Plant tissue culture

Here, all information about Plant Tissue Culture
HISTORY OF PLANT TISSUE CULTURE
THE TECHNIQUE OF PLANT TISSUE CULTURE
Plantlet Regeneration and Transfer to Soil
A Classification of Tissue Culture Techniques
EMBRYO CULTURE
MERISTEM CULTURE
ANTHER OR POLLEN CULTURE
TISSUE AND CELL CULTURES
SOMATIC HYBRIDIZATION

1. Plant tissue culture

Plant tissue culture broadly refers to the cultivation in vitro of all plant parts, whether a single cell, a
tissue or organ in a vessels using artificial media under aseptic conditions.



This new technique has enabled us to increase the knowledge in the following field of
studies.
 Totipotency, nutrition, metabolism, division, differentiation and preservation of plant
cells.
 Morphogenesis and plant regeneration from individual cells or tissues through the
process namely organogenesis or somatic embryogenesis.
 Variations generated through in vitro culture.
 Evolution of haploids through anther and pollen culture including ovule culture.
 Wide hybridization programmes through ovule, ovary and embryo cultures to overcome
both pre zygotic and post zygotic sterility mechanisms
 Micro propagation of plant materials
 In vitro selection of mutants tolerant to biotic and abiotic stresses.
 In vitro culture and secondary metabolite biosynthesis.
 Plant genetic engineering through in vitro culture methods and DNA transfer technique.

Thus plant cell, tissue and organ culture permeates plant biotechnology and cements together its
various aspects like Physiology, Biochemistry, Genetics and Cell Biology.
Like other subjects, plant cell and tissue culture has its own origin and development. The
chronology of major events in this field is presented for the benefit of the new entrants into this
field.

2. HISTORY OF PLANT TISSUE CULTURE

The in vitro techniques were developed initially to demonstrate the totipotency of plant cells
predicted which are characteristic of zygotes, i.e., their ability to develop into complete plants.
Efforts to demonstrate that is, tissue culture. This was made possible by brilliant contributions
from R.J. Gautheret in France and by Haberlandt in 1902. Totipotency is the ability of plant
cells to perform all the functions of development, totipotency led to the development of
techniques for cultivation of plant cells under defined conditions, P.R. White in U.S.A. during
the third and the fourth decades of 20th century. Shoot bud regeneration tobacco suspension
cultures were reported by White in 1939, but the first plant from a mature plant cal was
regenerated in 1959 by Braun. Most of the modern tissue culture media have been derived from
work of Skoog and coworkers during 1950s and 1960s. The first embryo culture was done by
Hannin in 1904 who cultured mature embryos of some crucifers. This technique was soon
applied by Laibach in 1925, to recover hybrid progeny from an interspecific cross in Linum.
Haploid plants from pollen grains were first produced by Maheshwari and Guha in 1964 by
culturing anthers of Datura. The technique has been further developed by many workers, more
notably by J.P. Nitsh, C. Nitsh and coworkers, who showed that isolated microspores of tobacco
produce complete haploid plants. Plant protoplasts are naked cells from which cell wall has
been removed. In 1960, Cocking produced large quantities of protoplasts by using cell wall
degrading enzymes. The techniques of protoplast production have now been considerably
refined, and whole plants regenerate readily from protoplasts. Protoplasts of different plant
species have been fused to obtain somatic hybrid plants. Thus within a brief period, the
techniques have made great progress: from the sole objective of demonstrating totipotency of
plant cells, the techniques now find application in both basic and applied researches. Our
objective here is to examine their present applications and future possibilities for improvement of
crop plants.
THE TECHNIQUE OF PLANT TISSUE CULTURE
The tissue culture technique aims primarily to achieve the following two objectives:
(1) To keep the plant cells and organs free from microbes, and
(2) To ensure the desired development in the cells and organs by providing suitable nutrient
media and other environmental conditions.
Microbes can be managed by using modern equipments and careful handling during various
operations. The second objective remains an area of active research and is likely to do so for
quite some time in the future; at present, it relies mainly on the manipulation of culture medium,

3. especially growth regulators, and, to a lesser extent, other factors, including environmental
conditions.
Explant:-
The plant tissue or organ excised and used for in vitro culture is known as explant. Virtually,
any plant part may be used as an explant, the choice depending mainly on the objective of the
culture and the regeneration potential of different organs of a plant species. For example, for the
recovery of an interspecific hybrid, young hybrid embryos have to be used as explants. The
appropriate explant is removed from the mother plant, and freed from microbes by surface
sterilization prior to its culture in vitro.

 Node : node culture
 Hairy root : hairy root culture
 Leaf : leaf culture
 Anther : anther culture
 Pollen : pollen culture
 Embryo : Embryo culture
 Cell : Cell culture
 Protoplast : Protoplast culture
 Callus : Callus culture

4. Surface Sterilization:-
The explants must be surface sterilized to
eliminate bacterial and fungal spores present
on their surface. This is commonly achieved
by treating them with 1-2 per cent solution
of sodium or calcium hypochlorite or with
-0.1% solution of mercuric chloride. The
explant is then rinsed several times with
fertilized distilled water to remove the
disinfectant. This and the subsequent
handling of explants or cultured cells and
organs has to be done under aseptic
conditions,

i.e., in an environment free from bacterial
and fungal spores. Laminar flow clean air
work stations provide aseptic conditions
readily, reliably and cheaply.



Sterilization:-
Plant tissue culture media are very rich, and they readily support microorganism growth. Growth
of a microorganism in a culture tube of plant cells/tissues/organs is called contamination.
Contamination must be avoided otherwise the cultures will be overrun by the contaminants.
Therefore, microbes present in culture media, culture vessels, instruments, etc. are inactivated by
a suitable treatment; this is called sterilization. Sterilization technique depends mainly on the
material to be sterilized.
1. Flame sterilization: - Instruments
like forceps; scalpels, needles, etc.
are dipped in 95% alcohol and
flamed just before use. Care should
be taken to cool them before use.

2. Dry heat: - Mouths of test tubes, culture flasks, etc. are ordinarily heated on a burner or
spirit lamp.

3. Ethanol (70%):- Laminar air flow
cabinet bench surface, outer surface
of culture vessels, hands of the
worker, etc. are wiped with 70%
ethanol and the alcohol is allowed to
evaporate.

5. 4. Autoclaving: - Culture media,
empty culture vessels, etc. are
autoclaved at 121°C and 15 p.s.i.
(pounds per square inch) for, usually,
15-20 minutes. The effectiveness of
sterilization depends mainly on the
temperature and time.


5. Air filter:-. The air blowing through laminar hoods is sterilized by HEPA (high efficiency
particulate air) filters.
6. Filter Sterilizatiom:- Thermo labile constituents like ABA, GA3, enzymes, etc. are filtered
through membrane filters of 0.45 µm pore size. The filter assembly itself must be sterilized,
usually by autoclaving, before use.

Nutrient Medium:-

The medium used for culture of plant cells and organs is known as nutrient medium, culture
medium, simply, medium. The medium contains inorganic salts to provide the 12 elements,
excluding C, H, O, necessary for plant growth; these elements are N, P, K, S, Ca, Mg (the six
macronutrients), Fe, , Cu, Zn, B, and Mo (the six micronutrients). In addition, certain vitamins, a
carbon source (generally sucrose) and, where needed, growth regulators like auxins and/or
cytokinins are also provided. 2,4-D (0.5-2.0 mg 1) is the most commonly used auxin; NAA and
IAA are also used. Similarly, kinetin and benzyl aminopurine (BA) are the most commonly used
cytokinins; some other cytokinins like zeatin, 2-ip, etc. are occasionally used. There are many
standard nutrient media available, but none of them is suitable for every purpose. Often, the
experimenter has to make some modifications to develop a medium suitable for his own needs.
Sometimes, complex organic supplements like coconut water, casein hydrolysate and yeast
extract are also used. The pH of the medium is generally adjusted to about 5.5 using IN KOH or
HCl as per need.

The medium may be solidified by using agar (⁓ 6g/1) or it may be used
as liquid. The medium is distributed into appropriate culture vessels, e.g., test tubes, culture
flasks, petriplates, etc., autoclaved at 15 p.s.i. for 15-20 min to free it from microbes, allowed to
cool, and stored for 2-3 days before use to be sure of proper sterilization. Sterilized explants are
then placed on to/into the nutrient medium; this operation is done under aseptic conditions.
When liquid medium is used, the culture flasks have to be constantly agitated or shaken on a
gyratory shaker (at about 100-200 rpm) to facilitate aeration. The cells on an agar medium
develop into an unorganised mass known as callus; consequently, they are called callus

6. cultures. In the liquid medium, on the other hand, a suspension of free cells and small cell
masses is obtained, and such cultures are known as suspension culture.



Environmental Conditions:-

Plant tissue cultures are maintained under a controlled environment, particularly in terms of
temperature and light. The temperature may vary from 18-25°C depending upon species and the
purpose of culture. Light is not essential for cell and tissue cultures, but it is often beneficial for
plantlet regeneration and for embryo and meristem cultures. The culture room or the incubator
contamination.

Culture room Incubator
.

Subculturing:-

After a period of time, it becomes necessary
to transfer organs and tissues to fresh media.
This is particularly true for tissue and cell
cultures, where a portion of tissue is used to
inoculate new culture tubes or flasks; this is
known as subculturing.

In general, callus cultures are subcultured
every 4-6 weeks, while suspension cultures
need to be subcultured every 3-14 days.
Theoretically, plant cell and tissue cultures
may be maintained indefinitely by serial
subculturing.

7. Plantlet Regeneration and Transfer to Soil

The application of plant tissue culture technology to crop improvement depends on regeneration
of complete plantlets and their successful transfer to soil. Production of various organs, e.g., root,
shoot, etc., in cultured tissues is known as organ regeneration or organogenesis. It is possible
to regenerate complete plantlets from tissue cultures of a large number of species, including rice,
maize, barley, oats, sugarcane, potato, pea, soybean, chickpea, alfalfa, etc. But plantlet
regeneration in many species, including some of the pulses, has not been obtained thus far.
Regeneration capacity appears to be genetically controlled, and it may be possible to improve a
suitable breeding programmed. For example, in alfalfa (M. sativa), two cycles of recurrent
selection improved the regeneration capacity from 12 to 67 per cent.
In many cases, transfer of whole plants from test tubes to soil is relatively easy. The
plants may be pretreated prior to their transfer to soil with different media designed to make
them hardy or even with some microbes (mainly bacteria). In a simple laboratory procedure,
plants may be transferred into small pots and covered with a suitable material/vessel, e.g.,
inverted beakers to prevent excess transpiration.


After 3-4 days, the covers are removed for increasing periods of time till they are finally
removed, but the pots are still kept in diffuse light for the next 5-10 days. Hardening on a large
scale is done in mist chambers: the plantlets are initially kept in low intensity diffuse light and
high (95%) relative humidity for few days. The light intensity s then gradually increased, while
humidity is gradually decreased over a period of time. The plants may then be transferred into a
greenhouse, and after about 1-2 weeks y may be planted in soil and kept in sunlight. Seedling
survival may vary from 50 to 100%, depending mainly on plant species.


A Classification of Tissue Culture Techniques

The tissue culture techniques are grouped into the following four categories on the basis of plant
part used as explant and the type of development in vitro: (1) embryo culture, (2) meristem
culture, (3) anther or pollen culture, and (4) cell culture. This classification is for convenience
and is Useful in discussion on the subject.

8. EMBRYO CULTURE

In embryo culture, young embryos are removed from developing seeds and cultured on a
suitable nutrient medium to obtain seedlings. If necessary, excised embryos may be placed
onto/into cultured endosperms, usually, of the same species. Young embryos require a high
osmotic concentration in the medium ns, embryos grow older, and the osmotic concentration
needs to be reduced. Cultured embryos generally de le complete development, and develop into
seedlings prematurely. Sometimes, embryos from mature se may also be used for embryo
culture, e.g., in Iris, orchids, etc. Generally, it is difficult to culture embs before a certain stage of
development, for example, before the globular stage in the case of barley, But in some species,
even few-celled embryos have been cultured successfully. Elaborate media for embryo culture
have been devised, but the tissue culture media may also be used for this modifications. Suitable
techniques are available for embryo culture in many crop species, and may be easily developed
for other species, if needed.



Applications of embryo culture:-

 Production of rare hybrids from intergeneric and interspecific crosses
 Development of disease resistant plants
 Production of haploids
 Overcoming seed dormancy
 Shortening of breeding cycle
 Propagation of rare plants
 Prevention of embryo abortion


MERISTEM CULTURE


The cultivation of axillary or apical shoot meristems is known as meristem culture. It involves
the development of an already existing shoot meristem and regeneration of adventitious roots.
However, often there is regeneration of adventitious shoots as well. Meristem cultures have been
extensively used for quick vegetative propagation of a large number of plant species. Ordinarily,
it is not necessary to excise or isolate the apical meristem, and usually 5-10mm shoot apices

9. containing the shoot apical meristem are used as explants. The shoot-tip may be cut into fine
pieces to obtain more than one plantlet from each shoot-tip. Nodal explants may also be used for
meristem culture. Generally, the standard tissue culture media are suitable for this purpose with
some modifications, where necessary.
Rapid multiplication may be achieved in one of the following two ways. The axillary buds
present in the explant and in the newly developing branches are stimulated to develop into
branches; this is achieved by a relatively higher concentration of a suitable cytokinin added into
the medium (enhanced axillary branching). After an appropriate period, each axillary branch is
excised and transferred to a fresh medium suitable for enhanced axillary branching.






Alternatively, the branches may be transferred to a rooting medium and subsequently transferred
to soil. But in some species, enhanced axillary branching cannot be achieved, e.g., in blue berry,
Dalbergia sissoo (sisam), etc., and each axillary bud/shoot-tip develops into a single shoot. In
such species, nodal cuttings (each containing 1-2 nodes) are excised from the shoots and cultured
for further multiplication.
The rates of multiplication may range from merely 16-fold in 52 weeks (Aechmea fasciata) to 9
x 10 in 52 weeks (Chrysanthemum). It is advisable to use the explants, taken from field-grown
plants, for 4-6 cycles of multiplication only in order to minimize the risk of somaclonal
variations and their unintended multiplication. In some species, e.g., banana, variants may arise
at a relatively high frequency if the conditions of culture are not carefully controlled.


Applications of meristem culture:-

• Production of virus free germplasm.
• Virus free plants serve as excellent experimental materials for evaluating the detrimental
effects of infections by various viruses.
• Mass production of desirable genotypes.
• Meristem culture can also help to eliminate other pathogens, e.g. mycoplasma, bacteria,
fungi.

10. ANTHER OR POLLEN CULTURE


Haploid plants may be obtained from pollen grains by placing anthers or isolated pollen grains
on a suitable culture medium; this is known as anther or pollen culture. Anthers may be taken
from plants grown in the field or in pots, but ideally, these plants should be grown under
controlled temperature, light and humidity. Often, the capacity for haploid production declines
with the age of donor plants. Exposure of the excised flower buds to a low temperature for some
time, e.g, at 5°C for 72 hr for tobacco, prior to the removal of anthers for culture may markedly
enhance the recovery of haploid plants. In some species, however, a brief exposure of the anthers
to a high temperature has a promotory effect, e.g. at 32°C for 8 hr in Brassica napus.

The medium requirements may vary with the species, the genotype, the age of the donor
plants and anthers, and the conditions under which the donor plants are grown. For example,
pollen grains of Datura and tobacco produce embryos on an agar medium containing only 2-4%
sucrose, while an elaborate medium had to be formulated for cereals. Sucrose is essential for
anther cultures; the concentration may range from 3% for barley to 6% for wheat and potato. For
most plant species a complete tissue culture medium is required; appropriate concentrations of
auxius and cytokinins may often be required. Anther cultures are generally maintained in
alternating periods of light (12-18 hr; 5,000-10,000 lux m?) at 28°C and darkness (12-6 hr) at
22°C, but the optimum conditions vary with species. The walls of responsive anthers turn brown
and after 3-8 weeks they burst open due to the developing callus/embryos. After the seedlings
(from embryos) or shoots (from callus) become 3-5 cm long, they are transferred to a medium
conducive to good root development. Finally, the plantlets are transferred to soil in the same way
as other in vitro-regenerated plantlets.

The optimum stage of pollen varies with the species. For many species, including Datura,
tobacco etc. the optimum stage is just before or just after the first pollen mitosis, while the early
biucleate stage is the most suitable for Atropa belladona and Nicotiana sylvestris, and is
absolutely essential for Nicotigno knightiana. In cereals and most other plant species, the best
stage appears to be the carly or mid- uninucleate stage, i.e., before the first pollen mitosis. In
tobacco, beginning of starch accumulation in pollen grains marks the end of their embryogenic
potential. Many crop species like tobacco, barley, wheat, etc. exhibit pollen dimorphism, i.e.,
most of their pollen grains are bigger, stain deeply with acetocarmine and contain plenty of
starch, while a small proportion (⁓0.7%) of pollen grains is smaller and stains faintly with
acetocarmine. The smaller pollen grains are called s-grains, and they respond during anther
culture; the frequency of responding pollen grains can be enhanced over that of s-grains by
certain pretreatments, e.g., chilling.

The early divisions in the responding pollen grains may occur in one of the following five
ways. (i) The nucleate pollen grain may divide symmetrically to yield two equal daughter cells,
both of which undergo further division, e.g., in Datura innoxia (Pathway I). (ii) In some other
cases, e.g., in tobacco, barley, wheat, triticale, chillies, etc., the unicleate pollen divides
unequally (as it does in nature). The generative cell degenerates, and the callus/embryo
originates from the vegetative cell (Pathway II). (iii) But in a few species, e.g., in Hyosciamus

11. niger, the pollen embryos originate from generative cell alone; the vegetative cell either does not
divide or divides only to a limited extent forming a suspensor like structure (Pathway III). (iv)
In some species, e.g., in Datura innoxia, the uninucleate pollen grains divide unequally
producing generative and vegetative cells, both of which contribute to the developing embryo
(Pathway IV). (v) Finally, in B. napus, the first division is symmetrical, and the pollen embryos
develop exclusively from the vegetative cell (Pathway V).

The responsive pollen grains become multicellular and ultimately burst open to release
the cell mass. This cell mass may either assume the shape of a globular embryo and give rise to
an embryo or it may develop into a callus depending on the plant species. In some species, e.g.,
rice, wheat, rye, maize, etc., the pollen grains can be induced to produce embryos or calli by
simply altering the medium composition. Pollen embryos are normally produced in anther
cultures of B. campestris, B. napus, several Nicotiana spp. (including N. tabacum and N.
rustica), etc. In such cases, the plantlets obtained from germination of pollen embryos are
generally haploid, but some polyploids are also produced. In many species like rice, barley,
wheat, tomato, triticales, etc., pollen grains produce callus, from which plantlets may be
regenerated under suitable culture conditions. In these cases, the ploidy level of plants varies
considerably more than it does in cases where embryos are produced. In case of indica rice,
about 50% of the plants derived from anther culture are diploid; these plants are of pollen origin,
and are actually doubled haploids.





Applications of anther or pollen culture:-
 Production of haploid plant
 Production of useful gametoclonal varition.

TISSUE AND CELL CULTURES

Application of plant tissue culture in crop improvement depends solely on regeneration of whole
plants from them. Both callus and suspension cultures of many species regenerate whole plants.
In many cases there is production of somatic embryos, e.g., carrot, wheat, soybean, coffee, tea. In
many other species shoot buds differentiate from the cell masses followed by differentiation of
roots, e.g., tobacco, rice wheal barley, coffee, etc. Regenerated plants often show variation in
policy level due to cytogenetic instability of plant tissue cultures, which increases with the
duration of in vitro culture, while their regeneration capacity decreases. The tissue culture
technique has several applications in crop improvement some of which are being exploited at
present.

12. 1. Shoot Regeneration
Shoot buds differentiate from cell cultures of many crop species, e.g., tobacco, alfalfa,
etc. Shoot buds are unipolar, i.e., have only plumule, and have vaseular connections with
the callus tissue. In general, a high cytokinin to auxin ratio supports shoot regeneration,
while a high auxin to cytokinin ratio promotes root regeneration. But in some species,
e.g., alfalfa, a two-step process of shoot regeneration may occur: first, callus is produced
on a medium having a high 2, 4-D to kinetin ratio, and second, shoot regeneration occurs
when the callus is transferred to a growth regulator-free medium. Gibberellins generally
suppress shoot regeneration.

Under conditions supporting shoot regeneration, localized groups of meristematic
cells, called meristemoids, are formed. Subsequently, shoot buds differentiate from,
usually, the outer cell layers of these meristemoids. There is some evidence that starch
accumulation in localized areas precedes shoot differentiation from them. Shoot buds
ultimately clongate to form shoots, which can be detached and rooted to give rise to
complete plants.

2. Somatic Embryogenesis
Somatic embryos generally originate from single peripheral or deep-seated cells of callus.
These cells divide to form a group of cells, which usually becomes cutinized on its
periphery. This group of cells divides and progresses through globular, heart-shaped,
torpedo-shaped and cotyledonary stages similar to zygotic embryos. Somatic
embryogenesis has four well recognised phases: (1) induction, (2). development, (3)
maturation, and (4) germination. In the induction phase, cells attain the capacity for
embryogenesis, and they may progress up to the globular stage. Generally, induction is
achieved by exposure to a high concentration of an auxin like 2,4-D, especially if the
cells in the explant are differentiated. But in case of less differentiated cells of young
zygotic embryos, a cytokinin may be enough for induction.

Development of somatic embryos beyond the globular stage constitutes the
development phase. This may occur on the induction medium itself, more generally on a
growth regulator-free medium or, in many cases, on a specially devised medium. In
maturation phase, somatic embryos do not grow in size, but they become tolerant to
desiccation. Maturation is promoted by ABA or a high sucrose concentration. Finally, the
somatic embryos germinate to produce seedlings (germination phase); this is usually
promoted by GA. Often somatic embryos develop from cells of other somatic embryos
and/or seedlings derived from them; such embryos are called secondary somatic
embryos. Repeated cycles of secondary somatic embryo production is called recurrent
somatic embryogenesis; this is useful in large scale production of somatic embryos.

13. Application of somatic embryogenesis:-

 Somatic embryogenesis may replace micropropagation for the rapid propagation of
econonically important plants
 Somatic embryos can meet specific breeding objectives by rapidly multiplying
germplasm that is initially present as embryonic materiai e.g maternal embryos ,haploid
embryos and interspecific hybrid embryos that normally abort due to non availability of
endosperm tissue
 Raising somaclonal variations from tree Species.
 Production of synthetic seeds
 Source of regenerable protoplast system
 Embryogenic callus, suspension culture and somes have been employed as sources of
protoplast isolation for a range of species.
 The nucellus usually degenerated during the development of seeds but in citrus species
embryogenic callus derived from nucellus remains totipotent for many years, can be used
as source material for regeneration of plants.



SOMATIC HYBRIDIZATION

Production of hybrid plants through the fusion of protoplasts of two different plant
species/varieties is called somatic hybridization, and such hybrids are known as somatic
hybrids. The technique of somatic hybridization involves the following four steps: (i) isolation
of protoplasts, (ii) fusion of protoplasts, (i) selection of hybrid cells, and (iv) proliferation of the
hybrid cells and regeneration of hybrid plants from them.

1. Protoplast Isolation
Isolation of protoplasts is readily achieved by treating the cells/tissues with a suitable
mixture of cell wall degrading enzymes. Usually, a mixture of pectinase or macerozyme

14. (0.1-1.0%) and cellulase (1-2%) is appropriate for most plant materials. Osmotic
concentration of the enzyme mixture and of subsequent media is elevated (usually, by
adding 500-800 mmol L1 sorbitol or mannitol plus 50-100 mmol L CaCl2) to stabilize
the protoplasts and to prevent them from bursting. The cells and tissues are incubated in
the enzyme mixture for few to several hours; naked protoplasts devoid of cell wall are
gradually released in the enzyme mixture. Protoplasts have been isolated from virtually
all plant parts, but leaf mesophyll is the most preferred tissue at least in case of dicots. In
general, fully expanded leaves are surface sterilized, their lower epidermis is peeled off
with a pair of forceps and the peeled areas are cut out with a scalpel and suspended in the
enzyme mixture. After the period of incubation, protoplasts are washed with a suitable
washing medium to remove the enzymes and the debris. The protoplasts may be cultured
on a suitable medium in a variety of ways; they readily regenerate cell wall, and undergo
mitosis to form macroscopic colonies, which can be induced to regenerate whole plants.

2. Protoplast Fusion
A number of strategies have been used to induce protoplast fusion; of these, the following
three have been relatively more successful. Protoplasts of the desired strains/species are
mixed in almost equal proportion, generally, while still suspended in the enzyme mixture.
The protoplast mixture is then subjected to a high pH (10.5) and high Ca2+ concentration
(50 mmol L) at 37°C for about 30 min (high pH-high Ca* treatment). This technique is
quite suitable for some species, while for some others it may be toxic. Polyethylene
glycol (PEG)-induced protoplast fusion is the most commonly used as it induces
reproducible high frequency fusion accompanied with low toxicity to most cell types.
The protoplast mixture is treated with 28-50% PEG (MW 1,500-6,000) for 15-30 min,
followed by gradual washing of the protoplasts to remove PEG; protoplast fusion occurs
during washing. The washing medium may be alkaline (pH 9-10) and may contain a high
Ca2+ concentration (50 mmol L); this approach is a combination of PEG and high pH-
high Ca+ treatments, and is usually more effective than either treatment alone.

The above fusion techniques are nonselective in that they induce fusion between
any two or more protoplasts, A more selective and less drastic approach is the
electrofusion technique, which utilizes low voltage current pulses to align the protoplasts,
and pairs of protoplasts are then selected with a micromanipulator and placed in separate
microclectrofusion chambers. The protoplasts are three of by a short pulse of high
voltage, which induces disturbances in plasma lemma that cause the protoplast to fuse.
The entire operation is carried out using a specially designed equipment under a
microscope,

3. Selection of Hybrid Cells
The protoplast suspension recovered after a fusion inducing treatment consists of the
following cell (1) unfused protoplasts of the two species/strains, (11) products of fusion
between two or more proton of the same species (homokaryons), and (iii) ‘hybrid'
protoplasts produced by fusion between on more protoplasts of each of the two species
(heterokaryons). In somatic hybridization, only the heterokaryons that result from fusion
between one protoplast of each of the two species/strains are of interest. However, they
form only a small proportion (usually, 0.5-10%) of the protoplast population Therefore, it

15. is critical that an effective strategy is used for their selection. Some visual markers
pigmentation of the parental protoplasts, may be used for the identification of hybrid cells
under microscope; hybrid cells are then mechanically isolated and cultured. Where such
features are not available the protoplasts of the two parental species may be separately
labelled with different fluorescent agents but this approach is time consuming, and
requires considerable skill and effort.

Some strategies make use of some properties (usually, deficiencies) of the
parental species, which are not expressed in the hybrid cells due to complementation
between their genetic systems. These properties may be naturally present in the parental
species or may be artificially induced through mutagenesis. These strategies are simple,
highly effective and the least demanding, but their application is drastically limited by the
nonavailability of suitable properties (both natural and induced) in most of the parental
species of interest. Genetic engineering has been used to resolve the above difficulty. In
this approach, separate selectable marker genes, e.g., for antibiotic resistance, are
introduced in the two parents to be fused, and the hybrid cells are seleced on a medium
supplemented with both the antibiotics (selection agents) concerned.
A more general and widely applicable strategy is to culture the entire
protoplast population without applying any selection for hybrid cells. All the types of
protoplasts form calli, and the hybrid calli are later identified on the basis of callus
morphology, chromosome constitution, protein and enzyme banding patterns, DNA
markers, etc. In some cases, the identification may be delayed till the plants are
regenerated. Many workers tend to favour this approach since it does not depend on the
presence of appropriate specific features in the parental species.

4. Regeneration of Hybrid Plants
Once hybrid calli are obtained, plantlets are regenerated from them since this is a
prerequisite for their use in crop improvement. Further, the hybrid plants must be at least
partially fertile, in addition to having some useful property, for use in breeding schemes,
Somatic hybrids have been regenerated from a number of species combinations but it has
not been possible to recover hybrid plantlets and/or calli from several other combinations.
Some somatic hybrids retain the full or nearby full somatic complements of the two
parental species; these are called symmetric hybrids. Such hybrids provide unique
opportunities for synthesizing novel species, which may be of theoretical and/or practical
interest. Frequently, somatic hybrids (symmetric) between distantly related sexually
incompatible species are sterile. In such cases, somatic hybridization 1 e other S: such 3n
plants may be partially fertile. These 3n hybrids can be used for gene! Chromosome
introgression from the species contributing the haploid protoplast.

Many somatic hybrids exhibit the full somatic complement of one species, while
all or nearly all the chromosomes of the other species are lost during the preceding
mitotic divisions; such hybrids are referred to as asymmetric hybrids. Such hybrids are
likely to show a limited introgression of chromosome segments from the eliminated
genome(s) due to drastically enhanced chromosomal aberrations and/or mitotic crossing
over in vitro. Asymmetric hybrids can be obtained even from those combinations, which
normally produce symmetric hybrids, by irradiating the protoplasts of one of the parental

16. species with a suitable dose of X-rays or gamma-rays to induce extensive chromosome
breakage.

Sexual hybrid cells, i.e., zygotes, contain chromosome complements (genomes)
from both the parental species, but the cytoplasmic genes (plasmon) from the female
parent only. In contrast, somatic hybrid cells contain both nuclear as well cytoplasmic
complements (present in chloroplasts and mitochondria) from both the parental species.
But somatic hybrid plants, in general, contain chloroplasts of only one of the two parental
species, and only a small proportion of them retains the chloroplasts of both the species.
The same applies to the mitochondria of the two fusion parents. A somatic hybrid plant
may have both chloroplants and mitochondria from the same or different fusion parents.
There considerable evidence that the genomes of both chloroplasts and mitochondria,
particularly the latter, undergo recombination in the hybrid cells, thereby producing
recombinant organelles in the progeny.


5. Cybrids
Cybrids or cytoplasmic hybrids are cells containing nucleus of one species but
cytoplasm from both the parental species. They are produced in variable frequencies in
normal protoplast fusion experiments due to (i) fusion of a normal protoplast of one
species with an enucleate protoplast or a protoplast having. inactivated nucleus of the
other species, (ii) elimination of the nucleus of one species from a normal heterokaryon,
or (ii) gradual elimination of the chromosomes of one species from a hybrid cell during
the subsequent mitotic divisions. Cybrids may be produced in relatively high frequency
by () irradiating the protoplasts of one species prior to fusion, or (ii) by preparing
enucleate protoplasts (cytoplasts) of one species and fusing them with normal protoplasts
of the other species. However, plants regenerated from cybrid cells generally have
chloroplasts/mitochondria of only one parent.

6. Applications of Somatic Hybridization
Somatic hybridization once promised much more than it has been able to deliver; the
various realized and potential applications are briefly summarised below.

1. Symmetric Hybrids. Symmetric hybrids can be produced between sexually
incompatible species, and between nonflowering or sterile strains of the same species.
Some of our allopolyploid crop like B. napus have narrow genetic base; this can be
rectified by producing symmetric hybrids from their parental species. Symmetric
hybrids have been crossed with crop species for transferring useful genes into the
latter, and even for cytoplasm transfer by repeated backerossing. Some symmetric
hybrids, either themselves or the lines derived from them may be useful novel
material for basic studies. Possibly, some symmetric hybrids possess such desirable
features that may be useful in breeding programmes.

2. Asymmetric Hybrids. These can be useful for gene and cytoplasm transfers. The
feasibility of these transfers has been demonstrated in several cases. Transfers using

17. asymmetric hybrids would be easier than that using symmetric hybrids since the latter
would require more number of backcrosses.

3. Cybrids. The objective of cybrid production is to combine the cytoplasmic genes of
one species with the nuclear and cytoplasmic genes of another species. But the
mitotic segregation of 'savageness leads to the recovery of plants having plasmagenes
of one or the other species only; only a small proportion of the plants remain "cybrid',
which would further segregate into the two parental types. This provides the
following unique opportunities: (i) transfer of plasmagenes of one species into the
nuclear background of another species in a single generation, and even in (ii) sexually
incompatible combinations, (iii) recovery of recombinants between the parental
mitochondrial or chloroplast genomes, and (iv) production of a variety of
combinations of the parental and recombinant chloroplasts with the parental or
recombinant mitochondria. When cybrids are produced by irradiating the protoplasts
of one species prior to fusion, they provide the additional opportunity for the (v)
recovery of chromosome 682 segment introgressions from the lost genome, in
combination with variations in the Plasmon.

Cybrids and asymmetric hybrids allow the mitochondria of one species to be
combined with a chloroplasts of the other species involved in the fusion. This is
achieved because the mitochondria e chloroplasts of the two fusion parents present in
a symmetric/an asymmetric hybrid/cybrid cell during the subsequent mitotic
divisions. As a result, most of the hybrid/cybrid plants contain mitochondria of only
one of the two fusion parents. The same is true for the chloroplasts as well. Some of
the cybrid plants contain mitochondria from one fusion parent, but the chloroplasts
from the other parent This feature has been used to correct the deficiency of chlorosis
at low temperatures due the Ogura'male sterility cytoplasm (see, Section 22.8.1.2 for
details). It must be emphasized that this can not be achieve by any comventional
breeding strategy.