1. INTRODUCTION

Animal cell culture (ACC) is the process of culture of animal cells outside the

tissue (in vitro) from which they were obtained. The process of ACC is carried

out under strict laboratory conditions of aseptic, sterility and controlled

environment involving temperature, gases and pressure. It should mimic the in

vivo environment (medium) successfully such that the cells are capable of

survival and proliferation in a controlled manner.

The artificial environment is generally known as media. A media comprises an

appropriate source of energy for the cells which they can easily utilize and

compounds which regulate the cell cycle. A typical media may or may not contain

serum. The latter is called a serum-free media. Some of the common sources of

serum can be fetal bovine serum, equine serum and calf serum.

Theoretically, cells of any type can be cultured upon procurement in a viable state

from any organ or tissue. However, not all types of cells are capable of

strangeness in such an artificial environment because of many reasons on which

the artificial environment may fail to mimic the biochemical parameters of the

source environment. Some good examples include the absence of growth

regulators, cell to cell signal molecules, etc

2. HISTORY OF ANIMAL CELL CULTURE

It was Jolly, who (1903) showed for the first time that the cells can survive and

divide in vitro. Ross Harrison, (1907) was able to show the development of nerve

fibres from frog embryo tissue, cultured in a blood clot. Later, Alexis Carriel

(1912) used tissue and embryo extracts as cultural media to keep the fragments of

chick embryo heart alive.

In the late 1940s, Enders, Weller and Robbins grew poliomyelitis virus in culture

which paved way for testing many chemicals and antibiotics that affect

multiplication of virus in living host cells. The significance of animal cell culture

was increased when viruses were used to produce vaccines on animal cell cultures

in late 1940s.

For about 50 years, mainly tissue explants rather than cells were used for culture

techniques, although later after 1950s, mainly dispersed cells in culture were

utilized. In 1966, Alec Issacs discovered Interferon by infecting cells in tissue

culture with viruses. He took filtrates from virus infected cells and grew fresh

cells in the filtered medium. When the virus was reintroduced in the medium, the

cells did not get infected. He proposed that cells infected with the virus secreted a

molecule which coated onto uninfected cells and interfered with the viral entry.

This molecule was called “Interferon”.

Chinese Hamster Ovary (CHO) cell lines were developed during 1980s.

Recombinant erythropoietin was produced on CHO cell lines by AMGEN

(U.S.A.). It is used to prevent anaemia in patients with kidney failure who require

dialysis. After this discovery, the Food and Drug Administration (U.S.A) granted

the approval for manufacturing erythropoietin on CHO cell lines. In 1982, Thilly

and co-workers used the conventional conditions of medium, serum, and O2 with

suitable beads as carriers and grew certain mammalian cell lines to densities as

high as 5x106 cells/ml.

A lot of progress has been also made in the area of stem cell technology which

will have their use in the possible replacement of damaged and dead cells. In

1996, Wilmut and co-workers successfully produced a transgenic sheep named

Dolly through nuclear transfer technique. Thereafter, many such animals (like

sheep, goat, pigs, fishes, birds etc.) were produced. Recently in 2002, Clonaid, a

human genome society of France claimed to produce a cloned human baby named

EVE.

For animals, if the explant maintains its structure and function in culture it is

called as an ‘organotypic culture’. If the cells in culture reassociate to create a

three dimensional structure irrespective of the tissue from which it was derived, it

is described as a ‘histotypic culture’.

TYPES OF CELL CULTURES

Cell cultures are derived from either primary tissue explants or cell suspensions.

Primary cell cultures typically will have a finite life span in culture whereas

continuous cell lines are, by definition, abnormal and are often transformed cell

lines.

3. Primary Cell Culture

The maintenance of growth of cells dissociated from the parental tissue (such as

kidney, liver) using the mechanical or enzymatic methods, in culture medium

using suitable glass or a plastic container is called Primary Cell Culture. A

primary culture is that stage of the culture after isolation of the tissue but before

the first subculture. The primary cell culture could of two types depending upon

the kind of cell in the culture

1. Anchorage dependent/ Adherent cells - Cells shown to require attachment for

growth are set to be Anchorage Dependent cells. The Adherent cells are usually

derived from tissues of organs such as kidney where they are immobile and

embedded in connective tissue. They grow adhering to the cell culture.

2. Suspension Culture/Anchorage Independent cells - Cells which do not

require attachment for growth or do not attach to the surface of the culture vessels

are anchorage independent cells/suspension cells. All suspension cultures are

derived from cells of the blood system because these cells are also suspended in

plasma in vitro e.g. lymphocytes.

3.1 Isolation of the Tissue

Before working with animal or human tissue, it is essential to be sure that work

does not violate medical-ethical rules or the current legislation. For instance,

culturing of cells from living human embryos is prohibited by law in some

countries. Once you have made a choice of the tissue to be used for culturing.

You may sterilize the site with 70% alcohol and remove the tissue aseptically and

transfer it to balanced salt solution (BSS) or to a suitable culture medium. The

tissue may also be stored in a refrigerator before transferring it to BSS or to a

culture medium, because viable cells may be recovered from chilled tissue several

days after explantation. Different protocols are available for isolation of tissues

like mouse embryo, hen's egg, human biopsy material, etc.

3.1.1 Enzymatic disaggregation

Cell-cell adhesion in tissue is mediated by a variety of homotypic interacting

glycopeptides (cell adhesion molecules), some of which are calcium dependent

and hence are sensitive to chelating gent such as EDTA or EGTA. Intracellular

matrix and basement membranes contain other glycoproteins, such as fibronectin

and laminin which are protease sensitive and proteoglycans, which are less so but

sometime can be degraded by glycanases. The two important enzymes used in

tissue disaggregation are collagenase and trypsin.Crude trypsin is the most

common enzyme used in tissue disaggregation.

Use of Trypsin for disaggregation of tissue is called trypsinization.It may be of

two types;

a. Warm trypsinization

b. Cold trypsinization

Crude trypsin is the most common enzyme used for disaggregation. It is tolerated

by a variety of cells and is effective for many tissues. Its residual activity is

neutralized by the serum of the medium or by a trypsin inhibitor (e.g. Soybean

trypsin inhibitor), in the case of serum-free medium. The cells are exposed to the

warm enzyme (36.5°C) for a minimum period. The dissociated cells are collected

every half an hour. The trypsin is removed by centrifugation after 3-4 hours,

which is required for complete disaggregation. Cold trypsinization involves

soaking of tissue in trypsin at O°C to allow penetration of enzyme, followed by

incubation at 36.5°C for a shorter period.

3.1.2 Mechanical disaggregation

Enzymatic disaggregation is labour intensive and involves damage of cells.

Therefore, mechanical disaggregation of cells is sometimes preferred. In, this

method, tissue is carefully sliced and the cells which spill out are collected.

Alternatively the cells are either,

(i) pressed through the sieves of gradually reduced mesh, or

(ii) forced through a syringe and needle ,or even

(iii) Repeatedly pipetted.

Although the method may cause mechanical damage the cell suspension is more

quickly obtained than in the enzymatic disaggregation. Therefore, when the

availability of tissue is no limitation and the efficiency of yield unimportant,

mechanical disaggregation may be used to obtain good yield of cells in a shorter

time, but at the expense, of very much more tissue.

3.2 Separation of viable and nonviable cells

The dissociated cells obtained as above, usually described as primary cells, grow

well when seeded on culture plates at high density. These are adherent primary

cultures, but primary cultures can also be maintained in suspension.

In the first case (adherent culture), nonviable cells wilt be removed at the first

change of medium. In suspension, on the other hand, non viable cells are

gradually diluted out, when cell proliferation starts. However, non viable cells can

also be removed from primary disaggregate by centrifuging the cells on a mixture

of Ficoll and sodium metrizoate, when viable cells are collected from the interface

after centrifugation.

3.3 Subculture and its propagation

The need to subculture implies that the primary culture has increased to occupy

all of the available substrate. The first subculture gives rise to a secondary culture,

the secondary to a tertiary, and so on, although in practice, this nomenclature is

seldom used beyond the tertiary culture. Once a primary culture is subcultured , it

becomes known as a cell line. A Cell Line or Cell Strain may be finite or

continuous depending upon whether it has limited culture life span or it is

immortal in culture. On the basis of the life span of culture, the cell lines are

categorized into two types:

a) Finite cell Lines - The cell lines which have a limited life span and go through

a limited number of cell generations (usually 20-80 population doublings) are

known as Finite cell lines. These cell lines exhibit the property of contact

inhibition, density limitation and anchorage dependence. The growth rate is slow

and doubling time is around 24-96 hours.

b) Continuous Cell Lines - Cell lines transformed under laboratory conditions or

in vitro culture conditions give rise to continuous cell lines. The cell lines show

the property of ploidy (aneupliody or heteroploidy), absence of contact inhibition

and anchorage dependence. They grow in monolayer or suspension form. The

growth rate is rapid and doubling time is 12-24 hours.

Table-Some animal cell lines and the products obtained from them

Cell line Product

Human tumour Angiogenic factor

Human leucocytes Interferon

Mouse fibroblasts Interferon

4. Techniques of animal cell culture

The development of primary cultures and cell lines, a variety of tissues and

disaggregation methods are used to give good yield of separate cells. The tissues

needs to be obtained under aseptic and sterile conditions, using the ‘primary

explanation technique’ developed by Harrison (1907), Carrel (1912) and

others. The primary explantation technique is used for cultivation of pieces of

fresh tissue derived from an organism, and was almost the exclusive technique

used for animal tissue culture till about 1945. Different forms of primary

explantation techniques are still widely used and will continue to be used for a

very long time.

These techniques differ only in the type of vessels (flasks, test, tubes, etc.) used

for growing the tissue, but are uniform in principle. The primary explantation

techniques are also used for embryo and organ culture, but are variously modified

to become specialized techniques are classified into the following:

(i) Slide cultures (ii) Carrel flask cultures

(iii) Roller test tube cultures.

4.1 Slide or coverslip cultures

In this technique, slides or cover slips are prepared by placing a fragment of tissue

(explantation) onto a coverslip, which is subsequently inverted over the cavity of

a depression slide. This is the oldest method of tissue culture and is still quite

widely used. This has a number of advantages and disadvantages. The application

of slide culture is limited but it may be very useful for morphological studies

through the use of time-lapse cinemicrographic investigations. There are several

methods for preparation of slide culture.

4.1.1 Single coverslip with plasma clot

This technique developed by Harrison (1907) has been most commonly used

during the last more than fifty years and includes the following steps;

(i) prepare medium in two parts, one containing 50% plasma in BSS (balanced

salt solution) and the other containing 50% embryo extract in serum;

(ii) under sterile and aseptic conditions, using a capillary pipette, place one drop

of plasma containing solution in the centre of each of one or more cover slips

(22mm)

(iii) transfer a fragment (one or two pieces) of tissue;

(iv) add the embryo extract containing solution and mix thoroughly before

clotting starts and then locate the explant;

(v) place two small spots of petroleum jelly (using a glass rod) near the concavity

of a depression slide and invert this slide over the coverslip; apply gentle pressure,

so that jelly sticks to coverslip;

(vi) allow culture medium to clot;

(vii) turn over the slide and seal the margins of coverslip with paraffin;

(viii) label and incubate at 37°C.

Preparation of Single Coverslip Cultures

A. Place one drop of Plasma

B. Add an Explant

C. Add a Drop of Embryo Extract

D. Spread Medium and Place Explant

A

B C D

G. Seal with Paraffin

F. Invert on a Cover Slip E. Place Two Spots Petroleum Jelly on Slide

G F E

4.1.2. Double Coverslip with Plasma Clot

Maximov's Double Coverslip Method for Preparation of Slide Cultures Top View Side View

This technique was developed by Maximov. It includes the following steps;

(i) a small drop of BSS is paced on a large coverslip (40mm);

(ii) a square or round coverslip (22mm) is placed over BSS in the centre of large coverslip.

These two steps are then followed by the steps listed above for single coverslip method. A

large depression slide is used and the entire preparation is attached to it by petroleum jelly

and wax in such a way that the small coverslip is not in contact with the slide at any point.

4.1.3. Single coverslip with liquid medium (lying and hanging drop cultures).

Following steps are involved in this method:

(i) suitably prepared explants are placed in culture medium in a watch glass;

(ii) the explants are drawn into the tip of a capillary pipette, and one explant is

deposited in the centre of each coverslip;

(iii) the liquid medium can be spread out in a very thin circular film with the

explant protruding above the surface;

(iv) a depression slide with petroleum jelly is applied immediately and

preparation turned over with a quick flip to prevent the fluid from running out of

the crevice between the slide and the coverslip;

(v) the coverslip is sealed and the slide incubated at 37°C, upright or inverted; the

tissue grows on the coverglass.

Advantages Disadvantages

1. It is simple and relative

inexpensive.

2. Cells in living state are

spread out in a manner

suitable for microscopy and

photography.

3. Cells grow directly on

coverslip and can be fixed

and stained to make

permanent slides

Supply of oxygen and nutrients

is rapidly exhausted, so that the

medium quickly becomes

acidic and requires transfer of

rapidly growing tissue.

Sterility can not be maintained

for a long period

Only very small amounts of

tissues can be cultured.

4.1.4. After-care of slide cultures

Single coverslip cultures are very useful for short-term studies but they are

difficult to handle subsequently except by a process of transfer. Therefore, double

coverslip method is recommended, whenever it is desirable to leave the explant in

its original location to obtain long-term tissue cultures. This would require

washing, feeding, patching and transfer of cultures.

a.) Washing and feeding double coverslip cultures

Washing and feeding involve the following steps:

(i) remove seal using razor blade and remove large coverslip along with small one

still attached to it and flip it over, so that the culture remains attached to small

coverslip and is uppermost in orientation:

(ii) detach small coverslip from the large coverslip using needle and forceps and

transfer it along with tissue to a Columbia staining dish (watch glass or Petri dish

may also be used) containing balanced salt solution (BSS);

(iii) remove small cover slip treated as above (one at a time) and place it again on

the large coverslip with culture up; while removing small coverslip from dish with

BSS, the amount of BSS brought with it may be controlled by the rate of removal

(too much liquid will interfere in feeding operation and too little will allow air

bubbles);

(iv) feed the culture by adding a drop of feeding solution (e.g. BSS : serum: 50%

embryo extract = I: I: 1) to the small coverslip;

(v) Attach a clean depression slide, using petroleum jelly, as done earlier.

b.) Patching the plasma clot in slide cultures

If there is evidence of liquefaction, plasma clot should be patched as follows;

(i) wash small coverslip with culture as above;

(ii) in a watch glass take two drops of a mixture of plasma and BSS and to this

add two drops    of a mixture of serum and embryo extract; mix his quickly and

place a drop on each washed coverslip having a culture;

(iii) a clean depression slide with petroleum jelly is then attached, as usual.

c.) Transfer of slide culture

The coverslip cultures may need to be transferred using the following steps:

(i) remove and wash coverslip with culture, and using a Bard-Parker knife, cut

through the outgrowth;

(ii) the square tissue may be cut into two or four pieces, each transferred to new

coverslip and treated as a new explant.

4.2 Flask cultures

The main use of flask cultures is in the establishment of a strain from fresh

explants of tissue. A good Carrel Flask has excellent optical properties for

microscopic examination, even though polystyrene culture flasks can also be used

provided they have a wide neck for handling the explants. The flask technique has

the following advantages:

(i) tissue can be maintained in the same flask for months or even years;

(ii) large number of cultures can be easily prepared and large amount of tissue can

be maintained for a considerable period of time.

Preparation of flask cultures.

Following steps are involved in the preparation of flask cultures:

(i) place upto six D3.5 Carrel Flasks in a rack with their necks flamed and

pointing to the right;

(ii) place a drop of plasma on the floor of flask and spread this plasma out in a

circle;

(iii) with the help of spatula, transfer the desired number of explants to the plasma

and allow clotting to occur;

(iv) after the plasma clots and explants fixed in position, add extra medium; for

thick clots 1.2ml of dilute plasma and for thin clots 1.2ml of dilute serum is added

instead of plasma; the whole thing is left for clotting;

(v) the flasks are gassed with gas phase (5% CO2 in air).

There are two types of flask techniques:

(i) Thick clot cultures', which allow rapid growth suitable for short-term cultures

and

(ii) Thin clot cultures which can be maintained for a considerable period of time.

Preparation of flask cultures

A. Add plasma to flask B. Spread with spatula

D. After clotting add medium C. place explant