Cell culture

Introduction to
CELL CULTURE
Dr SV Suresh Kumar
Professor of Pharmacognosy
CES College of Pharmacy, Kurnool
Andhra Pradesh
1Dr Suresh Solleti, CESCOP

CELL CULTURE
 Culturing of single cells in vitro in an artificial nutrient
media is known as cell culture.
 Cell cultures are carried out to investigate potentialities
and properties of individual cells.
 Isolation of single cells:
1. From intact organ
2. From callus

Isolation of single cells:
 This can be achieved from intact organs and from the
callus.
 From intact organ: Mechanical and enzymatic methods
are used to isolate the cells from intact organs.
 The plant part preferably leaf tissue is subjected for grinding in
a mortar with pestle along with addition of grinding medium
consisting of sucrose 20 μmol . MgCl2 10 μmol,Tris
Hydrochloride buffer 20 μmol (pH 7-8).The resulting mixture is
subjected for centrifugation and cells are taken up into a liquid
media.
 The plant tissue is treated with an enzyme known as
macerozyme along with 1% potassium dextran sulphate, which
causes the digestion of middle lamella there by liberating the
cells.

 The enzyme also causes the weakening of cell wall thus
creating the problems associated with osmosis. Hence the cells
are provided with osmotic protection with a substance known
as osmoticum.
 Commonly used osmoticums are sorbitol 450-800 mmol/l, KCl
335 mmol/l, Magnesium sulphate 40 mmol/l.
 From Callus:
 Callus preferably friable callus upon agitation in suitable liquid
media results in the dissociation of the cells from callus.
 The agitation will also helps in uniform distribution, breaking
up of lumps and gaseous exchange between culture air and
atmosphere.

Culturing of cells:
 The cells obtained in the isolation will be subjected for
culturing by which the cells build up quickly.
 This can be achieved with batch and continuous cultures.
 Batch Cultures:
 The inoculum from the suspension culture is used to initiate the
batch cultures.
 The duration of the lag phase depends on the growth phase of
the cells in stock culture at the time of culturing or sub culturing
& size of the inoculum i.e cell density.
 An initial cell density of 9-15 x 103 cells /ml is the best.
 Low cell density cell cultures will not grow unless the medium is
enriched with metabolites necessary to grow cells.

 Batch cultures are characterised by a constant change in the
pattern of cell growth and metabolism and also by change in
the composition of the medium.
 In these cultures, exponential growth with constancy of cell
doubling time may be achieved, but there is o period of
study state growth in which relative concentrations of
metabolites and enzymes are also constant.

 Continuous cultures:
 The continuous cultures are also known as mass cultures.
 In these cultures the cell are never allowed to enter in to the
stationary phase.
 These are of closed continuous and open continuous cultures.
 Closed continuous culture-they are characterised by the
addition of fresh medium and outflow of equal amount of old
medium. Cell biomass continues to increase as the growth
proceeds.
 Open continuous cultures are characterised by addition of fresh
medium while harvesting the equal amount of the old medium.
This allows indefinite maintenance of cultures at constant and
sub maximal growth.

 Open cultures are of chemo static and turbidistatic types
depending upon by which the old media is replaced with fresh
media.
 As in culture media i.e in actively growing suspension cultures,
the inorganic phosphate is rapidly utilised and consequently it
soon becomes a limiting factor.The amount of inorganic
phosphate is the basis for chemostatic methods.
 In the turbidistatic methods the turbidity of the culture is taken
into consideration.The turbidity is preselected on the basis of
biomass density and can be maintained by intermittent flow of
medium and washout of cells.

Assessment of cell growth:
 Cell counting : In the known volume of cell culture the cells
are counted using haemocytometer.
 To facilitate the cell counting, the cell aggregates are treated
with 5-8% chromic acid or 0.1-0.25% pectinase solutions, which
causes the breaking up of lumps.
 After the treatment the cells are washed with saline then the
cells are counted.

 Packed cell volume:
 To determine the PCV a known volume of the uniformly
dispersed culture is transferred to a 15 ml graduated
centrifuge tube, then subjected for centrifugation at 200xg
for 5 min.
 The sediment formed is measured and expressed as ml of
pellet for ml of culture.
 Cell wet weight:
 To determine the cell wet weight a known volume of the
uniformly dispersed culture is collected on pre weighed
circular filter of nylon fabric supported in hartley funnel.
 Then the cells are washed with water to remove the
medium, under vacuum and weighed and represented as
grams of cells/ml of culture.

 Cell dry weight:
 To determine the cell dry weight a known volume of the uniformly
dispersed culture is collected on pre weighed circular filter of
nylon fabric supported in hartley funnel.
 Then the cells are washed with water to remove the medium,
under vacuum and dried at 60°C for 12 hours and weighed & and
represented as grams of cells/ml of culture.

 Non invasive method:
 In this method, the culture flask is fitted with ruler, and is
tilted at an angle of 30° to 60° for 5 min and the height of the
sediment is reordered. Change in the height of the sediment
with the age of culture would represent the change in fresh
weight of cells.

Assessment of cell viability:
 Phase contrast microscopy:
 Using phase contrast microscopy, the cytoplastic
streaming and the presence of healthy nucleus will be
observed.Thus the viable cells are counted for the given
volume of culture.

 Reduction of tetrazolium salts:
 When the cell cultures are incubated with 2,3,5 triphenyl
tetrazolium chloride (TTC), the viable cells converts theTTC
into a red colored substance known as 1,3,5 triphenyl
Formazan, which is estimated spectrophotometrically.

 Fluorescein diacetate (FDA) method:
 Cell cultures are incubated with 0.5% FDA for 5 min.
 FDA being non polar and non fluorescing, enters the cells
and is cleaved by esterase activity in the living cell resulting
into polar Fluorescein.
 Since Fluorescein is not freely permeable across the plasma
membrane, it accumulates mainly in the cytoplasm of intact
cells, thus those cells exhibit green colour fluorescence.

 Evans Blue method:
 Cell cultures are incubated with 0.025% Evans blue for 5 min.
 Evans blue dye has been used as a viability assay on the basis
of its penetration into non-viable cells i.e dead and damaged
cells.
 The dead or damaged cell takes the blue colour.

Culturing of single cells
 Bergmann’s cell plating technique:
 The most popular technique of single cell culture is
Bergmann's cell plating technique.
 Suspension cultures which carry cell aggregates in
addition to free cells should be filtered through a sieve
which would allow only single cells and small cell
aggregates to pass through.The large cell aggregates are
discarded and only the fine suspension is plated.
 Free cells are suspended in a liquid medium at a density
twice the finally desired plating cell density.
 Melted agar (0.6-1%)-containing medium of otherwise the
same composition as the liquid medium is cooled and
maintained at 35°C in a water bath.

 Equal volumes of the two media are mixed and rapidly
spread out in Petri dishes in such a manner that the cells
are evenly distributed and fixed in a thin layer (1 mm
thick) of the medium after it has cooled and solidified.
 The dishes are sealed with parafilm.
 The plates may be observed under an inverted microscope
and single cells marked on the outside of the plate by a
fine marker to ensure the isolation of pure single cell
clones.
 If the plating cell density is determined at the time of
culturing and a known volume of suspension is transferred
to each plate it should be possible to make a quantitative
assessment of plating efficiency using the formula:
PE= No. of colonies per plate at the experiment
No of cell units initially per plate X100

 Usually plating at cell densities of less than 104 to 105 cells/ml
yields high plating efficiency.

Single cell techniques:
 Filter paper raft nurse technique:
 It involves cultivating individual cells on top of an actively
growing callus separated by a piece of filter paper.
 In practice, individual cells are isolated from suspension
cultures or a friable callus with the aid of a micropipette or
microspatula.
 Several days before cell isolation, sterile 8 x 8 mm squares
of filter paper are placed aseptically on top of the
established callus of the same or different species.
 The filter paper is wetted by liquid and nutrients from the
nurse tissue piece.The isolated single cell is placed on the
wet filter paper raft.
 After a macroscopic colony develops from the cell it is
transferred to agar medium for further growth and
maintenance of the cell clone.

 An isolated cell which generally fails to divide when plated
directly on the medium used for callus cultures is able to
divide under the nursing effect of the callus.
 Apparently, the callus supplies the cell with not only the
nutrients from the culture medium but something more that
is critical for cell division.The cell division factor(s) can
diffuse through the filter paper.

 Micro chamber technique:
 In this method a drop of the medium carrying a single cell is
isolated from suspension cultures, placed on a sterile
microscope slide and ringed with sterile mineral oil.
 A drop of oil is placed on either side of the culture drop and a
coverglass placed on each drop.
 A third coverglass is then placed on the culture drop bridging
the two coverglasses and forming a microchamber to
enclose the single cell aseptically within the mineral oil.
 The oil prevents water loss from the chamber but permits
gaseous exchange.
 The whole microchamber slide is placed in a petri-dish and
incubated.When the cell colony becomes sufficiently large
the coverglass is removed and the tissue is transferred to
fresh liquid or semi-solid medium.
 The microchamber technique permits regular observation of
the growing and dividing cell.

 Micro drop technique:
 The technique requires a specially designed cuprak dishes
which have smaller outer chamber and larger inner
chamber.
 The inner chamber carries numerous numbered wells,
each with a capacity for 0.25 to 25 µl droplet of nutrient
medium.
 Individual cell is transferred to each well of the chamber
along with conditioned medium.

 The outer chamber is filled with sterile water to maintain
humidity inside the dish.
 After covering with lid, the dish is sealed with parafilm
and maintained at optimal light and temperature
conditions.
 When cell group develops it is transferred to medium to
form callus.

Types of PTC
 Root tip cultures:
 From the root cultures it is possible to produce genetically
uniform roots with in few days.
 It can be used for the commercial production of useful
substances.
 The seed are surface sterilised and transferred to a petri-dish
lined with whatmann no. 1 filter paper and moistened with
distilled water.
 The petri-dish is wrapped in aluminium foil and incubated
for 5 days in dark at 25°C, which results in the germination of
seeds with development of root tip(s).
 About 10 mm long root tips are taken and transferred to
flasks containing culture medium and incubated for 10 days
at 25°C.This results in production of root and many lateral
roots

 Shoot tip culture
 Microbe free plants can be best produced through shoot tip
culture.
 One important method of vegetative propagation.
 About 10mm long shoots are removed from healthy plants
and these explants are subjected to surface sterilization in
hypochlorite solution for 15 min.
 Then the distilled water washed explants are placed on
petri-dish lined with moistened filter paper for dissecting the
shoot apex.
 The terminal 0.5mm of the shoot apex is carefully excised
and is placed culture vessel containing the cultured medium.
 Then the vessels are covered non absorbent cotton followed
by aluminium foil and subjected for incubation at 27°C with
light intensity of 1000 lux at 70% relative humidity for 16
hours.

 After formation of the callus, the callus transformed on to
the medium with high cytokinin concentration to induce
shoots followed by high auxin concentration to induce roots
formation.
 Alternatively the tips of the shoots can be cultured on the
medium, resulting in the production of clumps of shoots
from either axillary or adventitious buds.This method is
known as micro propagation.

Anther culture
 When the pollen/anthers of the correct stage are
collected and grown on suitable media, it is called as
anther culture.
 It is used to produce haploid plants.
 The selected flower buds are subjected to surface
sterilisation with alcohol or hypochlorite solution for about
10-30 min.
 They are rinsed repeatedly with sterile distilled water.
 By using sterile forceps and a dissecting needle, the
anthers are carefully excised from the flower buds.
 The filaments must be removed prior to culture as they
form callus at the cut ends.
 The excised anthers may be cultured on solidified culture
medium or placed on a filter paper bridge over a liquid
medium.

 Then is incubated at 25°C, in presence or absence of light.
 The plantlets are formed after 4-5 weeks of inoculation.
Many plantlets are produced from single anther.
 These plantlets are carefully separated quite early and
cultured on fresh root inducing medium containing agar to
develop roots, thus forming entire plants.
 Alternatively the anthers are allowed to produce callus
and from which shoots and roots respectively.

1. Pollen on media
2. Callus formation
3. Shoot initiation
4. Root initiation
5. Whole plant.

Embryo culture
 Culturing of the embryo excised from the seeds on
sterilised culture medium is called embryo culture.
 The selected fruit is surface sterilised for 15 min with
alcohol and then washed thoroughly with distilled water.
 The fruit is cut and the seed or ovule is taken out with a
sterile forceps and placed on a sterile petri dish.
 The seed or ovule is cut open and the embryo is
separated.
 The excised embryo is immediately placed on the nutrient
medium of culture vessel under aseptic conditions.
 The explanted embryos on culture vessels are kept at
room temperature in diffused light.
 This resulting in the production of seedlings which are
transferred to sterile growth promoting medium for
further development.

 Mature embryos are usually selected as the very small
globular embryos require a delicate balance of hormones.
 The dormancy period of seeds can be shortened.
 By the embryo culture it is possible to know the viability of
seeds.
 By this culture it is possible to know the nutritional
requirements of the embryos at various developmental
stages.
 Used to propagate rare
plants.

Cell culture

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