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2. INTRODUCTION TO PLANT TISSUE CULTURE:
 It is the process of producing plants from tissue of the desired plant in an artificial
nutrient medium under controlled environment.
 The plants so grown would be exactly similar to the mother plant in all aspects.
 The science of plant tissue culture takes its roots from path breaking research in
botany like discovery of cell followed by propounding of cell theory. In
1839, Schleiden and Schwann proposed that cell is the basic unit of organisms.
They visualized that cell is capable of autonomy and therefore it should be possible
for each cell if given an environment to regenerate into whole plant. Based on this
premise, in 1902, a German physiologist, Gottlieb Haberlandt developed the
concept of in vitro cell culture. He isolated single fully differentiated individual
plant cells from different plant species like palisade cells from leaves of Laminum
purpureum, glandular hair of Pulmonaria and pith cells from petioles of Eicchornia
crassiples etc and was first to culture them in Knop’s salt solution enriched with
glucose. In his cultures, cells increased in size, accumulated starch but failed to
divide. Therefore, Haberlandt’s prediction failed that the cultured plant cells could
grow, divide and develop into embryo and then to whole plant. This potential of a
cell is known as totipotency, a term coined by Steward in 1968.

3. STERILIZATION TECHNIQUES:
 Sterilization Methods Used in Tissue Culture
Laboratory - All the
materials, e.g., vessels, instruments, medium, plant
material, etc., used in culture work must be freed
from microbes. This is achieved by one of the
following approaches:
(i) dry heat treatment,
(ii) flame sterilization,
(iii) autoclaving,
(iv) filter sterilization,
(v) wiping with 70% ethanol, and
(vi) surface sterilization.

4. EXPERIMENTS:
Protocol.1- Tissue Culture Media Preparation
PRINCIPLE:
Murashige and Skoog medium or (MSO or MS0 (MS-zero)) is a plant
growth medium used in the laboratories for cultivation of plant cell
culture. MSO was invented by plant scientists Toshio Murashige and
Folke K. Skoog during Murashige's search for a new plant growth
regulator. It is the most commonly used medium in plant tissue
culture experiments. A series of experiments demonstrated that
varying the levels of these nutrients enhanced growth substantially
over existing formulations. It was determined that nitrogen in
particular enhanced growth of tobacco in tissue culture.

5. Protocol-2- Explant Preparation and Surface
Sterilization
PRINCIPLE:
Surface sterilization treatments applied on the explants were:
Dipping in Ethyl alcohol 70% for 5 minutes (mins). (T1)
Dipping in Chlorox 3% (commercial sodium hypochlorite, active
ingredients 5.2%) for 20 mins. (plus Tween 20). (T2)
Dipping in undiluted Chlorox for 20 mins. (plus Tween 20). (T3)
Dipping in Ethyl alcohol 70% for 5 mins. then in Chlorox 3% (plus
Tween 20) for 20 mins. (T4)
dipping in Ethyl alcohol 70% for 5 mins. then in Mercuric
chloride 0.3% for 5 mins. (T5) After each treatment explants
were rinsed three times in autoclaved distilled water.

6. Protocol-3-EMBRYO CULTURE:
PRINCIPLE:
Organogenesis is the formation of individual organs such as shoots and roots either
directly on the explant in which pre-formed meristem are lacking or the meristems
develop de novo from callus. Although callus is an actively growing undifferentiated
mass of cells, differentiation can take place at random, but may be associated with
centres of morphogenesis, which can give rise to organs such as shoots, roots and
embryos. To induce shoot organogenesis from the callus, concentration of growth
regulators is varied in the medium. Cytokinins such as BAP and kinetin promote
shoot organogenesis.
RESULT:

7. Protocol-4- Culture of Anther for Production of
Androgenic Haploids
PRINCIPLE:
Anther and Microscope Culture - One of the very popular methods for production of
haploids is through culturing anthers or microspores on artificial culture medium.
This leads to the growth of microspores into saprophytes. After the initial reports of
successful production of haploids from anther culture in Datura (Guha and
Maheshwari, 1966, 1967), haploids have been obtained in more than 150 species
belonging to 23 families of angiosperms (Maheshwari et a1., 1980). These include a
wide variety of economically important species.
More often, anthers rather than microspores are cultured, since the extraction and
culture methods for microspores differ and have been successful only in a few species
(Datura inoxia, Nicotiana sylvestris, N. tabacum, Oryza sativa, etc.).

8. Protocol-5-Meristem culture
PRINCIPLE:
Organogenesis is the formation of individual organs such as
shoots and roots either directly on the explant in which
pre-formed meristem are lacking or the meristems
develop de novo from callus. Although callus is an actively
growing undifferentiated mass of cells, differentiation can
take place at random, but may be associated with centres
of morphogenesis, which can give rise to organs such as
shoots, roots and embryos. To induce shoot
organogenesis from the callus, concentration of growth
regulators is varied in the medium. Cytokinins such as
BAP and kinetin promote shoot organogenesis.

9. Protocol-6- Meristem tip culture for production of
Virus –free Plants
 Theme: Shoot apical meristem lies in the 'shoot tip' beyond the
youngest leaf or first leaf primordium ; it measures upto about 100 µm in
diameter and 250 µm in length. Thus a shoot-tip of 100-500 11m would
contain 1-3 leaf primordia in addition to the apical meristem.In
practice, shoot-tips of up to 1 mm are used when the objective is virus
elimination. Shoot-tip culture is widely used for rapid clonal propagation
for which much larger, e.g., 5-10 mm, explants are used. Therefore, most
cases of meristem culture are essentially shoot-tip cultures. Nodal
explants of various sizes are also commonly employed for rapid clonal
propagation. When the objective is vegetative propagation, the size of
shoot-tip used for culture is not important. The upper few millimeters
(ca.5-6mm) in a shoot apex is considered to be free from virus in those
plants which are systematically infected. Due to active cell division (faster
than virus multiplication), absence of vascular connection and high auxin
concentration the shoot meristem remains virus free. If the meristem tip
is used as an explant for propagation, virus free plats can be obtained.

10. Protocol-7- Induction of Somatic
Embryogenesis (Monocot and Dicot System)
 In somatic embryogenesis the embryo arises from somatic cells, tissue or organs under in vitro
conditions. Somatic embryo is a bipolar structure and has no vascular connection with the
maternal cultured explant. The somatic embryos are functionally equivalent to zygotic embryos
but the process of embryogeny is different. It induces four developmental phases i.e.0, 1, 2, and
3 phases.
Phase 0: In phase 0 the competent single cells (state 0 cells) form embryogenic cell clusters
(state 1 cells) in the presence of auxin. During this phase the cell cluster formed from single
cells gains the ability to develop into embryos when auxin (a PGR used to induce somatic
embryogenesis) is removed from the medium giving rise to state 1 clusters.
 Phase 1: Phase 1 is induced by transfer of state 1 cell clusters on to auxin free medium. In this
phase the cell cluster proliferate slowly and undifferentiately.
 Phase 2: In phase 2, rapid cell division occurs in certain parts of cell clusters leading to
formation of globular embryos.
Phase 3: In phase 3, globular embryos develop into plantlets via heart shaped, torpedo shaped
and cotyledonary stage embryos.
Somatic embryos could be induced either directly from the explant tissue in the absence of
callus formation (direct somatic embryogenesis) or via the callus from the explant (Indirect
somatic embryogenesis). Embryogenic cells are small, isodiametric in shape, filled with dense
cytoplasm and have a conspicuous nucleus. In comparison to this, on-embryogenic cells are
relatively large, vacuolated and lack dense cytoplasm.

11. Protocol-8- Protoplast Isolation, Culture and
Regeneration
PRINCIPLE:
Mechanical Method of Isolation of Protoplast - In
mechanical method, cells are kept in a suitable plasmolyticum
(in plasmolysed cells, protoplasts shrink away from cell wall)
and cut with a fine knife, so that protoplasts are released from
cells cut through the cell wall, when the tissue is again
deplasmolysed. This method is suitable for isolation of
protoplasts from vacuolated cells (e.g. onion
bulbs, scales, radish roots). However, this method gives poor
yield of protoplasts and is not suitable for isolating protoplast
from meristematic and less vacuolated cells. The mechanical
method, though, was used as early as 1892, is now only rarely
used for isolation of protoplasts.

12. Protocol-9- Microculture Chamber Technique for
Single Cell Isolation
PRINCIPLE:
 Single Cell Culture - Establishment of a single cell culture provides an
excellent opportunity to investigate the properties and potentialities of
plant cells.
Such studies contribute to our understanding of the interrelationships and
complementary influences of cells in multicellular organisms. Several
workers have successfully isolated single cell division and even raised
complete plants from single cell cultures.
Using cell cultures in studies designed to describe the pathways of cellular
metabolism was another aspect that initially attracted the attention of
plant biologists. It was soon realized that single cell systems have great
potential for crop improvement.
 The microculture chamber technique was first developed by Jones et al.
(1960) and later was used by Vasil and Hildebrandt (1965) after some
modifications. This method consists of culturing 30-50μ/ of medium
containing one or more protoplasts on a microscope slide enclosed by a
cover glass resting on two other cover glasses placed on either side of the
drop. The cultures are sealed with sterile paraffin oil and incubated in light
at 23-25º C.

13. Protocol-10- Encapsulation of somatic embryos / shoot buds for
Production of Synthetic seeds
PRINCIPLE:
Synthetic seeds are prepared by encapsulating the somatic embryos obtained from
tissue culture in a protective jelly capsule, which is usually prepared with sodium
alginate. From the synthetic seeds whole plant can be recovered under in vitro,
greenhouse and field conditions. Alternatively, shoot buds, conservation and
maintenance of rare and threatened species.
Synthetic Seeds - In the conventional plant tissue culture for clonal propagation,
storage and transportation of propagules for transplantation is a major problem.
To overcome this problem, in recent years the concept of synthetic or artificial seeds
has become popular, where somatic embryos are encapsulated in a suitable matrix
(e.g. sodium alginate), along with substances like mycorrhizae, insecticides,
fungicides and herbicides. In. India, this technique of synthetic seeds is being
standardized and practiced for sandalwood and mulberry at BARC (Bombay) under
the leadership of Dr. P.S. Rao. Synthetic seeds have many advantages including the
following:
(i) they can be stored upto a year without loss of viability;
(ii) they are easy to handle, and useful as units of delivery;
(iii) they can be directly sown in the soil like natural seeds and do not need
hardening in greenhouse.
The only limitation of synthetic seeds, is the high cost of their production. However,
this may go down in future, so that these synthetic seeds will become popular at the
commercial scale in due course of time.

14. Protocol-11- Establishment of Cell Suspension
Culture
PRINCIPLE:
 Suspension Culture Growth and Subculture - Cell suspensions are
clonally maintained by the routine transfer (subculture) of cells in the early
stationary phase to a fresh medium. During the incubation period the
biomass of the suspension cultures increases due to cell division and cell
enlargement. This continues for a limited period since the viability of cells
in suspension after the stationary phase decreases due to the exhaustion of
some factors or the accumulation of toxic substances in the medium. At this
stage an aliquot of the cell suspension with uniformly dispersed free cells
and cell aggregates is transferred to afresh liquid medium of the original
composition. The timing of a subculture is very important.
 Types of Suspension Cultures - There are mainly two types of suspension
cultures, batch cultures and continuous cultures. Batch cultures are
maintained by propagating a small aliquot of the inoculum in the moving
liquid medium and transferring it to a fresh medium at regular intervals.
Generally cell suspensions are grown in flasks (100-250 ml) containing 20-75
ml of the culture medium: The biomass growth in batch cultures follows a
fixed pattern.

15. Protocol-12- Culture of single Cells (Bergmann’s cell plating
technique)
 PRINCIPLE: PRINCIPLE:
 In this technique, free cell are suspended in a liquid medium (if cell aggregates are
there, the culture is filtered), and a culture medium with agar (0.6-1%) is cooled and
maintained at 35ºC in a water bath. Equal volumes of these liquid and agar media are
mixed and rapidly spread in a Petri dish, so that cells are evenly distributed in a thin
layer, after solidification. The Petri dishes are sealed with parafilm and examined with
inverted microscope to mark single cells (marking is done on outer surface of the
dish). The Plates are incubated in dark at 25°C and cell colonies developing from
marked single cells are used to obtain single cell cultures. Various other methods (e.g.
filter paper raft technique; microchamber technique) have also been developed to
grow individual cells (Bhojwani and Razdan, 1983)
In this technique, free cell are suspended in a liquid medium (if cell aggregates are
there, the culture is filtered), and a culture medium with agar (0.6-1%) is cooled and
maintained at 35ºC in a water bath. Equal volumes of these liquid and agar media are
mixed and rapidly spread in a Petri dish, so that cells are evenly distributed in a thin
layer, after solidification. The Petri dishes are sealed with parafilm and examined with
inverted microscope to mark single cells (marking is done on outer surface of the
dish). The Plates are incubated in dark at 25°C and cell colonies developing from
marked single cells are used to obtain single cell cultures. Various other methods (e.g.
filter paper raft technique; microchamber technique) have also been developed to
grow individual cells (Bhojwani and Razdan, 1983)

16. Protocol-13- Maintenance of cell line
PRINCIPLE:
Maintenance of Cultures Cell Lines - When a primary culture produced on
a substrate or in suspension has increased to the extent that all the available
substrate is occupied or the medium largely consumed, there arises the
need to subculture it. From a very heterogeneous primary culture
containing many types of cells derived from the original tissue, during sub
culturing (passages or transfer) a more homogeneous 'cell line' emerges.
The culture now called a cell line can be propagated, characterized and
stored. The potential increase in cell number and uniformity of cells, open
up a much wider range of possibilities. The term' cell line implies the
presence of several cell lineages either similar or distinct. Among these cell
lineages, if a particular cell lineage has specific properties, which are
identified in bulk of the cells of that culture, it is described then as a 'cell
strain'. A "cell line" or 'cell strain' may be finite or continuous, depending
upon whether it has limited culture life spans or it is immortal in culture.
Finite cell lines grow upto 20-80 population doublings before I extinction.
Some commonly used cell lines and cell strains and a comparison of the
properties of mite and continuous cell lines is presented