick) is common practice for some applications, e.g. virology. To
achieve a successful primary culture depends on many factors, such as the species and the
age of donor, but a major factor is the choice of tissue. Materials from mature or adult
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
organisms are less reliable as a source of actively dividing cells than embryonic tissues. One
notable exception is tumour tissue; but, this gives variable success. Some cell types, such as
plasma cells, are the end product of a long sequence of differentiation events and are therefore
unlikely to survive in culture. For this reason, embryonic tissue is a recommended source
despite the fact that they may have different characteristics from adult cells, and may not ever
mature.
Once the primary culture has been established, successful subculture to generate a cell line is
usual, and allows the possibility of cloning, characterization and preservation, and thus greater
uniformity. It should be remembered that cells from a primary vertebrate culture usually have a
finite lifespan dependent on the species. In order to overcome this, and generate an immortal
cell line, a transformation event must occur, either as a natural somatic mutation or as an
induced event (in vivo or in vitro).
The method of tissue dissociation will be dependent upon the type of tissue and the long-term
purpose of the culture. In some tissues, like spleen, the cells are loosly associated and mild
physical shearing techniques will release them from the tissue. Mechanical techniques can
cause significant cell damage. In many cases enzymatic digestion is an absolute necessity.
Unfortunately most enzymes have severe deleterious effects on cells. These may be reversible
or they may cause lysis. The lack of success in deriving cultures from adult tissue may, at least
in part, be due to the extreme conditions necessary for dissociation of cells from the ageing
tissue matrix. The criteria for choice of enzyme and dissociation conditions will need to be
established by trial and error though previously published data may provide guidance.
During the process of adaptation to tissue culture conditions, the cell undergoes a significant
loss of many biochemical and morphological characteristics associated with the tissue of origin
(Griffiths and Riley 1985). The extent of this de-differentiation will depend on the cell type and
the suitability of the culture conditions. Nevertheless, primary cultures are considered more
representative than some of the ubiquitous cell lines, in particular in the expression of tissuespecific properties.
Tumour tissue is an extremely useful source of cells for primary cultures, especially if an
immortalized cell line is required. Such cells tend to lack many differentiated characteristics
and, even as primary cultures, display in vitro tumourgenic properties, such as loss of contact
inhibition of growth (Griffiths and Riley 1985).
It must be borne in mind that most primary cultures are made up of a heterogeneous cell
population. However, by careful selection and isolation, it is possible to obtain a homogenous
population. Even so, sub-culture will almost certainly lead to loss of more specialized cells and
with them differentiated properties unless care is taken to select out the correct lineage. An
essential feature of this developmental stage is to identify accurately the characteristics of the
cell population of interest by all available techniques.
Enzymatic techniques for tissue dissociation
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
The procedure used to isolate cells from tissues must be designed to maximize the yield of
functionally viable and dissociated cells remaining after digestion.
Before preparing cell cultures the following list of parameters should be addressed so that a
decision on the suitability of the prepared material can be made.
Type of tissue, species of origin, age of the animal, dissociation medium used, enzyme(s) used,
impurities in any crude enzyme preparation used, concentration(s) of enzyme(s) used,
temperature and incubation times.
Enzymes which may be used
•
Collagenase
This enzyme, more accurately referred to as purified clostridiopeptidase A, is usually inefficient
in dissociating tissue and the collagenase most commonly used for tissue dissociation is a
crude preparation containing clostridiopeptidase A in addition to a number of other proteases,
polysaccharidases and lipases.
Four basic types of collagenase are now identified and available commercially:
Type 1 contains average amounts of assayed activities (collagenase, caseinase, clostripain,
and tryptic activities). It is generally recommended for obtaining fat cells, adrenal cells and liver
cells.
Type 2 contains greater clostripain activity. It is generally used for heart, bone, muscle, thyroid,
cartilage and liver tissue.
Type 3 is selected for low proteolytic activity. It is usually used for obtaining mammary cells.
Type 4 is selected because of low tryptic activity. It is commonly used for obtaining the cells of
the islets of Langerhans from pancreatic tissue and other such applications where receptor
integrity is crucial.
•
Trypsin
Trypsin is a pancreatic serine protease with specificity for peptide bonds involving the carboxyl
group of the basic amino acids arginine and lysine. It is one of the most highly specific
proteases known, although it also exhibits some esterase and amidase activity.
Purified trypsin alone is usually ineffective for tissue dissociation since it shows little selectivity
for extracellular proteins. Combinations of purified trypsin and other enzymes, such as
elastase, have proven effective for dissociation of the cells of many tissues.
‘Trypsin’ is also the name commercial suppliers have given to pancreatin, a crude mixture of
proteases, polysaccharidases, nucleases and lipases extracted from porcine pancreas. Crude
‘trypsins’ (National Formulary (NF) 1:250 or 1:300), are widely used for dissociating tissues,
perhaps because the tryptic and contaminating proteolytic and polysaccharidase activities do
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Applied Veterinary Virology: The isolation and identification of viruses using cell cultures
bring about a preferential attack of the extracellular matrix. It appears, however, that crude
trypsin and crude collagenase dissociate tissues by different mechanisms, and difficulties are
often encountered when using NF 1:250 preparations - the most common being incomplete
solubility, lot-to-lot variability, and cell toxicity.
In tissue culture laboratories, purified trypsin is used to release cells into suspension from
monolayers growing on the interior surfaces of culture vessels.
•
Elastase
Pancreatic elastase is a serine protease with a specificity for peptide bonds adjacent to neural
amino acids. It also exhibits esterase and amidase activity. While elastase will hydrolyze a wide
variety of protein substrates, it is unique among proteases in its ability to hydrolyze native
elastin, a substrate not attacked by trypsin, chemotrypsin or pepsin. It is produced in the
pancreas as an inactive zymogen, proelastase, and activated in the duodenum by trypsin.
Elastase is also found in blood components and certain bacterial species.
Because elastin is found in highest concentrations in the elastic fibres of connective tissues,
elastase is frequently used to dissociate tissues which contain extensive intercellular fibre
networks. For this purpose, it is usually used with other enzymes, such as collagenase, trypsin
and chymotrypsin. Elastase is the enzyme of choice for the isolation of Type II cells from the
lung.
•
Hyaluronidase
Hyaluronidase is a polysaccharidase with as specificity for bonds between 2-acetoamido2deoxy-ß-D-glucose and D-glucoronate. It is often used for the dissociation of tissues, usually
in combination with crude protease, such as collagenase.
•
Papain
Papain is a sulfhydryl protease from the latex of the paw-paw or papaja tree, Carica papaya.
Papain has wide specificity and it will degrade most protein substrates more extensively than
pancreatic proteases. It also exhibits esterase activity. In some tissues, papain has proved less
damaging and more effective than other proteases.
•
Deoxyribonuclease 1
Often, as a result of cell damage, nucleic acid leaks into the dissociation medium, increasing
viscosity and causing handling problems. Deoxyribonuclease will digest the nucleic acids
without damaging the intact cells.
•
Pronase
This protease is a bacterial enzyme derived from Streptomyces frisens. In addition to gentle
disaggregation of tissues, it is often used as an alternative to trypsin for sub-culturing cell lines.
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Because of its mild action, pronase may have to be used in conjunction with other enzymes,
e.g. collagenase.
•
Dispase
This is a bacterial enzyme, obtained from Bacillus polymyxa. It is a neutral metalloenzyme
requiring calcium for activity. It is useful for tissues of low hyaluronic acid content and is usually
used in conjunction with other enzymes for complete digestion.
•
EDTA
Ethylenediaminotetraacetic acid at a concentration of 0,02 % (w/v) in calcium- and magnesiumfree phosphate-buffered saline (PBS) can be used in conjunction with trypsin to enhance tissue
disaggregation. Acting as a chelating agent, it dissociates intercellular links, such as tight
junctions. For this reason it is not suitable for use with dispase.
•
Trypsin inhibitor (soybean)
The trypsin inhibitor from soybean inactivates trypsin, but it exhibits no effects on the
esterolytic, proteolytic or elastolytic activities of porcine elastase.
Preparation and storage of enzymes
Once diluted with medium or buffer, proteolytic enzymes may undergo autolysis. Dissolve them
immediately before use. Special care must be taken with the deoxyribonuclease which is very
prone to shear denaturation. They must be mixed gently.
Reconstituted enzymes should not be stored at 2-8°C. If necessary, they can be aliquoted and
frozen at -20°C. Avoid repeated freeze-thaw cycles. All enzymes, upon reconstitution, can be
sterile filtered through a 0,22 μm membrane.
Generally, all enzymes, except trypsin, can be directly dissolved in a balanced salt solution or
buffer of choice. A stock solution of trypsin should be made initially by reconstituting the
enzyme in 0,001M HCl. This solution can be diluted into the digestion balanced salt solution
(BSS) or buffer immediately prior to use.
Optimization of cell dissociation
Although optimization of a cell isolation procedure for a particular cell type is dependent upon
the adequate recovery of cells having various required characteristics, some guidelines can be
established.
There is a complex relationship between cell yield and viability. In general, there is an area of
optimized recovery balanced between yield and viability; working near the middle of this range
will reduce variability in the results of the cell isolation procedure.
Trouble shooting
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•
Low yield/low viability
Over/under-dissociation, cellular damage. Change to less digestive enzyme and/or decrease
working concentration (e.g. from trypsin to collagenase/from Type 2 collagenase to Type 1)
•
Low yield/high viability
Under-dissociation and/or incubation time monitor both yield and viability response. If yield
remains poor, evaluate a more digestive enzyme and/or the addition of a secondary enzyme(s)
•
High yield/low viability
Good dissociation, cellular damage. Enzyme overly digestive and/or at too high a working
concentration. Reduce concentration and/or incubation time and monitor yield and viability
response. Try diluting the proteolytic action by adding bovine serum albumin (BSA) (0,1-0,5%
w/v) or soybean trypsin inhibitor (0,01-0,1% w/v) to the tissue suspension being dissociated.
Try using less proteolytic enzyme although yield may be affected and should be monitored
•
High yield/high viability
The place to be. Consider evaluating the effect of dissociation parameters to learn their
limitations for future reference
Sub-culturing of cell lines
Routine sub-culturing - adherent
When an attached cell line has either covered the surface available for growth, or depletes the
nutrients in the surrounding medium, it must be sub-cultured into new vessels, e.g. flasks or
dishes.
Attached cells will exhibit strong to very light adherence, depending on cell type. For example,
a confluent monolayer of the dog kidney cell line, MDCK, adheres to surfaces very strongly and
requires both a protease, e.g. trypsin and a chelating agent e.g. EDTA to detach the cells.
Conversely, the Chinese hamster cell line (CHO) can be loosened by gentle tapping of the
culture vessel, or pipetting off the monolayer. However, the majority of attached cell lines will
require at least the addition of a protease to dissociate the cells from the surface on which they
are growing.
To prevent either microbial infection or cross-contamination of cell lines, it is essential to
maintain separate reagent bottles for each cell line, e.g. such as PBS, trypsin, medium. If only
small quantities are required, aliquot the reagent into individual bottles. Mark each bottle in use
with the cell line name and date of use.
Additionally, each cell line must be handled separately, always working with those which have
been characterized first. Allow a 10 - 15 min ‘clearance time’ between working with different cell
lines, making sure that the flow cabinet has been thoroughly decontaminated with 70 % alcohol
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by wiping all internal surfaces, including the sides and glass screen, and that all reagents for
the previous cell line are removed. If gloves are worn, these must also be changed. Hands
must be washed after each subculture.

Prewarm all the cell culture medium and PBS to the culture temperature. Allow the trypsin
or trypsin-EDTA solution to reach room temperature. Remember that repeated warming
will inactivate trypsin

Microscopically examine the culture for its level of confluence of the cells, cellular
morphological appearance and any signs of microbial contamination

Decant the culture medium from the cells by either pouring or pipetting it off.
If the
medium is poured off (in case of flasks) invert the culture so that the medium runs down
the side of the flask opposite to that to which the cells are adhered. Remember that
decanting rather than pipetting off medium increases the risk of contamination.
It is
essential, whenever possible, to exclude antibiotics from the medium, in order to detect
any infections at an early stage

Wash the cells with a volume of Ca2+/Mg2+ - free PBS equal to at least half the volume of
growth medium. Decant the PBS. Repeat, if the cells are of the type known to be strongly
adherent

Add trypsin or trypsin-EDTA solution to the cells, 1 - 2 ml per 25 cm 2 of flask surface, and
gently spread over the entire surface by tilting the vessel. Leave inside the cabinet for 15 60 sec, at room temperature, and then decant or pipette off most of the solution

Place the cultures in an incubator at their normal growing temperature, e.g. 24 °C for
insect cell lines and 37 °C for mammalian cell lines. The cells should detach after 2 - 10
min as seen by gently tilting the vessel and observing if the cell layer moves. Gently tap
the side of the vessel to increase detachment. Detached cells impart a milkiness to the
suspension

Microscopically examine the culture flask to determine if all the cells have detached. Reincubate if necessary

Collect the cells by pipetting culture medium over the surface of the vessel, i.e. 2 - 5 ml per
25 cm2 of flask surface. Mix carefully to disperse the cells into a single-cell suspension.
Measure the total suspension volume. Remember that trypsin activity is neutralized by
serum proteins. It is important to ensure that the volume of serum in the culture medium of
the new culture vessels is equal to, or exceeds, the volume of trypsin in the vessel
subcultured. Cells will not attach if the trypsin is not neutralized

At this point the cells can be counted . When culturing a new cell line it is advisable to
make a viable cell count to estimate the maximum cell density achieved, i.e. cell/cm 2, and
the expected viability
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
Transfer the cell suspension into prepared culture vessels by inoculating the appropriate
volume of the cell suspension. For routine subculture a split ratio is used (1:2 or 1:3 etc.).
This is calculated as the ratio of surface areas; for example, a 75-cm2 has a split ratio of 1
into 3 (1:3), i.e. three 75 cm2 Roux flasks or only one 225 cm 2 flasks

Place the inoculated vessels in an incubator at the correct temperature. If a CO 2 incubator
is used leave culture flask caps loose
Check the cultures after a few hours for cell attachment and pH. If the cultures are too acid
(yellow), or alkaline (purple), the CO2 concentration must be immediately checked and
adjusted. To prevent the loss of cultures due to a failure of the supply of CO 2 keep the flask
saps tightened after cultures have equilibrated, i.e. 1 – 2 h.
The duration between sub-cultures will depend on the incubation temperature and cell type.
The majority of mammalian cell lines require sub-culturing every 3 - 7 days. If the duration is
longer than 5 days, change the medium of the cultures every 3 - 4 days.
Quantification of cells
To measure performance, to achieve reproducibility, or to make comparative studies, a means of
quantifying the cell population is needed. Classically, direct count of cell numbers using a microscopic
counting chamber (haemocytometer), usually in conjunction with a vital dye (e.g. trypan blue) to
distinguish viable and non-viable cells, is used. However, all vital stains are subjective and cannot
give absolute values, and by counting cell numbers no differences in cell size/mass are taking into
account. The method is simple, quick and cheap, and requires only a small fraction of the total cells
from a cell suspension.
Automation of cell counting is possible with electronic counters, especially for non-clumping single
suspension cells. Other methods for total cell numbers include:
•
counting cell nuclei after dissolving the cytoplasm. This is particularly useful for large clumps of
cells where cells are inaccessible (e.g. in matrices), or where cells are difficult to trypsinize off
substrates (e.g. micro-carriers)
•
if cell mass, rather than number is the important factor then a cell constituent has to be measured
•
an indirect measurement can be used e.g. glucose or oxygen consumption rates. Other
metabolites include lactic and pyruvic acid and carbon dioxide. There are reservations about
these methods as these metabolic rates are not constant throughout the growth cycle of a
specific cell and may also be influenced by changes in the culture, which is the subject of the
investigation. These biochemical methods are best used over a time-course in culture so that
successive readings will show a definite trend in the culture dynamics, e.g. stationary, growing or
dying. A combination of one or two methods is the ideal.
Haemocytometer cell counts
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In order to ensure that cell cultures have reached the optimum level of growth before routine
subculture or freezing, it is helpful to obtain an accurate cell count and a measure of the
percentage viability of the cell population.
The most common routine method for cell counting which is efficient and accurate is with the
use of a haemocytometer.
A thick, flat counting chamber coverslip rests on the counting chamber at a distance of 0,1 mm
above the base of the slide. The base of the slide has rulings accurately engraved on it,
comprising 1 mm squares, some of which are further divided into smaller squares.
When a cell suspension is allowed to fill the chamber, the cells can be observed under a
microscope and are counted in a chosen number of ruled squares. From these counts, the cell
count per ml of suspension can be calculated. Hybridoma cells and others which grow in
suspension may be counted directly. Cell lines which are attached will need to be removed
from the tissue culture flask by trypsinization. Since accuracy of counting requires a minimum
of approximately 105 cells/ml it may be necessary to re-suspend the cells in a smaller volume of
medium.
To ensure that a cell culture is growing exponentially, it is useful to know the percentage of
dead cells and hence the stage of growth of the cells. This can be estimated by their
appearance under the microscope, as live healthy cells are usually round, refractile and
relatively small in comparison to dead cells, which may appear larger, crenated and nonrefractile when in suspension. The use of viability stains such as trypan blue ensures a more
quantitative analysis of the condition of the culture. Trypan blue is a stain which will only enter
across the membranes of dead/non-viable cells.
When a cell suspension is diluted with a solution containing trypan blue, viable cells stay small,
round and refractile. Non-viable cells become swollen, larger and dark blue. Both the total
count of cells/ml and percentage of viable cells can be determined.
The use of different cell cultures
Monolayer cell culture techniques provide versatile and useful methods for isolating and propagating
many viruses and for performing serological tests. The effects of viruses on the cells may manifest as
cell degeneration or cytopathic effect (CPE), development of haemagglutinins or other antigens,
haemadsorption phenomena, or interference with the effects of an ‘indicator virus’ which is added
later.
In general four different types of cell cultures can be used. These are primary cultures (directly
derived from the disaggregated cells of monkey kidney, human amnion or other tissues), or
secondary cultures (derived from the former by passage) in which the cells are still morphologically
similar to those in the primary cultures, established cell lines (e.g. Hela) or also refer to as continuous
cell lines, which are capable of indefinite serial growth in culture and semi-continuous lines.
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Established or continuous cell lines have special uses and free the laboratory from dependence on a
source of primary tissue. Semi-continuous lines of human embryo cells, unlike established lines,
retain their characteristic diploid chromosomal configuration; they are sensitive to a wide range of
viruses but unfortunately they cannot be sub-cultured indefinitely and tend to lose their viability
between the 30th and 50th passage. Preservation of early passages of these cells at temperatures of –
70°C or lower provides an almost unlimited supply from one embryo. Low temperature storage is also
useful for keeping stocks of continuous cell lines, which may be lost by microbial contamination, or
may lose virus sensitivity with continuous cultivation, and for preservation of freshly trypsinized cells
for primary cultures.
A number of important conditions must be satisfied for successful cell culture:
•
The temperature should be 36-37°C (except for certain cells, e.g. epithelial cells, which grow
better at a slightly lower temperature
•
The pH between 7,2 and 7,4. Fortunately, most cells survive and grow in a wider pH range (6,87,8) and at a temperature of 33°C or lower
•
The osmotic pressure of the medium is critical: it is mainly determined by dissolved crystalloids
but may be severely affected by increases in glucose concentration. Inorganic ions essential for
immediate survival of cells are sodium, potassium, calcium, magnesium, iron, carbonate,
phosphate and possibly sulphate. Cell cultures require amino acids and vitamins (which the
addition of serum provides) or they are given as a synthetic mixture. As a source of
carbohydrate, glucose is usually added
•
Both oxygen and carbon dioxide are required, and are provided either as a mixture of CO 2 and
air supplied to the culture vessel or by sealing the vessel tightly to retain CO 2 produced by cell
metabolism
•
Aseptic techniques and the most scrupulous preparation of glassware, media and reagents are
essential
Because of the risks of contamination and infection, cell culture in the diagnostic laboratory is
conducted in closed vessels consisting mainly of stoppered tubes or flat-sided bottles. Cultures are
set up in an initial growth medium of which the simplest is Hanks Balanced Salt Solution (BSS)
supplemented with lactalbumin hydrolysate and serum. Fastidious cells may require more complex
chemically-defined media containing amino acids and vitamins such as (Eagle’s MEM or Parker’s
Medium 199) which are also supplemented with serum for growth. Once the cells have formed a
confluent monolayer, cultures are changed to a maintenance medium designed to maintain cultures
for as long as possible in a healthy state without stimulating cell growth. This is achieved by reducing
the serum concentration, or omitting it altogether. For growth of viruses in cell cultures, it may be
necessary to use a particular inoculation medium whose composition depends more on the growth
properties of the virus than of the cells.
The use of organ cultures (in contrast to tissue cell cultures) in a diagnostic virus laboratory is at
present of limited value. The object of this technique is to maintain the architecture and normal
function of the tissue as occurs in vivo and is usually achieved by culturing small fragments of organs
in chemically defined media. Organ cultures of ciliated nasal or tracheal epithelium from human
embryos have led to the recognition of new human respiratory viruses (e.g. coronaviruses) which are
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not detectable by conventional methods. More recently these techniques have been adapted to the
maintenance of human embryonic intestine for the study of diarrhoea illnesses and of human
embryonic conjunctivae for the isolation of some strains of enteroviruses. In Veterinary Clinical
Virology, the tracheal rings of chickens are generally used for the identification of coronaviruses.
Table 2: The use of different cell lines for viral isolation
Foot- and- mouth disease virus
Primary porcine kidney, BHK, Vero
Rinderpest virus
Bovine kidney, Vero
Infectious bovine rhinotracheitis
Vero, MDBK
Bovine virus diarrhoea virus
Primary bovine foetal kidney
Bovine ephemeral fever virus
BHK
Bluetongue virus, African horsesickness virus, and other
BHK, Vero, insect cell lines e.g. C6/36
orbiviruses
Equine rhinovirus 1
Vero
Equine rhinovirus 2
RK-13
Canine distemper virus
Vero
Canine hepatitis virus
Vero, MDCK
Newcastle disease virus
Vero
Rabies virus
BHK, Vero
Lumpy skin disease virus
Primary bovine dermis, Primary equine lung, MDBK
Rift Valley fever virus
BHK, Vero, insect cells e.g. C6/36
Wesselsbron disease virus
Vero, insect cells e.g. C6/36
Canine parvovirus
Feline kidney
African swine fever virus
Leukocytes, porcine kidney
Equine herpesvirus
Primary equine lung
Feline enteric coronavirus
CRFK, NLFK
Feline herpesvirus
CRFK, NLFK
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Feline calicivirus
CRFK, NLFK
Equine arthritis virus
RK-13, Vero
C6/36
Aedes albopictus clone C6/36
CRFK
Crandell feline kidney
NLFK
NL- feline kidney
VERO
African green monkey cells
BHK
Baby hamster kidney cells
RK-13
Rabbit kidney
BHK
Baby hamster kidney cells
MDBK
Madin-Darby bovine kidney
MDCK
Madin-Darby canine kidney
Contamination of cell cultures
Bacterial and fungal contamination of cell cultures occurs when they are inoculated with clinical
material naturally contaminated with micro-organisms. The incorporation of high concentrations of
antibiotics into transport medium and other reagents used for clinical specimens, and their routine use
in all cell cultures, reduce it to a very low level. Cultures contaminated with bacteria and yeasts
rapidly become turbid and the pH generally falls. Under the low power of the microscope, yeasts are
distinguished from bacteria by their larger size, budding forms also commonly being present. Mould
contamination is characterized by a rapidly proliferating macroscopic mass whose form depends upon
the particular contaminant.
Mycoplasma is a generic term given to organisms of the order Mycoplasmatales which can infect cell
cultures. Those which belong to the families Mycoplasmataceae
Acholeplasmataceae (Acholeplasma) are of particular interest.
(Mycoplasma)
and
The first observation of mycoplasma infection of cell cultures was by Robinson et al., (1956). The
incidence of such infection has since been found to vary from laboratory to laboratory. Mycoplasmas
do not necessarily manifest themselves in the manner of most bacterial or fungal contaminants, e.g.
pH change or culture turbidity. It is important therefore to adopt an active routine detection procedure.
Mycoplasmas have been shown to elicit various effects, including:
•
Induction of chromosome aberrations (Aula & Nichols 1967)
•
•
Induction of morphological alterations, including cytopathology (Butler & Leach 1964)
Interference in the rate of growth of cells (McGarrity et al., 1980)
•
Influence nucleic acid (Levine et al., 1968) and amino acid (Stanbridge et al., 1971) metabolism
•
Induce membrane alteration (Wise et al., 1978) and even cell transformation (MacPherson &
Russel 1966)
A range of assay techniques are available for the detection of mycoplasma contamination. These
include staining, culture, DNA probes and co-cultivation. To remove the risk of false negatives two
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methods at least should be employed. The enrichment broth and agar culture and Hoechst 33258
DNA staining are recommended techniques.
The fluorochrome dye Hoechst 33258 binds specifically to DNA. Cultures infected with mycoplasmas
are seen under fluorescence microscopy as fluorescing nuclei with extranuclear mycoplasmal DNA,
whereas uninfected cell cultures contain fluorescing nuclei against a negative background.
Mycoplasmas may appear as filamentous forms, some of which may branch, indicating a culture in
logarithmic growth, or as cocci, which is typical of an aged mycoplasma culture. Contaminating fungi
or bacteria will also stain using this technique but will appear much brighter and larger than
mycoplasmas.
The main advantage of this dye technique is that it is speedy and that non-cultivable mycoplasma
strains, which have been detected in cell cultures, can be observed.
Cryopreservation
Polge et al. (1949) made the observation that glycerol enhanced the survival of fowl semen at –79°C.
The technique was then used for other cell types. Lovelock and Bishop (1959) discovered the
cryoprotective properties of DMSO. Mazur (1970) showed that cells cooled slowly in the presence of a
cryoprotectant shrink, and do not contain intracellular ice. DMSO increases the membrane
permeability and the reversal of intracellular ice formation. The prevention of intracellular damage on
thawing is essential to successful cryopreservation (Mazur 1977).
A major achievement permitting the development of animal cell technology has been the
determination of the parameters for routine cell cryopreservation. The importance of a stable, reliable,
secure supply of material held at temperatures below –130°C cannot be overstated. At its most
simple, the technology is based on slow freezing and fast thawing, together with high protein
concentration and the presence of an agent which increases membrane permeability. The other,
perhaps obvious, ingredient is ‘healthy’ cells.
The first consideration is whether glass or plastic ampoules should be used; both have their
advantages and disadvantages. A correctly sealed glass ampoule will not permit entry of liquid
nitrogen which might be the case with a screw-capped plastic ampoule. This is important in two
respects: the presence of liquid nitrogen during rapid warming could risk the danger of explosion due
to the rapid expansion of gas; liquefied gas is not sterile and could carry contaminating objects into
the ampoule. However, glass ampoules are comparatively difficult to seal, and have to be thoroughly
tested before freezing. An additional problem with glass is labelling; ceramic ink is needed which
requires specialized equipment and a certain degree of expertise. For most users, plastic ampoules
are more convenient as they are pre-sterilized and will label with marker pens. In addition, there is a
commercially available heat-shrinking sheath which can be placed over the ampoules to minimize the
risk of entry of liquid.
The means of freezing at a controlled rate is another major consideration. A controlled-rate freezing
apparatus is considered by some as an expensive luxury. Nevertheless, successful freezing is
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virtually guaranteed when alternative methods are used. Some pre-validation of the methodology to
be employed is strongly recommended.
Long term storage
Long-term storage of cell stocks requires good record keeping and maintenance of an inventory
with each removal logged to ensure adequate stock levels.
A wide range of liquid nitrogen refrigerators is available. Whatever the choice, they must be
alarmed to prevent accidental drying out. In addition, the ampoules must be kept in more than
one location.
In the routine handling of cell stocks, it is essential to avoid wide fluctuations in storage
temperature. Frequent removal of the inventory system from the refrigerator with the associated
variation in storage temperature will reduce viability. It is recommended that stocks are
periodically checked for maintenance of viability - every 5 years.
Mechanical freezing
Cryopreservation using –80°C or –130°C in a mechanical freezer is probably the least reliable
of methods. However, low-temperature freezers are widely used to cryopreserve cell lines.
To achieve slow cooling, ampoules must be heavily insulated. A block of polystyrene containing
individual holes sufficiently large to take ampoules should be prepared. The polystyrene must
be 1 - 2 cm deep all around the ampoule, with no air spaces. The insulation required is placed
in the block, which is then placed near the middle of the freezer and left overnight (16 –24 h)
before being transferred to a liquid nitrogen storage vessel.
Before using this method on a regular basis, a series of tests should be made to monitor the
cell viability after freezing. If it varies significantly from the viability of the cells prior to freezing,
i.e. a drop of more than 15 – 20%, the insulation will have to be modified. This method is not
recommended for preparing master or reference stocks for long term storage.
Two-stage freezing
Ampoules are kept at a holding temperature of –20° to –40°C for up to 24 – 48 h and then
transferred directly to –196°C.
Similar results can be achieved for a small number of ampoules by using a small device which
holds them in liquid nitrogen vapour in the neck of a Dewar flask. After a holding time of 10-20
min the ampoules are plunged into liquid nitrogen and then transferred to their final storage
location.
Preliminary experimental work is required to determine the optimum conditions. In all cases, the
cells should be examined for viability following cryopreservation.
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For certain specialized cell types, it may be considered necessary to include particular essential
growth factors to the freeze medium in order to maintain surface receptor stability during
cryopreservation. For example, hybridomas often cause problems on revival.
It is essential to use freshly prepared freezing mixture every time. DMSO should be obtained
from a supplier who is able to offer the most recently prepared stocks available. Once received,
it should be filter sterilized (0, 2 μm) through a filter specifically designed for DMSO and stored
in a glass container with an air-tight stopper or lid at –20°C. This will ensure that no problems
develop due to oxidation of the DMSO. In those cases where DMSO causes differentiation,
glycerol should be used instead.
Thawing
•
Achieve rapid thawing by transferring ampoules directly to a water bath containing water at
37°C. If an ampoule contains potentially hazardous material, it is advisable to add 1 - 2%
(w/v) chloramine-T to the water in the water bath.
•
Take care not to submerge the cap of the plastic ampoule in order to prevent
contaminated water from entering the ampoule. A simple method is to use a plastic rack
designed to hold tubes of the same diameter as that of the ampoules, and to place it in the
correct depth of water. Alternatively, a piece of foam polystyrene foam containing holes
into which the ampoules are fitted; this will float on the surface of the water.
•
Once thawed, transfer the ampoules to the sterile work area. Submerge glass ampoules in
70% ethanol for a brief period and thereafter allow them to dry. A pre-scored ampoule can
be snapped open using a paper tissue soaked in 70% ethanol. Unscored types must first
be scored with a diamond before soaking in 70% ethanol, after which they are opened.
Plastic ampoules are opened whilst gripped with a tissue soaked in 70% ethanol.
•
Using a pipette, transfer the ampoule contents into either a flask or a centrifuge tube
containing prewarmed growth medium. A slow transfer is recommended. If it essential to
remove the cryoprotectant at this stage, centrifugation should be at the lowest speed
necessary to form a cell pellet. For cultures started directly from an ampoule, the first
medium change should be at 24 h after transfer unless recommended otherwise.
Start-up cell density is of critical importance. The cells must be encouraged to become
conditioned to the medium as rapidly as possible and prevent a protracted lag phase. For
adherent cells, an initial density of 2-5 x 104 cells /ml is normally appropriate.
•
Examine the cells daily using an inverted microscope and subculture them as soon as
confluency or maximum cell density is reached. This is frequently indicated by a medium
colour change to orange or orange-red, i.e. acid pH.
Safety considerations
During the handling of frozen ampoules, a full-face mask and gloves must be worn. Ampoules
containing pathogens should be thawed on an outer sealed container. In order to prevent
explosion of the ampoules on their removal from the liquid phase, it is recommended that
transfer to the gaseous phase is carried out 24 h prior to thawing. To reduce the risk of injury
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from exploding ampoules during thawing, it is advisable to place them in a perforated container
before they are placed in a water bath.
Demonstration / confirmation of the presence of viruses in infected cell
cultures
Introduction
When evidence of viral infection is observed in inoculated cell cultures, either by the
development of CPE or by a positive haemadsorption test, the affected cultures should be subcultured in order to demonstrate propagation of the effects. If the CPE observed is minimal, one
may increase the likelihood of a successful passage if the CPE is allowed to progress. Since
some viruses are cell associated one should ensure that cells are transferred as well as fluid
when making a passage.
Once evidence of the presence of a virus is obtained, an identification procedure can be
selected based on the assessment of the most likely viruses to display the effects noted and
which appear consistent with the nature of the specimen and the patient’s illness, i.e. one
would not look for influenza virus as an isolate from a stool specimen. Although numerous
methods of identification have been developed, usually only a few of them are routinely used by
any particular laboratory. With the exception of the biophysical characterization, which is used
to obtain the taxonomic assignment of a virus that proves difficult to identify, the methods used
are mostly immunological. The most commonly used techniques are immunofluorescence and
neutralization tests, and ‘reversed’ serology tests, such as the haemagglutination inhibition and
complement fixation (the latter in general used anymore).
The immunofluorescence test (direct or indirect) is often the most convenient way to determine
the identity of many common pathogenic viruses. These techniques are most useful whenever
only one or one of a small number of agents is considered most likely to be involved. In the
absence of reagents or facilities for performing immunofluorescent identification, perhaps the
next most convenient means of viral identification is the use of ‘reverse’ serology. This is where
the virus-containing culture is used as an unidentified serologic antigen, which is to be tested
for reactivity with a number of specific reference sera. Haemagglutination inhibition is well
suited for this type of identification in that the reactivity of the antigen is known before the test is
carried out; complement-fixation is more difficult to use, both technically and in that the
reactivity of the unknown antigen is not assured before the test is run. Serotyping of viruses can
also be accomplished by means of the neutralization test.
Handling of specimens for the isolation and identification of viruses
Preparation of clinical specimens for viral isolation
Most clinical material requires processing before being tested in order to remove extraneous,
non-virus matter. In general, it is necessary to prepare a reasonably concentrated,
homogeneous suspension of the specimen free from particulate matter which might block
hypodermic needles and pipettes or obscure specific effects in tissue cultures and other test
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systems. Certain types of specimen e.g. swabs, nasal washings, lesion scrapings, and
cerebrospinal fluid usually contain little if any solid debris, in which case they may be tested
without prior treatment.
Nasal washings
Centrifuge the nasal washing fluid at 2000 rpm for 15 min at 4°C to remove debris. This
procedure may be omitted if no debris is present. Carefully pipette off the supernatant fluid.
Freeze and store the unused supernatant fluid and remainder of the specimen. Depending on
the time period of storage material can be stored either at 4°C or at –20°C.
Cerebrospinal fluid
No processing is generally required. If cellular material is obviously present, transfer the fluid to
a centrifuge tube and proceed as for nasal washings. Antibiotics are usually added. The type
and concentration of the antibiotics used depends on the laboratory, but in general a broad
spectrum antibiotic is used.
Urine
No processing is generally required. If cellular material is obviously present, transfer the fluid to
a centrifuge tube and proceed as for nasal washings. Antibiotics are usually added.
Saliva
No processing is generally required. Antibiotics are usually added.
Biopsy or necropsy tissue and organ specimens for viral isolation
Weigh approximately 1 g of the specimen and place it in a Ten-Broeck tissue grinder of suitable
size or a pestle and mortar with sterile sand. Add sufficient phosphate-buffered saline
containing extra antibiotics to make a 10% w/v suspension, and grind by hand until a uniform
homogenate is obtained. The time required for complete homogenization depends on the
nature of the tissue. Relatively non-fibrous tissues, such as brain may be adequately
suspended within 1 – 2 min, whereas a much longer time is required for tissue, such as muscle.
In the latter case, the grinder should be embedded in crushed ice to avoid overheating during
the grinding process. Suspensions of tissues of even greater resistance, such as keratinized
skin, are best made by grinding the tissue in a mortar containing washed sand and phosphate–
buffered saline. When homogenized, transfer the suspension to a centrifuge tube and
processed as for nasal washings.
Faeces
Faeces samples are not normally used for viral isolation in cell culture. Electron microscopy is a
rapid and sensitive method to identify many viruses. If cell cultures are used:
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•
One method is to weigh about 1 g of faeces into a screw-capped bottle containing about 5
g coarse beads approximately 5 mm in diameter. Add sufficient phosphate-buffered saline
containing a high concentration of antibiotics to make a 10% w/v suspension. Shake
vigorously by hand for 30 sec or until the faeces are uniformly dispersed. Centrifuge the
resulting suspension at 4000g for 20 min at 4°C. High speed centrifugation is desirable
since it removes the majority of bacteria and certain toxic substances from the stool
suspension. Carefully remove and save only the upper two-thirds only of the supernatant
fluid, discarding the remainder. Freeze and store at approximately –20°C.
•
Faecal samples can also be extracted with chloroform before being used for the isolation
of viruses in cell cultures. This method entails: A suspension of the faeces is made using
phosphate buffered saline (PBS). Chloroform is then added, and the suspension shaken
and centrifuged. The supernatant is used to infect cell cultures.
Swabs
Procedures used for the preparation of nasal, throat, rectal and vesicle swabs are identical.
Macerate the swab well in the transport medium in which it was submitted, to disperse any
virus particles. Transfer the resulting suspension to a centrifuge tube, and proceed as for nasal
washings.
Tissue scrapings of lesions
Add coarse glass beads to the transport medium in which the scrapings were transmitted and
submitted. Shake vigorously by hand for about 5 min at room temperature in order to disrupt
any intact cells. Transfer the resulting suspension to a centrifuge tube and proceed as for nasal
washings.
Blood
Blood in heparin is generally used for viral isolation. The blood is added to the cells, left for an
hour and then the cells are washed with medium before adding fresh medium to the cells. In
some cases the blood can be toxic to the cells, but in general the cells will flourish. Blood
destined for serological investigation is best collected in containers, which can be centrifuged
subsequently.
Inoculation of cell culture
Care must be taken in the preparation of the material suspected to contain the antigen or
antigens. The isolation cabinet should be cleaned before using a disinfectant. Do not prepare
tissue specimens from more than one animal or more than one antigen at the same time in the
cabinet.
Viruses are generally isolated from diagnostic samples using cell monolayers in tubes or 25
cm2 flask cultures. Before use the medium in the flask is changed from a growth medium to an
equivalent volume of the maintenance medium of choice.
The serum content of the
maintenance medium is lower than that of the growth medium. This is to prevent the
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overgrowth of the cultured cells and their consequent rapid degeneration. If normal bovine
serum has been used in the growth medium all traces of it must be removed to prevent the
presence of antibodies which may interfere with the isolation of certain viruses. If a serum-free
maintenance medium is used, the monolayers are first washed free from residual growth
medium. This is done by aspirating the fluid and rinsing twice with maintenance medium before
finally adding the same volume of maintenance medium as there originally was of growth
medium to the flask.
The cultures are screened before inoculation. Medium changing carried out the day before
inoculation will ensure that the pH is not too high and will assist in the screening process.
Screening should be carried out microscopically. A volume of 0,2 ml of the prepared material
from the clinical specimens are inoculated into each of preferably four tubes/flasks which are
incubated immediately.
Four tubes/flasks in each batch are kept as uninfected negative
controls. In order to reduce the risk of cross-contamination between specimens, only one
specimen is inoculated at a time, each specimen container and the inoculated tubes being
properly stoppered and labelled before proceeding to the next. Moreover, since inoculation
inevitably involves contamination of the tube/flask lip during entry and exit of the inoculating
pipette, the tube/flask must be adequately flamed before closing. This requires care, since too
rapid heating may result in spluttering of infective fluid. However, it is important that no residual
virus remains on the lip.
Maintenance of inoculated cultures
Once specimens have been inoculated into cell cultures, all further manipulations which may
provide opportunities for cross-contamination of the cultures to occur should be avoided.
Primarily, this comprises the opening of the tubes for the purpose of changing the medium, or
adding buffers to adjust the pH. In addition blind passage should be avoided as far as possible,
to reduce the total number of cell cultures required and thus the general workload involved.
One approach to these problems is to increase the buffering capacity of the maintenance
medium in which the cultures are kept by the addition of synthetic buffers to the medium: the
HEPES buffer is widely used for this purpose, but the MOPS buffer is less expensive and works
well. A well-buffered maintenance medium containing galactose in the place of glucose reduces
the formation of acids by cellular fermentation is an alternative, but very expensive, medium.
Inoculated cell cultures are held, with daily to three times a week observations, for two weeks,
or as long as a given lot of cell cultures do not undergo spontaneous degeneration – observed
in the non-inoculated control cultures. Specimen toxicity, in the form of nonspecific CPE,
usually develops within a few hours after inoculation of cell cultures. If non-specific CPE
appears in inoculated cell cultures, especially if it occurs shortly after the inoculation, the
affected cultures should be used to inoculate fresh cultures (passed), to dilute out potential
toxic material in the inoculum.
Cytopathic effect
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An inoculated culture should be examined microscopically every day so that the onset of
changes may be observed and their progress followed. In this way, any virus-specific effect
(cytopathic effect or CPE) may be differentiated with certainly from any non-specific
phenomena. In practice, it is often not possible to undertake daily examinations, especially
when large numbers of tubes are involved. With some experience of the rates at which
changes induced by different viruses advance but also of the agents likely to be encountered in
given clinical materials, the times at which tubes are read may be profitably adjusted.
It is important, however, that a reading is always made on the first day after inoculation when
any non-specific cell changes due to toxicity of the clinical specimen will be disclosed. Toxic
changes are variable. Mild toxicity is frequently manifest as a diffusely spread granularity of the
cell cytoplasm. In toxicity of a more severe degree, the cells may become pyknotic, or bizarre
forms may be apparent. When stool specimens or rectal swabs are inoculated, traces of
residual trypsin from the gut may actually digest the cell monolayer from the glass. Non-specific
toxic changes of the foregoing kind are readily distinguished from almost all kinds of virusinduced changes which, in any case, are unlikely to appear during the first 24 hours following
inoculation. The importance of this first-day reading is that non-specific changes may progress
rapidly to total destruction of the culture, which at a later stage is unlikely to be distinguishable
from a culture destroyed by specific virus action. Non-specific changes observed on the day
after inoculation may often be reversed, or their progress halted, by removing the medium and
replacing it with an equal volume of fresh maintenance medium. Since fluids removed 24 h
after inoculation may contain large amounts of virus, the changing procedure should be done
with the utmost care in order to restrict cross-contamination.
It is convenient to adopt a numerical system of scoring when reading tubes for cytopathic
changes. The extent of involvement is usually scored on a scale from 0 to 4. At one end of the
scale, 0 represents a normal culture without CPE, while at the other extreme; a completely
degenerated culture is scored 4+. Varying degrees of involvement are given in between values.
Thus, a culture with one or a few foci of infection is scored 1+, while a tube in which
approximately 10% of the monolayer is affected is rated 2+. When 50 – 75% of the monolayer
is involved, a 3+ rating is given. It is obvious that such a system entails an element of
subjective interpretation. However, a single person can achieve a high degree of reproducibility,
so that comparisons within and between experiments are valid.
The appearance of CPE in certain cell cultures is characteristic for some viruses. Respiratory
syncytial and distemper virus induce non-haemadsorbing multinucleated giant cell formation in
many cell cultures. Adenovirus infection results in the aggregation of the cells into ‘clusters’.
Herpes simplex virus gives rise to ballooning of human embryo cells, and the shrinking and
rounding of other cell types. Enteroviruses and rhinoviruses cause rounding of cells and are
difficult to distinguish from each other by their CPE. Examination of cultures showing complete
CPE by electron microscopy may be helpful in ‘grouping’ the agent, thus possibly minimizing
subsequent specific identification procedures.
Cultures not showing CPE
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The majority of cultures inoculated with clinical material almost never develop CPE. This may
be because of:
•
No virus was present in the sample
•
Most tissue culture systems are far from the ideal for growing non-adapted viruses, and
therefore clinical material may not elicit a CPE although incubated for a time adequate for
the virus to develop on theoretical grounds – the same virus may produce CPE if given
more time. Cultures failing to give CPE can be sub-cultured into further tubes. Cells as well
as medium are transferred. Prior to sub-culturing the cells are detached from the tube wall
by vigorous pipetting of the medium, and 0,2 ml of the suspension is then inoculated into
another tube of the same kind and incubated under the same conditions as before. Tubes
failing to produce any CPE by the end of the second period are recorded as negative. This
is referred to as the blind passaging of cells.
•
The susceptibility of the type of cells used. Some cell cultures may not be the ideal cell
type for a specific virus. This can be very common in Veterinary Clinical Virology because
of the large number of different animal species that are dealt with. The clinical signs of the
animal as well as the expected virus in the sample must be considered.
•
Cell cultures, derived from insect cells (mosquitoes, etc.) will, because of their vector
capabilities for certain viruses, not produce CPE. Such cell cultures then become
persistently infected with the virus and, in a clinical diagnostic situation, one needs to ‘blind
passage’ these cultures to a mammalian cell culture in which the virus (if present) in most
cases will produce CPE.
Inclusion bodies
Viral replication proteins and assembled virions often accumulate in specific regions of the
nucleus (e.g. inclusion bodies or replication compartments in cells infected with herpesviruses
or adenoviruses) or cytoplasm (e.g. the Negri body in rabies-infected cells). The assembly of
these new structures in the infected cell often displaces host-cell components from specific
regions of the cell and leads to one form of CPE. The inclusion bodies, or areas of altered
staining in infected cells, are useful in diagnostic virology because they are found at locations in
the cytoplasm or nucleus (or both) which are characteristic of specific groups of viruses. Also,
they show characteristic staining properties in that they are basophilic or acidophilic. To clearly
see these inclusion bodies, cells are grown on coverslips, fixed and stained. Hematoxylin and
eosin is the stain of choice. These structures are used in diagnostic virology and are illustrated
by the following examples:
Parvovirus forms typical intranuclear inclusions, which can be used as a diagnostic tool.
Measles virus
The most striking and consistent feature of measles virus infection in vivo as in vitro is the
formation of multinucleated giant cells, which results from the fusion of infected cells with their
infected and uninfected neighbour cells. In tissue culture, these giant cells contain eosinophilic
cytoplasmic inclusion bodies.
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Reoviruses
Cells infected with reovirus develop characteristic cytoplasmic inclusions referred to as ‘viral
factories’ or ‘inclusion bodies’. These inclusions initially appear as phase-dense granular
material scattered in the cell cytoplasm, which eventually coalesce and move toward the
nucleus. Dense perinuclear inclusions subsequently appear and increase in size as infection
progresses. Inclusion body formation is delayed when infected cells are maintained at low
temperature. Different cell lines vary in temporal sequence of inclusion body formation following
infection. Viral inclusion bodies have been shown to contain dsRNA, virus-specific polypeptides
and both complete and incomplete viral particles.
Orbiviruses
Virus synthesis and maturation occurs in the cytoplasm of infected cells, where inclusions and
crystalline arrays of virions are sometimes found. These virus inclusion bodies apparently
represent centres of viral replication or assembly. Accumulations of tubular structures or virusspecific tubules are also seen in infected cells.
Fixation and staining of coverslips
Cells grown on coverslips are used to identify viruses especially those which induce the
formation of inclusion bodies and syncytia. Coverslips are first fixed in a fixative, such as
ethanol or Bouins fixative, and then stained with the desired stain. They are then mounted onto
microscopic slides for observation.
Syncytia
The formation of multinucleated giant cells results from the fusion of infected cells with their
infected and uninfected neighbour cells.
Measles virus, distemper virus and other viruses of the Paramyxoviridae family have this
unique characteristic, which can be used for their viral identification.
Plaque test
Cell monolayers are infected with certain viruses which are allowed time to enable them to
settle on and attach to the cells. The latter are then overlaid with a nutrient agar medium to limit
the spread of the viruses. As a result, localized, often multiple, areas of cellular destruction and
lysis (CPE) called plaques are formed. These may be detected if the monolayers are stained
with dyes, such as neutral red or trypan blue, that are used to distinguish living from dead cells.
This technique can also be used for the quantification of viruses in a solution or specimen, in
which case the titre of the virus sample is then expressed as plaque forming units per volume.
Plaque separation - if a mixture of viruses is suspected the same technique can be used to
obtain pure cultures of viruses. It is also known as the plaque purification method.
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Haemadsorption
The detection of haemagglutinins either by haemadsorption or haemagglutination is widely
used to detect infection by membrane-bound viruses, such as orthomyxoviruses and
paramyxoviruses, which obtain their outer envelope by budding through the cell membrane and
may cause little or no CPE. Prior to budding, viral coded proteins (haemagglutinins) are
incorporated into the cell membrane. Even with strains that produce CPE, haemagglutinins are
usually detectable at an earlier stage. Cell cultures which are sensitive to myxoviruses should
be tested for their presence by haemadsorption 5 – 7 days after inoculation. Un-inoculated
control cultures should be tested together along with the inoculated cultures to detect
haemadsorbing viral contaminants which are frequently found in primary monkey kidney
cultures.
The haemadsorption test is a particularly convenient screening test, as one need only add a
few drops of dilute erythrocyte suspension to the tubes containing the control cells and to those
inoculated with the specimens being tested. The cultures are then refrigerated briefly to allow
adsorption to occur and to inhibit the relapse of the adsorbed cells by viral neuraminidases. The
cultures are inverted and inspected to determine if erythrocytes are adhering to the infected
cells. The culture medium can be removed and monolayers washed in order to facilitate the
inspection, but this is not always necessary. If the haemadsorption test gives an ambivalent
result it is recommended that the more sensitive haemagglutination test be performed.
Haemagglutination
Haemagglutination (HA is a property of some viruses. For human viruses, the human ‘O’ group
red blood cells (RBC) are normally used for influenza and parainfluenza viruses, and certain
enteroviruses and adenoviruses. Chicken RBCs are used for Newcastle disease virus and
porcine RBC for parvovirus of dogs.
Many viruses have optimal temperature and pH requirements for HA, and these variables along
with the need to absorb sera for naturally occurring agglutinins to some species of RBC, make
HA tests particularly difficult to reproduce with certain specificity.
Binding is accomplished by an interaction of viral proteins (haemagglutinins) located in the
outer coat of the virion or virion envelope with receptor sites on the RBC. Agglutination is a
result of the bridging effect of one virion binding to two RBCs simultaneously and these in turn
being bound to other RBCs by additional virions. The result is a lattice-like aggregate of RBCs.
The ability of viruses to agglutinate RBCs is influenced by pH and temperature.
Agglutination of RBCs is useful for the concentration and purification of some viruses, and as a
rapid presumptive test for the presence of these viruses in fluids from infected cell cultures and
chicken embryos. It is especially useful for assaying viral activity of cell cultures infected with
haemagglutinating viruses that produce little or no discernible CPE. Clinical specimens such as
faeces can also be directly examined for haemagglutinating (HA) activity. For example, a
presumptive diagnosis of canine parvovirus can be made by demonstrating HA activity to
porcine RBCs in faecal samples.
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Figure 1: Schematic presentation of the haemagglutination test
Serology
Serum viral neutralization test
Serum viral neutralization test (SNT) is the most widely used method of detecting and
measuring antibodies to viruses of veterinary importance. This test is generally considered to
be the most reliable of all serologic tests, being less prone to variation and less subjective in its
interpretation.
The principle of the test is based on the fact that the demonstrable activity of the virus - whether
it is CPE in cell cultures, clinical signs, lesions or death in embryonated chicken eggs and
animals - can be inhibited by specific antibody to that virus.
SNTs are almost always performing using cell cultures. Stock viruses for use in the tests are
previously grown, aliquoted, and stored at ultra-low temperature. These viruses are titrated
several times to determine the amount of virus present. Performance of the test entails the
dilution of the test serum in microtitre plates, followed by the addition of an equal volume of
virus suspension diluted to contain approximately 100-300 infective doses. Following incubation
of 1 to 2 h at 37°C, indicator cell cultures at a certain concentration are added. Plates are
sealed, incubated at 37°C, and observed daily for development of viral CPE. The presence of
specific antibody in the test serum inhibits the production of CPE.
The SNT is also used to identify unknown viral isolates in essentially the same manner as
described above. The only difference is that different known antibodies must be used and
identification of the unknown virus is accomplished by the one which inhibits the development
of CPE.
Serotyping
To determine the serotype of a virus the neutralization tests is normally performed using
specific antibodies. This test is carried out using a standard challenge dose of virus, which is
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determined by preliminary titration of the isolate. Unfortunately, the neutralization tests are
time-consuming and expensive due to the need to use large numbers of cultures for the titration
and neutralization steps, and the need to await virus growth in controls and non-neutralizing
serum reactions. Also the subject is complicated by the existence of heterologous neutralization
reactions, poorly neutralizing strains (which may be due to aggregated virus) and prime strains,
which react in one-way crosses with standard prototype strains used to produce the reference
sera for the neutralizations.
Haemagglutination inhibition test
The haemagglutination inhibition test (HAI) test is used to measure specific antibody, which
often corresponds with neutralizing antibody, and to identify virus isolated using specific animal
or bird antisera. Test sera often contain non-specific inhibitors of agglutination and may result in
false positive antibody tests. These can be removed by the adsorbing of sera with RBC or by
the treatment of sera with kaolin and the ‘receptor destroying enzyme’ prior to testing.
The HAI test is similar in principle to the SN test except that the viral activity being inhibited is
haemagglutination. HI tests are quite sensitive and highly specific, and are particularly useful
for measuring antibody to those haemagglutinating viruses that either grow poorly in cell culture
or produce little or no discernible CPE. Examples of such viruses are Type A influenza viruses
of most animal species, Newcastle disease virus of birds, and porcine parvovirus. HI tests are
usually performed in microtitre plates. Dilutions of test sera (25 µl) are made, followed by the
addition of an equal volume of virus suspension diluted to contain approximately 4 to 8 HA
units. The appropriate RBC suspension is then added (50 µl) and the plates are gently mixed
and allowed to incubate for 1 to 2 h at 4°C (for most viruses). If the specific antibody is present
in the test serum, agglutination of RBC will be inhibited and the RBCs will settle out in a welldefined “button”. Agglutinated cells, in contrast, will settle out in a thin layer over the entire
bottom of the test well, or in a rough, irregularly fringed button.
Figure 2: Schematic presentation of the haemagglutination inhibition test
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Figure 3: Schematic representation of the haemagglutination inhibition test
Electron microscopy
A feature of many viral groups, which can be used with advantage for diagnostic purposes, is
that they are morphologically distinct. When viewed in electron microscope (EM) herpes, pox,
adeno, rota, influenza and wart viruses are readily distinguishable, even to the untrained eye.
Widespread use of the EM in diagnostic virology is limited principally by the large number of
particles (>106/ml) required before even one of them can be seen on the microscope screen.
The diagnostic use of the EM for direct examination of clinical specimens is thus restricted to
viruses which are concentrated naturally in the body, e.g. herpes simplex, small pox or
chickenpox virus in vesicles, wart or orf virus in skin lesions, rotavirus, astrovirus and others in
faeces. Throat swabs and respiratory secretions contain little virus and although viruses have
been identified directly by electron microscopy in such specimens, the EM can best be used
indirectly to identify virus after it has multiplied to a sufficient level in cell culture. Generally, any
cell culture exhibiting complete CPE will contain enough virus for it to be visible in the EM, and
examination of such cultures is worthwhile as time and reagents can be saved by the early
identification of the virus group to which it belongs. The information provided by the EM is
purely morphological, i.e. size, shape and surface detail. Specificity, therefore, is limited to the
virus group: although adenoviruses can be distinguished from herpes viruses, adenovirus type
2 is morphologically indistinguishable from adenovirus type 5, and herpes simplex type 1 from
herpes simplex type 2. Some differentiation within groups can be achieved by observing
whether the virus will react with type-specific antibody as shown by antibody molecules visibly
attached to a virus or by clumps or virus particles aggregated by antibody molecules. However,
this combination of serology and electron microscopy – immune electron microscopy - is mainly
used for identifying viruses of poor morphological differentiation and low titre. It can only be
recommended for identifying members within a virus group if high titred, type specific sera are
available.
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