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5. Embryo cultures

Embryo culture is the sterile isolation and growth of an immature or mature embryo in vitro

with the goal of obtaining a viable plant. Conventionally, the term embryo culture refers to

the sexually produced zygotic embryo culture. There are two types of embryo culture: mature

embryo culture and immature embryo culture (embryo rescue).

Mature embryo culture

Mature embryos are isolated from ripe seeds and cultured in vitro. Mature embryo cultures

are carried out when: the embryos remain dormant for long periods, embryos have low

survival in vivo, to avoid inhibition in the seed for germination or to convert sterile seeds to

viable seedlings. In some plants, seed dormancy may be due to chemical inhibitors or

mechanical resistance exerted by structures covering the embryo. Seed dormancy can be

successfully bypassed by culturing the embryos in vitro. Embryo culture is relatively easy as

they can be grown on a simple inorganic medium supplemented with energy source (usually

sucrose) to develop viable seedlings. This is possible since the mature embryos excised from

the developing seeds are autotrophic in nature.

Immature embryo culture

Embryo rescue involves the culture of immature embryos to rescue them from unripe or

hybrid seeds which fail to germinate. This approach is very useful to avoid embryo abortion

and produce a viable plant. Wild hybridization involving crossing of two different species of

plants from the same genus or different genera often results in failure. This is mainly because

the normal development of zygote and seed is hindered due to genetic barriers. Consequently,

hybrid endosperm fails to develop leading the abortion of hybrid embryo. The endosperm

may also produce toxins that ultimately kill the embryo. In the normal circumstances,

endosperm first develops and supports embryo development nutritionally. Thus, majority of

embryo abortions are due to failure in endosperm development. Embryo abortion can be

avoided by isolating and culturing the hybrid embryos prior to abortion. The most important

application of embryo rescue is the production of interspecific and inter-generic hybrids from

wild plant species.

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Culture Technique for Embryo Rescue:

The isolation of immature embryos often poses some difficulty. The aseptically isolated

embryos can be grown in a suitable medium under optimal conditions. In general, a complex

nutrient medium is required for culture methods involving embryo rescue. For adequate

nutritional support of immature embryos, embryo-endosperm transplant is used.

Embryo-endosperm transplant

Steps of the endosperm transplant technique used for culturing immature embryos: The

hybrid embryo from the ovule in which endosperm development has failed is taken out by

excision. Another normally developed ovule with endosperm enclosing an embryo is chosen.

This ovule is dissected and the normal embryo is pressed out. This leaves a normal

endosperm with an exit hole. Now, the hybrid embryo can be inserted into the normal

endosperm through exit hole. This results in embryo-endosperm transplant which can be

cultured in a suitable medium. By using embryo-endosperm transplant, many interspecific

and inter-generic plants have been raised e.g., hybrid plants of legumes.

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Nutritional needs for embryo cultures

1. If the embryo is heterotrophic: the embryo is mostly dependent on the endosperm and

maternal tissues for nutrient supply.

2. If the embryo is autotrophic: the embryo has the metabolic capability to synthesize

substances required for its growth which slowly makes it independent. The nutrient

supply is highly variable at this phase which mostly depends on the plant species. In

general, the composition of the medium for culturing immature embryos is more

complex than that required by mature embryos which can grow on a simple inorganic

medium. Further, the transfer of embryos from one medium to another is frequently

needed in order to achieve full development of embryos.

Applications of embryo culture: This way of culturing is needed for:

1. Prevention of embryo abortion

Incompatibility barriers in interspecific and inter-generic hybridization programs leading

to embryo abortion can be successfully overcome by embryo rescue. In fact, many distant

hybrids have been obtained through embryo rescue techniques. Some distant plant species

crossed and the resistance traits developed by employing embryo rescue.

2. Overcoming seed dormancy

Seed dormancy is caused by several factors—endogenous inhibitors, embryo immaturity,

specific light and temperature requirements, dry storage requirements etc. Further, in

some plants the natural period of seed dormancy itself is too long. Embryo culture is

successfully applied to overcome seed dormancy, and to produce viable seedlings in these

plant species.

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3. Shortening of breeding cycle

Some of the plants in their natural state have long breeding cycles. This is mostly due to

seed dormancy attributed to seed coat and/or endosperm. The embryos can be excised and

cultured in vitro to develop into plants within a short period. For instance, Hollies, a

Christmas decoration plant can be grown in 2-3 weeks through embryo cultures in

contrast to 3 years period required through seed germination.

4. Overcoming seed sterility

Certain plant species produce sterile seeds that do not germinate e.g. early ripening

varieties of cherry, apricot, and plum. Seed sterility is mostly associated with incomplete

embryo development which leads to the death of the germinating embryo. Using embryo

cultures, it is possible to raise seedlings from sterile seeds of early ripening fruits e.g.

apricot, plum.

5. Clonal Propagation

Clonal propagation refers to the process of asexual reproduction by multiplication of

genetically identical copies of individual plants. The term clone is used to represent a

plant population derived from a single individual by asexual reproduction. Embryos are

ideally suited for in vitro clonal propagation. This is due to the fact that embryos are

juvenile in nature with high regenerative potential.

Synthetic seed

Synthetic seeds have great potential for large scale production of plants at low cost as an

alternative to true seeds. It is often described as a novel analogue to true seed consisting of a

somatic embryo surrounded (or not according to type) by an artificial coat (like solidified

media) which is at most equivalent to an immature zygotic embryo. There are various

advantages of synthetic seeds such as;

1. Better clonal plants could be propagated similar to seeds;

2. Preservation of rare plant species extending biodiversity could be realized;

3. More synchronized harvesting of important agricultural crops would become a reality

4. Ease of handling,

5. Potential long-term storage and low cost of production

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Today synthetic seeds represent capsules with a gel envelope, which contain not only somatic

embryos but also axillary or apical buds. These plant materials are encapsulated in protecting

material (eg: hydrogel or alginate gel) and can be developed into a plant. The coating protects

the explants from mechanical damage during handling and allows germination and

conversion to occur without inducing undesirable variations. They behave like true seeds and

sprout into seedlings under suitable conditions.

Rules for the Production of Synthetic Seeds

The general procedure of synthetic seed production varies according to the type of artificial

seed produced, need of artificial seeds and the economic feasibility.

The development of the ideal viable, quiescent, low-cost artificial seed can be summarized as

follow:

The optimization of the clonal production system (optimizing protocols to synchronize

and maximize the development of normal mature embryos capable of conversion to

normal plants).

Post-treatment of mature embryos to induce quiescence,

Development of an encapsulation and coating system.

Optimization of the encapsulation system.

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Optimization of requirements for greenhouse and field growth (watering, fertilizer,

transplantation, etc.).

Identification and control of any pest and disease problems that may be unique to

artificial seeds and

Determination of the economic feasibility of using the artificial seed delivery system

for a specific crop compared with other propagation methods (cost–benefit analysis of

encapsulation versus other options).

The need for synthetic seed

Zygotic embryo seeds carry traits from both parents. Production of seeds carrying certain

traits requires homozygous parents for such traits which is not easy and time consuming.

After the discovery of somatic embryogenesis in 1950 it was possible to have an alternative

of conventional zygotic seeds. Somatic embryo arises from the somatic cells of a single

parent. They differ from zygotic embryos since somatic embryos are produced through in

vitro culture, without nutritive and protective seed coats and do not typically become

quiescent. Somatic embryos are structurally equivalent to zygotic embryos, but are true

clones, since they arise from the somatic cells of a single parent. The structural complexity of

artificial seeds depends on requirements of the specific crop application. Therefore, a

functional artificial seed may or may not require a synthetic seed coat, be hydrated or

dehydrated, quiescent or non-quiescent, depending on its usage. The field that seeks to use

somatic embryos as functional seed is termed ‘‘artificial or synthetic seed technology’’.

Types of synthetic seeds

There are various types of artificial seeds:

i. uncoated non quiescent somatic embryos, which could be used to produce those crops

micropropagated by tissue culture;

ii. uncoated, quiescent somatic embryos would be useful for germplasm storage.

iii. Non quiescent somatic embryos in a hydrated encapsulation constitute a type of artificial

seed that may be cost effective for certain field crops that pass through a greenhouse

transplant stage such as carrot, celery, seedless watermelon, and other vegetables and

iv. dehydrated, quiescent somatic embryos encapsulated in artificial coatings are the form of

artificial seed that most resembles conventional seed in storage and handling qualities. These

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consist of somatic embryos encased in artificial seed coat material, which then is dehydrated.

Under these conditions, the somatic embryos become quiescent and the coating hardens.

Theoretically, such artificial seeds are durable under common seed storage and handling

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conditions. Upon rehydration, the seed coat softens, allowing the somatic embryo to resume

growth, enlarging and emerging from the encapsulation.

Artificial seeds and their germination

6. Callus and cell suspension cultures

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Callus culture

Explant tissues generally show distinct planes of cell division, various specializations of cells,

and organization into specialized structures such as the vascular system. In contrary, callus

formation from explant tissue involves the development of progressively more random planes

of cell division with no polarity, less frequent specialization of cells and loss of organized

structures. Consequently, callus is defined as a mass of undifferentiated cells (meristematic

mode of action) arising from any kind of explant under in vitro culture conditions. It is also

naturally formed on in vivo plants in response to wounding.

How explant is changed to callus?

Mature plant cells generally do not divide in the intact plant, but they can be stimulated to

divide by wounding, by infection with certain bacteria, and by plant hormones, including

auxins and cytokinins. The course of callus development from an explant can be divided into

induction, and multiplication phases. In induction phase, the cells of explants prepare

themselves to divide, its metabolism is activated and the cell size remains constant. The

duration of this phase varies with the physiological state of the cells in the initial explant and

the culture conditions employed. In multiplication phase, regressive change involving the

return to the meristematic state starting from the peripheral wounded layers of the explants

and forming a certain growth pattern known as callus . This process was performed with the

help of the wound hormone, traumatic acid. The callus in this stage can be subcultured back

on the proliferation medium.

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The callus tissue from different plant species may be different in structure and growth habit.

Callus formation takes place under the influence of exogenously supplied growth regulators

present in the nutrient medium and the type of explant. The type of growth regulator

requirement and its concentration in the medium depends strongly on the genotype and

endogenous hormone content of an explant. Auxins like 2,4-D have strong effect on initiation

of cell division in tissue culture.

Callus growth passes several phases through sigmoid curve theory:

Lag phase: no or relatively very slow growth will be observed; only cell expansion

occurs as they established themselves on the new fresh medium.

Exponential phase: start by moderate growth to reach maximum increase in growth as the

cells actively growing synthesizing proteins, nucleic acid, phospholipids, as well as

multiplication of organelles and utilization of energy as ATP.

Linear phase: Cell division slows but rate of cell expansion increases.

Deceleration phase: Rate of cell division and elongation decrease compared to linear

phase (still rising)

Stationary phase: Number and size of cells remain constant.

Decline phase: due to the degradation of compounds over the synthesis processes and /or the

release of extracellular material which accumulated in the medium and cannot be recovered

by senescent cells.

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Callus tissue derived from the original explant can be established and maintained in an actively

growing state by the transfer of fragments to a fresh medium at regular 4-6 week intervals

(subculture). The nature of the callus tissue, its texture, compactness, friability and coloration

depends on the genotype, culture conditions and age of the explant. Growth of callus culture can be

monitored by some measurements, such as fresh and dry weights, growth indices, cell number or

mitotic indices.

Cell suspension culture (Cell culture)

Suspension culture is a type of culture in which single cells or small aggregates of cells

multiply while suspended in agitated liquid medium (using shaking incubator) to provide both

aeration and dispersion of cells. Like callus culture, the cells are also sub-cultured into new

medium.

Cell suspension cultures may be done in batch culture or continuous culture system. In the

later system, the culture is continuously supplied with nutrients by the inflow of fresh

medium with subsequent draining out of used medium but the culture volume is constant.

This culture method is mainly used for the synthesis of specific metabolite or for biomass

production.

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7. Protoplast (naked cell without cell wall) cultures

Cells of primary plant tissues possess cellulosic walls with a pectin-rich matrix, the middle

lamella, joining adjacent cells. The living cytoplasm of each cell, bounded by the plasma

membrane, constitutes the protoplast. Normally, intimate contact is maintained between the

plasma membrane and the wall, since this membrane is involved in wall synthesis. However,

in hypertonic solutions, the plasma membranes of cells contract from their walls. Subsequent

removal of the latter structures releases large populations of spherical, osmotically fragile

protoplasts (naked cells, no cell wall), where the plasma membrane is the only barrier

between the cytoplasm and its immediate external environment.

Protoplast isolation is now routine from a wide range of species; viable protoplasts are

potentially totipotent. Therefore, when given the correct chemical and physical stimuli, each

protoplast is capable, theoretically, of regenerating a new wall and undergoing repeated

mitotic division to produce daughter cells from which fertile plants may be regenerated via

the tissue culture process. A remarkable progress has been made in the number of species for

which protoplast-to-plant systems exist.

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Isolation of protoplast

Protoplasts can be isolated from a range of plant tissues: leaves, stems, roots, flowers, anthers

and even pollen. Protoplast isolation may be carried out by Mechanical disruption method or

enzymatic method. Out of these two methods, an enzymatic method is preferred as it provides

better protoplast yield with low tissue damage while mechanical method causes maximum

tissue chopping with lower protoplast yields.

In mechanical procedures, releasing protoplasts involved incubation of tissues in hypertonic

solution (eg: concentrated sucrose solution) to shrink protoplast away from cell wall, then

plasmolysed tissues are cut with a sharp-edged knife to remove only the cell walls.

In this process some of the plasmolyzed cells were cut only through the cell wall, releasing

intact protoplasts while some of the protoplasts may be damaged inside many cells. The

protoplasts are released by osmotic swelling when strips of tissues are placed in a hypotonic

solution.

In enzymatic method, the plasmolysed cells were separated by degrading enzyme or mixed

enzymes as cellulase, hemicellulase, pectinase, and protease.

In both cases, debris is filtered and/or centrifuged out of the suspension and the protoplasts

are then centrifuged to form a pellet. On resuspension the protoplasts (isotonic) can be

cultured on media which induce cell division and differentiation.

The isolation and culture media used vary with the species and with the tissue from which the

protoplasts were isolated. Some salts and nutrients (eg: sucrose or the sugar alcohol as

sorbitol/mannitol) are used as osmoticum to prevent plasma membrane from rupturing.

Pretreatment of donor tissues (plasmolysis or cold treatment) is used to reduce cytoplasmic

damage and spontaneous fusion of protoplasts from adjacent cells.

Factors affecting protoplast release

The physiological status of the source tissue influences the release of viable protoplasts.

Several factors influence protoplast release, including the extent of thickening of cell walls,

temperature, duration of enzyme incubation, pH optima of the enzyme solution, gentle

agitation, and nature of the osmoticum.

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Uses (applications)

Protoplasts are used in a number of ways for research and for plant improvement. They can

be treated in a variety of ways (electroporation, incubation with bacteria, heat shock, high pH

treatment) to induce them to take up DNA.

1. Somatic hybridization

Isolated protoplasts were observed to fuse spontaneously, which is now used to produce

hybrids from 2 sexually incompatible species. Protoplast fusion is achieved through high

Ca++

, high pH, Polyethylene glycol (PEG) or electric field.

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2. Transformation

The protoplast culture may develop genetically transformed plant where the transgenic is put

successfully within the protoplast. Then the protoplast regenerates a cell wall, undergo cell

division and forms callus. The callus can be subcultured. Embryogenesis begins from callus

when it is placed on nutrient medium lacking mannitol and auxin. The embryo develops into

the transgenic seedlings and finally into mature genetically modified plants.

Protoplast fusion

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8. Haploid Cultures

Sporophyte: is an independent plant with diploid chromosome number. In higher plants, the

sporophyte is dominant and performs vegetative and sexual reproduction. Gametophyte: is an

independent plant with haploid chromosome number. In higher plants, the gametophytes are

very much reduced and represents the gametes only which fuse to form sporophyte. Life

cycle alternates between sporophyte (2n) and gametophyte (n). Plants with gametophytic

chromosome number in their sporophyte are referred to as haploids. Haploids can result

through the culture of haploid explants like ovules or pollens. The process of haploid

regeneration though unpollinated (unfertilized) female ovules or ovaries are usually described

as gynogenesis, androgenesis is used when starting with intact anther, while microspore

(pollen) culture refers to isolate microspores (pollen) from anthers before culture. Haploid

plants develop from either cultures may proceed directly or indirectly through a callus phase.

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Note

The regenerated haploid plants are generally sterile, requiring chromosome doubling for use

in breeding programs. Chromosomes can be doubled (to produce homozygous individual at

all loci) either spontaneously or artificially, and haploid plantlets are usually treated with

colchicine as a means of inducing chromosome doubling (dihaploids/doubled haploid)

resulting a completely homozygous plant (2n).

Factors affecting haploid cultures

Several factors affecting haploid cultures from androgenesis, gynogenesis and microspore as

genotype, media, culture conditions,…..

Genotype

The choice of starting material for an anther or microspore culture project is of the utmost

importance. In particular, genotype plays a major role in determining the success or failure of

an experiment.

Haploid plant production via androgenesis has been very limited or nonexistent in many plant

species. Furthermore, within a species, differences exist in the ability to produce haploid

plants. Even within an amenable species, such as tobacco, some genotypes produce haploids

at a much higher rate than do others. Because of this genotypic effect, it is important to

include as much genetic diversity as possible when developing protocols for producing

haploid plants via anther or microspore culture.

Gynogenesis has not been investigated as thoroughly or with as many species as has

androgenesis; therefore, less information is available concerning the various factors that

contribute to the successful production of haploids from the female than the male

gametophyte. However, several studies have identified genotype as a critical factor in

determining the success of a gynogenesis experiment. Not only are there differences between

species, but genotypes within individual species have responded differently.

Condition of donor plants

The age and physiological condition of donor plants often affect the outcome of androgenesis

experiments. In most species, the best response usually comes from the first set of flowers

produced by a plant. As a general rule, anthers should be cultured from buds collected as

early as possible during the course of flowering. Various environmental factors that the donor

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plants are exposed to, may also affect haploid plant production. Light intensity, photoperiod,

and temperature have been investigated, and at least for some species, these are found to

influence the number of plants produced from anther cultures. Specific optimum growing

conditions differ from species to species; in general, the best results are obtained from

healthy, vigorously growing plants.

Media

Androgenesis can be induced in tobacco and a few other species on a simple medium such as

that developed by Nitsch and Nitsch (1969). For most other species, the commonly used

media for anther culture include MS (Murashige and Skoog, 1962), N6 (Chu, 1978), or

variations on these media. In some cases, complex organic compounds, such as potato

extract, coconut milk, and casein hydrolysate, have been added to the media. For many

species, 2–3% sucrose is added to the media, whereas other species, particularly the cereals,

have responded better to higher (about 15%) concentrations of sucrose. The higher levels of

sucrose may fulfill an osmotic rather than a nutritional requirement. Other sugars, such as

ribose, maltose, and glucose, have been found to be superior to sucrose for some species. For

a few species, such as tobacco, it is not necessary to add plant growth regulators (PGRs) to

the anther culture media. Most species, however, require a low concentration of some form of

auxin in the media. Cytokinin is sometimes used in combination with auxin, especially in

species in which a callus phase is intermediate in the production of haploid plants. Anther

culture media is often solidified using agar. Because agar may contain compounds inhibitory

to the androgenic process in some species, the use of alternative gelling agents has been

investigated. Gelrite, agarose, and starch have proven superior to agar for solidifying anther

culture media in various species. The use of liquid medium has been advocated by some

researchers as a way to avoid the potentially inhibitory substances in gelling agents. Anthers

may be placed on the surface of the medium, forming a so-called “float culture.”

Alternatively, microspores may be isolated and cultured directly in liquid medium.

Media has also been identified as an important factor in gynogenesis. The most commonly

used basal media for recovering gynogenic haploids are MS, B-5 (Gamborg et al., 1968),

Miller’s (Miller, 1963), or variations on these media. Sucrose levels have ranged from 2–

12%. While gynogenic haploids have developed in a few species without the use of growth

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regulators, most species have required auxins and/or cytokinins in the medium. For those

species that undergo indirect gynogenesis, both an induction and a regeneration medium may

be required. Most ovule and ovary culture experiments have been conducted using solid

medium.

Pretreatment and Culture conditions

For some species, a pretreatment following collection of buds, but before surface

disinfestation and excision of anthers, has been found to be beneficial. Yields of tobacco

haploids are often increased by storing excised buds at 7 to 8˚ C for 12 days prior to anther

excision and culture. For other species, temperatures from 4 to 10˚ C and durations from 3

days to 3 weeks have been utilized. For any one species, there may be more than one

optimum temperature and length of treatment combination. In general, lower temperatures

require shorter durations, whereas a longer pretreatment time is indicated for temperatures at

the upper end of the cold pretreatment range mentioned above. Cold pretreatment of flower

buds at 4˚ C for 4 to 5 days has been effective in increasing yields of haploid embryos or

callus through gynogenesis in a few species.

Various cultural conditions, such as temperature and light, may also affect androgenic

response. Anther cultures are usually incubated at 24 to 25˚ C. In some species, an initial

incubation at a higher or lower temperature has been beneficial. Haploid plant production was

increased in Brassica campestris L. by culturing the anthers at 35˚ C for 1 to 3 days prior to

culture at 25˚ C (Keller and Armstrong, 1979). In contrast, androgenesis was promoted in

Cyclamen persicum Mill. by incubating cultured anthers at 5˚ C for the first 2 days of culture.

Some species respond best when exposed to alternating periods of light and dark, whereas

continuous light or dark cultural conditions have proven beneficial in other species. Other

physical cultural factors, such as atmospheric conditions in the culture vessel, anther density,

and anther orientation, have been studied and found to affect androgenic response in some

species; however, species have varied greatly in their response to these physical factors.

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