Introduction to plant tissue culture and propagation Lecture Note

PLANT TISSUE CULTURE

Plant tissue culture, also referred to as cell, in vitro, axenic, or sterile culture, is an important tool in both basic and applied studies, as well as in commercial application. Plant tissue culture is the aseptic culture of cells, tissues, organs and their components under defined physical and chemical conditions in vitro. The theoretical basis for plant tissue culture was proposed by Gottlieb Haberlandt in 1902

Principle:

Totipotency, - The ability to regenerate the entire organism from a single somatic cell, i.e., trigger the use of the genetic information present to direct the entire regenerative and developmental programs needed to create the whole organism from a single cell, Cyto differentiation: dedifferentiation and re-differentiation are the principles. Dedifferentiation is the capacity of mature cells to return to meristematic condition and development of a new growing point

Competency describes the endogenous potential of a given cell or tissue to develop in a particular way. For example, as embryogenically competent cells are capable of developing into fully functional embryos. The opposite is non-competent or morpho-genetically incapable.

BRIEF HISTORY:

1838 - Schwann and Schleiden put forward the theory which states that cells are totipotent, and in principle, are capable of regenerating into a complete plant. Their theory was the foundation of plant cell and tissue culture

1902 - Haberlandt proposed concept of in vitro cell culture

1904 - Hannig cultured embryos from several cruciferous species

1922 - Kolte and Robbins successfully cultured root and stem tips respectively

1926 - Went discovered first plant growth hormone –Indole acetic acid

1934 - White introduced vitamin B as growth supplement in tissue culture media for tomato root tip

1939 - Gautheret, White and Nobecourt established endless proliferation of callus cultures

1941 - Overbeek was first to add coconut milk for cell division in Datura

1946 - Ball raised whole plants of Lupinus by shoot tip culture

1954 - Muir was first to break callus tissues into single cells

1955 - Skoog and Miller discovered kinetin as cell division hormone

1957 - Skoog and Miller gave concept of hormonal control (auxin: cytokinin) of organ formation

1959 - Reinert and Steward regenerated embryos from callus clumps and cell suspension of carrot (Daucus carota)

1960 - Cocking was first to isolate protoplast by enzymatic degradation of cell wall

1960 - Bergmann filtered cell suspension and isolated single cells by plating

1960 - Kanta and Maheshwari developed test tube fertilization technique

1962 - Murashige and Skoog developed MS medium with higher salt concentration

1964 - Guha and Maheshwari produced first haploid plants from pollen grains of Datura (Anther

culture)

1966 - Steward demonstrated totipotency by regenerating carrot plants from single cells of tomato

1970 - Power et al.successfully achieved protoplast fusion

1971 - Takebe et al.regenerated first plants from protoplasts

1972 - Carlson produced first inter specific hybrid of Nicotiana tabacum by protoplast fusion

1974 – Reinhard introduced biotransformation in plant tissue cultures- starting of genetic engineering

1977 - Chilton et al. successfully integrated Ti plasmid DNA from Agrobacterium tumefaciens in plants

1978- Melchers et al. carried out somatic hybridization of tomato and potato resulting in pomato

1981- Larkin and Scowcroft introduced the term somaclonal variation

1983 - Pelletier et al. conducted intergeneric cytoplasmic hybridization in Radish and Grape

1984 - Horsh et al. developed transgenic tobacco by transformation with Agrobacterium

1987 - Klien et al. developed biolistic gene transfer method for plant transformation

2005 - Rice genome sequenced under International Rice Genome Sequencing Project

ORGANIZATION OF LABORATORY

Any laboratory designed for plant tissue culture or biotechnology must focus on cleanliness & maintaining of aseptic condition. The essential 7 fundamental matter is the contamination free condition in all steps of the procedure. Any laboratory, in which tissue culture techniques are performed, regardless of the specific purpose, must contain a number of basic requirements.

These are:

a)      A general washing area

b)      A media preparation, sterilization & storage area

c)      Environmentally controlled incubators or culture rooms

d)      An observation/data collection area

e)      Acclimatization area

a.      Washing area:

The washing area should contain good quality basin, large sink & well drainage facilities. It should have access to dematerialized water & double distilled water. Space for drying ovens or racks, automated dishwashers, acid baths, pipette washers & driers & storage cabinets should also be available in the washing area.

General guidelines for washing area;

1.      Reusable glassware for tissue culture should be emptied immediately & need to be soaked in water. Media or agar must never be allowed to dry on the glassware

2.      All glassware containing corrosive chemicals or fixatives should be separated from the rest of the tissue culture glassware.

3.      All glasswares contaminated or coming into contact with microorganisms should be autoclaved before washing.

4.      The contents of any containers should be discarded immediately after completion of an experiment.

5.      Flaks or beakers used for agar based media should be rinsed immediately after dispensing the media into culture vassels so as to prevent drying of the residual agar in the beaker prior to washing.

b.      Media Preparation Area:

This area comprises the central section of the laboratory, home to most of the activities. This area should have ample storage space for the chemicals, culture vessels & glassware required for media preparation & dispensing. The general laboratory section includes the area for media preparation for autoclaving the media & also for many of the activities that relate to the handling of tissue culture materials. Laboratory equipments required for media preparation room are as followsGas, water & electric supplies & compressed air & vacuum line.

1. Water heater
2. Different types of glasswares
3. Hot plate with magnetic stirrer
4. Coarse & sensitive balance
5. Spatula for use during weighing
6. Microwave oven for rapid heating media & agar mixture
7. pH meter
8. Distillation unit
9. De-ionizer
10. Metal racks for holding test tubes in the autoclave
11. Test tubes, flasks, plastic containers
12. Autoclave or cooker
13.  Storage tank for distilled and /or de-ionized water.

CHEMICALS FOR CULTURE MEDIA:

A. Inorganic elements:

a. Macro nutrient:

The need of macro nutrients is higher in tissue culture media. It provides both anion & cation for the plant cell. The name of each element with available form & important functions are given below:

Sl. No.	Name of the macro nutrient	Available form	Function
1.	Nitrogen (N)	KNO3 ,NH4 NO3	Both structural & functional role in protein synthesis
2.	Phosphorus (P)	KH2PO4	Activation in nucleotide synthesis
3.	Potassium (K)	KNO3	Essential for activation of many enzymes, maintenance of ionic balance of the cell.
4.	Calcium (Ca)	CaCl2.2H2O	Acts as a cofactor & largely bound to the cell wall & cell membrane, Essential for cation- anion balance by counteracting organic inorganic anions.
5.	Magnessium (Mg)	MgSO4.7H2O	Essential for photosynthesis & many other enzymatic reactions.
6.	Sulphur (S)	MgSO4.7H2O,K2SO4	Functional role in protein synthesis.

b. Micro nutrients

Micro nutrient is essential for plant cell tissue growth. The name of elements, available salt combination & function are given below:

Sl. No.	Name	Available form	Function
1.	Zinc (Zn)	ZnSO4.7H2O	Act as a component of a number of enzymes, plays active role in protein synthesis, specially in the synthesis of tryptophan
2.	Manganese (Mn)	MnSO4.4H2O	Help in photosynthesis
3.	Copper (Cu)	CuSO4.5H2O	Plays an important role in electron transport chain at the time of photosynthesis
4.	Molybdenum (Mo)		It participates in the conversion of nitrate to ammonium
5.	Boron (B)	H3BO3	It is required for the synthesis of cell wall & cell membrane
6.	Iron (Fe)	FeSO4.5H2O	Formation of protein, important for biosynthesis of chlorophyll
7.	Cobalt (Co)	CoCl2. 6H2O	Helpful for nitrogen fixation
8.	Chlorine (Cl)	CaCl2. 2H2O	To control the osmoregulation of cell development.

B. Organic Components

a. Vitamins:

Normally plants synthesis vitamins endogenously. When plant cells & tissues are grown on in vitro condition some essential vitamins are absolutely required.

Sl. No.	Name of the vitamin
1.	Thiamine (vitamin-B1)- Promotion of cell growth & development.
2.	Nicotinic acid B20 - Promotion of cell growth & development.
3.	Pyridoxin-HCl (B6) - Promotion of cell growth & development.
4.	Folic acid - Promotion of cell growth & development.
5.	Biotin -Promotion of cell growth & development.
6.	Riboflavin - Promotion of cell growth & development.
7.	Retinol (vitamin-A) -- Promotion of cell growth & development.

b.Myo-inositol:

It has several functions like sugar transport, carbohydrate metabolism, membrane structure & cell wall formation.

c. Sugar:

It can be supplied in the form of sucrose, glucose, and fructose. It is a source of carbon.

d. Amino acid:

Cultured tissues are normally capable of synthesis of amino acid. In spite of this, the addition of amino acids to the media is important for stimulating cell growth. Unlike inorganic nitrogen, amino acids are taken up more rapidly by plant cells. Glycine is the most common amino acid used in different tissue culture media. Some of the other amino acids like glutamine, asparagines, cystine etc. are also required for cell culture.

e. Plant growth regulators:

Plant growth regulators are the organic molecules which have different regulatory effects on growth & development in whole plants & plant tissues. It is the most critical component of any culture media accepted that without regulators, in vitro culture is often impossible. Plant growth regulators which are often used in plant tissue culture are the following.

i. Auxin: The major functions of auxin are cell division, cell elongation, organogenesis. It is frequently used as a rooting hormone. The most frequently employed auxins are IAA (Indole-3- acetic acid), IBA (Indole-3-butyric acid), NAA (Napthalene acetic acid), 2, 4-D (2 ,4- Dichlorophenoxy acetic acid). IAA is a naturally occurring auxin is added in concentration of 0.01-10 mg/l. The most effective auxin of callus proliferation for most cultures is 2, 4-D, but unfortunately it strongly suppresses organogenesis & should not be used in experiments involving root & shoot initiation.

ii. Cytokinin: Cytokinins are derivatives of adenine, which promote cell division, regulate growth and development in plant tissues. It is known as shooting hormone essential for induction of auxillary branching and adventitious shoot formation. The most widely used cytokinins are kinetin, zeatin, BAP (Benzyladenine), 2iP (2- isopentenyladenine).

iii. Other regulators: Other types of hormones which may be used in plant tissue culture include gibberellins (GA3), which promotes shoot elongation, and internodal elongation, ethylene and abscisic acid.

Aseptic Transfer Area/Inoculation room:

All the activities of sterile transfers are performed in this room. There must be a laminar air flow cabinet where all the precautions should be taken to prevent entry of any contaminant into the culture vial during the process of inoculation or subculture. Laminar air flow hoods are usually sterilized by switching on the hood and wipping the working sueface with 70% ethyl alcohol for 15 minutes before initiating any operation under the hood. Ultraviolet light (UV) is sometimes installed to disinfect the area; this light should only be used when people and plant materials are not in the room. This room is provided with:

1.      Laminar air flow cabinet: Inoculation & subculture by maintaining aseptic condition.

2.      Steribed sterilizer, Sprit lamp/Bunsen burner: Sterilization of the knives, scalpels, forceps etc.

3.      Stereo-microscope: observe for specific part.

4.      Ethyl alcohol: sterilization and flaming of small instruments.

5.      Tiles/glass plates use during sterile cutting.

6.      Hypochloride solution: sterilization of plant material.

c. Incubation Room/Culture Room: This is the room where light, temperature, humidity is maintained. All of these environmental considerations will vary depending on the size of the growth room.

Temperature: is an important consideration for the tissue culture and other factors like light, relative humidity, and shelving depend on it. Generally, temp. of the growth room remains in the range of 25± 2^oC. Temp. in the primary growth room can be maintained by air conditioner.

Lighting facility: Intensity of light in the room can easily be maintained by using fluorescent light with timer. However, most culture rooms are lighted at the 1000 lux (for 1000cft) with some going up 5000-10000 lux.

Light duration: 16-18 h/day.

Light quality: Spectral quality of light received by in vitro cultures is very important.

Relative humidity: Relative humidity (RH) is very difficult to control inside the room but humidifier can be used to control humidity. Humidity inside the room should be 70-75%

Shelves: Shelving with primary growth rooms can vary depending upon the situations & explants grown. Wood is recommended for the inexpensive easy to build shelves.

This room is provided with

1.      Temperature control (25± 2oC)

2.      Electricity supply essential for lighting, cooling and heating

3.      Shelves for culture racks

4.      Fluorescent tubes for lighting

5.      Timer for regulating day length

6.      Racks for culture vials

7.      Rotary shaker for suspension cultures

8.      Observations table.

d. Data collection Area: Culture room is prepared by glass wall. Qualitative data could be collected from outside of the culture room through the glass wall. The quantitative data could be collected from inside the culture room by following aseptic rules and regulation.

e. Acclimatization area:

Plants regenerated from in vitro tissue cultures are transplanted to vermiculite pots. The potted plants are ultimately transferred to greenhouses or growth cabinets and maintained for further observations under controlled conditions of light, temperature and humidity.

Major equipment and their function

Sl. No.	Name of the equipment	Function
1.	Autoclave machine, Pressure cooker	Sterilization of media, glassware &small instrument.
2.	Balance	Measurement of chemical from the range of µgm to Kg
3.	Hot plate magnetic stirrer	To mix the chemical & other ingredient of media

4.	pH meter	To determine the pH of various chemicals & media
5.	Refrigerator	To store all sorts of temperature-sensitive chemical & stock solution.
6.	Micro oven	To melt agar, agarose & other gelling agents.
7.	Hot air oven	For dry heat sterilization of cell & suspension culture
8.	Shaker	Use for gentle rotation of cell& suspension culture
9.	Filter sterilization unit with vacuum pump	Filtration of thermoliable compound like growth regulator, vitamin, amino acid etc.
10.	Microscope	To study the cell & tissue culture material at different stages of development
11.	Luxmeter	To measure the light intensity of the culture room
12.	Thermometer	To record the temperature reading of laboratory & culture room
13.	Centrifuge machine	To sediment cell & clean supernatant
14.	Laminar air flow cabinet	To avoid air remaining contaminant

 General rules to be followed in a tissue culture laboratory

1.      A laboratory should have an inventory & a complete up-to-date record of all the equipment along with their operating manual.

2.      A laboratory should have an inventory & a complete up-to-date record of all the chemicals including the name of manufacturer & grade.

3.      All chemicals should be assigned to specific areas preferably by their alphabetical order.

4.      Strong acid & bases should be stored separately.

5.      Special handling or storage procedure should be posted in the records so that retrieving of chemical is easy, because chemicals need storage at different temperatures ( for example room temperature 4^o, -20^o C)

6.      Chloroform, alcohol, phenol, which is volatile or toxic in nature, must be stored in a fume hood.

7.      Chemicals which are hygroscopic in nature must be stored in desiccators in order to avoid caking.

8.      Chemicals kept in refrigerator or freezers should be arranged either alphabetically or in small baskets.

Safety rules:

1. Eating, smoking and drinking is strictly prohibited in the tissue culture laboratory.

2. Toxic chemical must be handled with appropriate precautions and should be discarded into separate labeled containers. e.g. Organic compounds, halogens etc.

3. Broken glass and scalpel blades must be disposed into individual marked containers.

4. Pipettes, tips, Pasteur pipettes and other things used in the lab should be first collected in autoclavable bags and then it should be finally autoclaved and disposed in safe place.

5. Pipetting any solution should not be conducted without using any pipette.

6. First aid kits should be placed in every laboratory and every individual working in the laboratory should know its location and how to use its contents.

7. Fire extinguishers should be provided in each laboratory.

Steps involved in general techniques.

Regeneration of Plantlets:

1. Preparation of Suitable Nutrient Medium:

Suitable nutrient medium as per objective of culture is prepared and transferred into suitable

containers.

2. Selection of Explants:

Selection of explants such as shoot tip should be done.

3. Sterilization/ surface decontamination of Explants:

Surface sterilization of the explants by disinfectants and then washing the explants with sterile distilled water is essential.

4. Inoculation:

Inoculation (transfer) of the explants into the suitable nutrient medium (which is sterilized by filter-sterilized to avoid microbial contamination) in culture vessels under sterile conditions is done.

5. Incubation:

Growing the culture in the growth chamber or plant tissue culture room, having the appropriate physical condition (i.e., artificial light; 16 hours of photoperiod), temperature (-26°C) and relative humidity (50-60%) is required.

6. Regeneration:

Regeneration of plants from cultured plant tissues is carried out.

7. Hardening:

Hardening is gradual exposure of plantlets to an environmental condition.

8. Plantlet Transfer:

After hardening plantlets transferred to the green house or field conditions following acclimatization (hardening) of regenerated plants.

METODS OF STERILIZATION

The various methods of sterilization are:

1. Physical Method

(a) Thermal (Heat) methods

(b) Radiation method

(c) Filtration method

2. Chemical Method

3. Gaseous method

Nutrient medium

Culture media are largely responsible for the in vitro growth and morphogenesis of plant tissues. The success of the plant tissue culture depends on the choice of the nutrient medium. In fact, the cells of most plant cells can be grown in culture media. Basically, the plant tissue culture media should contain the same nutrients as required by the whole plant. It may be noted that plants in nature can synthesize their own food material. However, plants growing in vitro are mainly heterotrophic i.e. they cannot synthesize their own food.

Composition of Media:

The composition of the culture media is primarily dependent on two parameters:

1. The particular species of the plant.

2. The type of material used for culture i.e. cells, tissues, organs, protoplasts.

Thus, the composition of a medium is formulated considering the specific requirements of a given culture system. The media used may be solid (solid medium) or liquid (liquid medium) in nature. The selection of solid or liquid medium is dependent on the better response of a culture.

Major Types of Media:

The composition of the most commonly used tissue culture media is given in the following Table, and briefly described below.

White’s medium:

This is one of the earliest plant tissue culture media developed for root culture.

MS medium:

Murashige and Skoog (MS) originally formulated a medium to induce organogenesis, and regeneration of plants in cultured tissues. These days, MS medium is widely used for many types of culture systems.

B5 medium:

Developed by Gamborg, B5 medium was originally designed for cell suspension and callus cultures. At present with certain modifications, this medium is used for protoplast culture.

N6 medium:

Chu formulated this medium and it is used for cereal anther culture, besides other tissue cultures.

Nitsch’s medium:

This medium was developed by Nitsch and Nitsch and frequently used for anther cultures. Among the media referred above, MS medium is most frequently used in plant tissue culture work due to its success with several plant species and culture systems.

 Composition of common Tissue culture media

Synthetic and natural media:

When a medium is composed of chemically defined components, it is referred to as a synthetic medium. On the other hand, if a medium contains chemically undefined compounds (e.g., vegetable extract, fruit juice, plant extract), it is regarded as a natural medium. Synthetic media have almost replaced the natural media for tissue culture.

Expression of concentrations in media:

The concentrations of inorganic and organic constituents in culture media are usually expressed as mass values (mg/l or ppm or mg I-1). However, as per the recommendations of the International Association of Plant Physiology, the concentrations of macronutrients should be expressed as mmol/l–and micronutrients as µmol/l–.

Constituents of Media:

Many elements are needed for plant nutrition and their physiological functions. Thus, these elements have to be supplied in the culture medium to support adequate growth of cultures in vitro. A selected list of the elements and their functions in plants is given in the Table below.

Selected list of elements and their functions in plants

 

The culture media usually contain the following constituents:

1. Inorganic nutrients

2. Carbon and energy sources

3. Organic supplements

4. Growth regulators

5. Solidifying agents

Culture initiation and regeneration through different pathways.

Types of in vitro culture:

1. culture of intact plants e.g. seed culture in orchids

2. embryo culture e.g. immature embryo culture

3. organ culture e.g. meristem culture, shoot tip culture root culture anther culture

4. callus culture

5. single cell culture

6. Protoplast culture.

Culture initiation: selection of explants, sterilization, media optimization and establishment of the plants from in vivo to in vitro

Organogenesis

This is a major path of regeneration that involves the differentiation of culture cells or callus tissue into organs such as shoot and roots. Plant regeneration through the formation of shoots and roots is known as plant regeneration through organogenesis. Organogenesis can occur directly or indirectly from the explants depending on the hormonal combination of the medium and the physiological state of the explants. Miller and Skoog demonstrated that the initial formation of roots or shoots on the cultured callus or explant tissue depends on the relative concentration of auxins and cytokinins in the culture media. Medium supplemented with relatively high auxin concentration will promote root formation on the explants and high cytokinin concentration will promote shoot differentiation. In tissue culture practices there may be three types of medium in relative combinations of auxins and cytokinins, which promote either the shoot formation or root formation or both simultaneously. In the latter case, we can get the complete plantlets, having both shoot and roots, which can be directly transferred to the pots in the greenhouse. Whereas in other cases, after the formation of shoots, individual shoots are transferred to the rooting medium, which promote root formation. The rooted plantlets can be transferred to a greenhouse for acclimatization. Plant regeneration through organogenesis is commonly used for mass multiplication, for micropropagation, and for conservation of germplasm at either normal or subzero temperatures (cryopreservation)

Skoog and Miller (1957) were responsible to recognize the regulatory mechanism as a balance between auxin and cytokinin. As per their finding, a relatively high level of auxin to cytokinin favoured root formation and the reverse favoured shoot formation. Using this concept, it has now become possible to achieve organogenesis in a large number of plant species by culturing explants, calli and cell suspension in a defined medium. In organogenesis, the shoot or root may form first depending upon the nature of growth hormones in the basal medium. The genesis of shoot and root from the explants or calli is termed as caulogenesis (caulm = stem) and rhizogenesis (rhizo = root) respectively.

Organogenesis leading to complete plantlet regeneration is a multistage process consisting of at least three distinct stages.

1. shoot bud formation, 2. shoot development and multiplication 3. rooting of developed shoots.

Caulogenesis is a type of organogenesis by which only adventitious shoot bud initiation takes place in the callus tissue. When organogenesis leads to root development, then it is known as rhizogenesis. Abnormal structures developed during organogenesis are called organoids. The localized meristematic cells on a callus which give rise to shoots and/or roots are termed as meristemoids. Meristemoids are characterized as an aggregation of meristem-like cells. These can occur directly on an explant or indirectly via callus.

Thus, there are two kinds of organogenesis. A developmental sequence involving an intervening callus stage is termed 'indirect' organogenesis: Primary explant → callus → meristemoid → organ primordium. Direct organogenesis is accomplished without an intervening proliferate callus stage: Primary explant → meristemoid → organ primordium.  In vitro plant tissues may produce many types of primordia (adventitious buds and organs) including those that will eventually differentiate into embryos, flowers, leaves, shoots, and roots. These primordia originate de novo from a cellular dedifferentiation process, followed by initiation of a series of events that results in to an organ.

Embryogenesis/ Somatic embryogenesis

This is another major path of regeneration and development of plantlets for micropropagation or mass multiplication of specific plants. The cells, under a particular hormonal combination, change into the physiological state similar to zygotes (somatic zygotes) and follow an embryonic path of development to form somatic embryos. These somatic embryos are similar to normal embryos (seed embryos) developed from zygotes formed by sexual fertilization. The somatic embryos can develop into a complete plant. Since somatic embryos can germinate into a complete plant, these can be used for the production of artificial seeds. Somatic embryos developed by tissue or cell cultures can be entrapped in certain inert polymers such as calcium alginate and used as artificial seeds. Since the production of artificial seed is amenable to mechanization and for bioreactors, it can be produced in large numbers.

Embryogenesis Embryos have been classified into two categories: zygotic embryos and non-zygotic embryos. Zygotic embryogenesis Embryos developing from zygotes (resulting from regular fusion of egg) are called as zygotic embryos or often simply embryos. Non-zygotic embryogenesis Usually non- zygotic embryos are formed by cells other than the zygote. E.g. Parthenogenetic embryos - formed from unfertilized eggs or a fertilized egg without karyogamy. Androgenetic embryos – formed from microspores, micro-gametophytes or sperm.

Somatic embryos (also called as embryoids, accessory embryos, adventitious embryos and supernumerary embryos) – formed by somatic cells either in vivo or in vitro. A somatic embryo is an embryo derived from a somatic cell, other than zygote, usually on in vitro culture. The process of somatic embryo development is called as somatic embryogenesis.

Stages in development of somatic embryos

Somatic embryos generally originate from single cells which divide to form a group of meristematic cells. Usually, this multi-cellular group becomes isolated by breaking cytoplasmic connections with the other cells around it and subsequently by cutinization of the outer walls of this differentiating cell mass. The cells of meristematic mass continue to divide to give rise to globular (round ball shaped), heart-shaped, torpedo and cotyledonary stages.  Somatic embryo genesis begins with active division of cells which leads to increase in size but retains the spherical shape. At this stage the primary meristem (protoderm, ground meristem and procambium) becomes visible. Following this stage, the callus continues to divide and differentiate into a heart-shaped embryo, with initiation of cotyledon primordia.

As the cotyledon develops the embryo passes into the torpedo-shaped stage. The cells inside the cotyledonary ring divide to form shoot and root apical meristem and procambium differentiation takes place. In general, the essential features of somatic embryo development, especially after the globular stage, are comparable to those of zygotic embryo. The somatic embryogenesis can also be either direct or indirect depending up on the hormonal composition.

 

                         Diagrammatic representation on direct and indirect regeneration

Micropropagation

Micropropagation is the practice of rapidly multiplying stock plant material to produce a large number of progeny plants, using modern plant tissue culture methods

Technique of Micro propagation:

Micro propagation is a complicated process and mainly involves 3 stages (I, II and III). Some authors add two more stages (stage 0 and IV) for more comprehensive representation of micro- propagation. All these stages are represented in the following Figure, and briefly described hereunder.

 

Major stages involved in micropropagation

 Stage 0:

This is the initial step in micro- propagation, and involves the selection and growth of stock plants for about 3 months under controlled conditions.

 Stage I:

In this stage, the initiation and establishment of culture in a suitable medium is achieved. Selection of appropriate explants is important. The most commonly used explants are organs, shoot tips and axillary buds. The chosen explant is surface sterilized and washed before use.

 Stage II:

It is in this stage; the major activity of micro propagation occurs in a defined culture medium. Stage II mainly involves multiplication of shoots or rapid embryo formation from the explant.

 Stage III:

This stage involves the transfer of shoots to a medium for rapid development into shoots. Sometimes, the shoots are directly planted in soil to develop roots. In vitro rooting of shoots is preferred while simultaneously handling a large number of species.

 Stage IV:

This stage involves the establishment of plantlets in soil. This is done by transferring the plantlets of stage III from the laboratory to the environment of greenhouse. For some plant species, stage III is skipped, and un-rooted stage II shoots are planted in pots or in suitable compost mixture.

 The different stages described above for micro propagation are particularly useful for comparison between two or more plant systems, besides better understanding. It may however, be noted that not all plant species need to be propagated in vitro through all the five stages referred above.

 Micro propagation mostly involves in vitro clonal propagation by two approaches:

1.      Multiplication by axillary buds/apical shoots.

2.      Multiplication by adventitious shoots.

 Besides the above two approaches, the plant regeneration processes namely organogenesis and somatic embryogenesis may also be treated as micro propagation.

3.  Organogenesis: The formation of individual organs such as shoots, roots, directly from an explant (lacking preformed meristem) or from the callus and cell culture induced from the explant.

4.      Somatic embryogenesis: The regeneration of embryos from somatic cells, tissues or organs.

 1. Multiplication by Axillary Buds and Apical Shoots:

 Quiescent or actively dividing meristems are present at the axillary and apical shoots (shoot tips). The axillary buds located in the axils of leaves are capable of developing into shoots. In the in vivo state, however only a limited number of axillary meristems can form shoots. By means of induced in vitro multiplication in micro propagation, it is possible to develop plants from meristem and shoot tip cultures and from bud cultures.

 Meristem and Shoot Tip Cultures:

Apical meristem is a dome of tissue located at the extreme tip of a shoot. The apical meristem along with the young leaf primordia constitutes the shoot apex. For the development of disease-free plants, meristem tips should be cultured.

 Meristem or shoot tip is isolated from a stem by a V-shaped cut. The size (frequently 0.2 to 0.5 mm) of the tip is critical for culture. In general, the larger the explant (shoot tip), the better are the chances for culture survival. For good results of micro propagation, explants should be taken from the actively growing shoot tips, and the ideal timing is at the end of the plants dormancy period.

The most widely used media for meristem culture are MS medium and White’s medium. A diagrammatic representation of shoot tip (or meristem) culture in micro propagation is given in Fig  and briefly described hereunder.

 

Diagrammatic representation of shoot tip (or meristem) culture in micropropagation; I, II, III are stages.

In stage I, the culture of meristem is established. Addition of growth regulators namely cytokinins (kinetin, BA) and auxins (NAA or IBA) will support the growth and development.

In stage II, shoot development along with axillary shoot proliferation occurs. High levels of cytokinins are required for this purpose.

Stage III is associated with rooting of shoots and further growth of plantlet. The root formation is facilitated by low cytokinin and high auxin concentration. This is opposite to shoot formation since high level of cytokinins is required (in stage II). Consequently, stage II medium and stage III medium should be different in composition. The optimal temperature for culture is in the range of 20-28°C (for majority 24-26°C). Lower light intensity is more appropriate for good micro propagation.

 Bud Cultures:

The plant buds possess quiescent or active meristems depending on the physiological state of the plant. Two types of bud cultures are used— single node culture and axillary bud culture.

Single node culture:

This is a natural method for vegetative propagation of plants both in vivo and in vitro conditions. The bud found in the axil of leaf is comparable to the stem tip, for its ability in micro propagation. A bud along with a piece of stem is isolated and cultured to develop into a plantlet. Closed buds are used to reduce the chances of infections.

A diagrammatic representation of single node culture is below. In single node culture, no cytokinin is added.

 

Axillary bud culture:

In this method, a shoot tip along with axillary bud is isolated. The cultures are carried out with high cytokinin concentration. As a result of this, apical dominance stops and axillary buds develop. A schematic representation of axillary bud culture for a rosette plant and an elongate plant is given below

Fig. Schematic Representation of Axillary Bud Method of Vegetatively Propagating Plants. (a) Rosettle plants; (b) Elongate plants showing bud culture and single node culture. 

For a good axillary bud culture, the cytokinin/ auxin ratio is around 10: 1. This is however, variable and depends on the nature of the plant species and the developmental stage of the explant used. In general, juvenile explants require less cytokinin compared to adult explants. Sometimes, the presence of apical meristem may interfere with axillary shoot development. In such a case, it has to be removed.

 2. Multiplication by Adventitious Shoots:

 The stem and leaf structures that are naturally formed on plant tissues located in sites other than the normal leaf axil regions are regarded as adventitious shoots. There are many adventitious shoots which include stems, bulbs, tubers and rhizomes. The adventitious shoots are useful for in vivo and in vitro clonal propagation. The meristematic regions of adventitious shoots can be induced in a suitable medium to regenerate to plants.

 3. Organogenesis:

Organogenesis is the process of morphogenesis involving the formation of plant organs i.e. shoots, roots, flowers, buds from explant or cultured plant tissues. It is of two types — direct organogenesis and indirect organogenesis.

 Direct Organogenesis:

Tissues from leaves, stems, roots and inflorescences can be directly cultured to produce plant organs. In direct organogenesis, the tissue undergoes morphogenesis without going through a callus or suspension cell culture stage. The term direct adventitious organ formation is also used for direct organogenesis.

 Induction of adventitious shoot formation directly on roots, leaves and various other organs of intact plants is a widely used method for plant propagation. This approach is particularly useful for herbaceous species. For appropriate organogenesis in culture system, exogenous addition of growth regulators—auxin and cytokinin is required. The concentration of the growth promoting substance depends on the age and nature of the explant, besides the growth conditions.

Indirect Organogenesis:

When the organogenesis occurs through callus or suspension cell culture formation, it is regarded as indirect organogenesis. Callus growth can be established from many explants (leaves, roots, cotyledons, stems, flower petals etc.) for subsequent organogenesis.

Fig: Plant Regeneration Pathways

Micropropagation of plants by direct and indirect organogenesis

 The explants for good organogenesis should be mitotically active immature tissues. In general, the bigger the explant the better the chances for obtaining viable callus/cell suspension cultures. It is advantageous to select meristematic tissues (shoot tip, leaf, and petiole) for efficient indirect organogenesis. This is because their growth rate and survival rate are much better.

For indirect organogenesis, the cultures may be grown in liquid medium or solid medium. Many culture media (MS, B5 White’s etc.) can be used in organogenesis. The concentration of growth regulators in the medium is critical for organogenesis.

By varying the concentrations of auxins and cytokinins, in vitro organogenesis can be manipulated:

        i.            Low auxin and low cytokinin concentration will induce callus formation.

      ii.            Low auxin and high cytokinin concentration will promote shoot organogenesis from callus.

    iii.            High auxin and low cytokinin concentration will induce root formation.

4. Somatic Embryogenesis:

The process of regeneration of embryos from somatic cells, tissues or organs is regarded as somatic (or asexual) embryogenesis. Somatic embryogenesis may result in non-zygotic embryos or somatic embryos (directly formed from somatic organs), parthogenetic embryos (formed from unfertilized egg) and androgenic embryos (formed from male gametophyte).

 In a general usage, when the term somatic embryo is used it implies that it is formed from somatic tissues under in vitro conditions. Somatic embryos are structurally similar to zygotic (sexually formed) embryos, and they can be excised from the parent tissues and induced to germinate in tissue culture media.

Development of somatic embryos can be done in plant cultures using somatic cells, particularly epidermis, parenchymatous cells of petioles or secondary root phloem. Somatic embryos arise from single cells located within the clusters of meristematic cells in the callus or cell suspension. First a pro-embryo is formed which then develops into an embryo, and finally a plant.

 Direct Somatic Embryogenesis:

When the somatic embryos develop directly on the excised plant (explant) without undergoing callus formation, it is referred to as direct somatic embryogenesis (Fig 47.6A). This is possible due to the presence of pre-embryonic determined cells (PEDQ found in certain tissues of plants. The characteristic features of direct somatic embryogenesis is avoiding the possibility of introducing somaclonal variations in the propagated plants.

 Indirect Somatic Embryogenesis:

In indirect embryogenesis, the cells from explant (excised plant tissues) are made to proliferate and form callus, from which cell suspension cultures can be raised. Certain cells referred to as induced embryo genic determined cells (IEDC) from the cell suspension can form somatic embryos. Embryogenesis is made possible by the presence of growth regulators (in appropriate concentration) and under suitable environmental conditions.

 Somatic embryogenesis (direct or indirect) can be carried on a wide range of media (e.g. MS, White’s). The addition of the amino acid L-glutamine promotes embryogenesis. The presence of auxin such as 2, 4-dichlorophenoxy acetic acid is essential for embryo initiation. On a low auxin or no auxin medium, the embryo genic clumps develop into mature embryos.

Two routes of somatic embryogenesis are known — direct and indirect

 

Indirect somatic embryogenesis is commercially very attractive since a large number of embryos can be generated in a small volume of culture medium. The somatic embryos so formed are synchronous and with good regeneration capability.

 Artificial Seeds from Somatic Embryos:

Artificial seeds can be made by encapsulation of somatic embryos. The embryos, coated with sodium alginate and nutrient solution, are dipped in calcium chloride solution. The calcium ions induce rapid cross-linking of sodium alginate to produce small gel beads, each containing an encapsulated embryo. These artificial seeds (encapsulated embryos) can be maintained in a viable state till they are planted.

 Factors Affecting Micro propagation:

For a successful in vitro clonal propagation (micro propagation), optimization of several factors is needed.

Some of these factors are briefly described:

1. Genotype of the plant:

Selection of the right genotype of the plant species (by screening) is necessary for improved micro propagation. In general, plants with vigorous germination and branching capacity are more suitable for micro- propagation.

2. Physiological status of the explants:

Explants (plant materials) from more recently produced parts of plants are more effective than those from older regions. Good knowledge of donor plants’ natural propagation process with special reference to growth stage and seasonal influence will be useful in selecting explants.

3. Culture media:

The standard plant tissue culture media are suitable for micro propagation during stage I and stage II. However, for stage III, certain modifications are required. Addition of growth regulators (auxins and cytokinins) and alterations in mineral composition are required. This is largely dependent on the type of culture (meristem, bud etc.).

4. Culture environment:

Light:

Photosynthetic pigment in cultured tissues does absorb light and thus influence micro- propagation. The quality of light is also known to influence in vitro growth of shoots, e.g blue light induced bud formation in tobacco shoots. Variations in diurnal illumination also influence micro propagation. In general, an illumination of 16 hours day and 8 hours night is satisfactory for shoot proliferation.

Temperature:

Majority of the culture for micro propagation requires an optimal temperature around 25°C. There are however, some exceptions e.g. Begonia X Cheimantha hybrid tissue grows at a low temperature (around 18°C).

Composition of gas phase:

The constitution of the gas phase in the culture vessels also influences micro propagation. Unorganized growth of cells is generally promoted by ethylene, O2, CO2 ethanol and acetaldehyde.

Factors Affecting in Vitro Rooting:

A general description of the factors affecting micro propagation, particularly in relation to shoot multiplication is given above. For efficient in vitro rooting during micro- propagation, low concentration of salts (reduction to half to one quarter from the original) is advantageous. Induction of roots is also promoted by the presence of suitable auxin (NAA or IBA).

Applications of Micro propagation:

Micro propagation has become a suitable alternative to conventional methods of vegetative propagation of plants. There are several advantages of micro propagation.

 High Rate of Plant Propagation:

Through micro propagation, a large number of plants can be grown from a piece of plant tissue within a short period. Another advantage is that micro propagation can be carried out throughout the year, irrespective of the seasonal variations. Further, for many plants that are highly resistant to conventional propagation, micro propagation is the suitable alternative. The small sized propagules obtained in micro propagation can be easily stored for many years (germplasm storage), and transported across international boundaries.

 Production of Disease-free Plants:

It is possible to produce disease-free plants through micro propagation. Meristem tip cultures are generally employed to develop pathogen-free plants .In fact, micro propagation is successfully used for the production of virus-free plants of sweet potato (Ipomea batatus), cassava (Manihot esculenta) and yam (Discorea rotundata).

Production of Seeds in Some Crops:

Micro propagation, through axillary bud proliferation method, is suitable for seed production in some plants. This is required in certain plants where the limitation for’ seed production is high degree of genetic conservation e.g. cauliflower, onion.

Cost-effective Process:

Micro propagation requires minimum growing space. Thus, millions of plant species can be maintained inside culture vials in a small room in a nursery. The production cost is relatively low particularly in developing countries (like India) where the manpower and labour charges are low.

Automated Micro propagation:

It has now become possible to automate micro propagation at various stages. In fact, bio-reactors have been set up for large scale multiplication of shoots and bulbs. Some workers employ robots (in place of labourers) for micro- propagation, and this further reduces production cost of plants.

Disadvantages of Micro propagation:

Contamination of Cultures:

During the course of micro propagation, several slow-growing microorganisms (e.g. Eswinia sp, Bacillus sp) contaminate and grow in cultures. The microbial infection can be controlled by addition of antibiotics or fungicides. However, this will adversely influence propagation of plants.

Brewing of Medium:

Micro propagation of certain plants (e.g. woody perennials) is often associated with accumulation of growth inhibitory substances in the medium. Chemically, these substances are phenolic compounds, which can turn the medium into dark colour. Phenolic compounds are toxic and can inhibit the growth of tissues. Brewing of the medium can be prevented by the addition of ascorbic acid or citric acid or polyvinyl pyrrolidone to the medium.

Genetic Variability:

When micro propagation is carried out through shoot tip cultures, genetic variability is very low. However, use of adventitious shoots is often associated with pronounced genetic variability.

Vitrification:

During the course of repeated in vitro shoot multiplication, the cultures exhibit water soaked or almost translucent leaves. Such shoots cannot grow and even may die. This phenomenon is referred to as vitrification. Vitrification may be prevented by increasing the agar concentration (from 0.6 to 1%) in the medium. However, increased agar concentration reduces the growth rate of tissues.

Cost Factor:

For some micro propagation techniques, expensive equipment, sophisticated facilities and trained manpower are needed. This limits its use.

In vitro Micrografting

Micrografting is an in vitro grafting technique which involves the placement of a meristem or shoot tip explant onto a decapitated rootstock that has been grown aseptically from seed or micropropagated cultures. Special techniques have been used for increasing the percentage of successful micrografts with the use of growth regulators, etiolation treatments, antioxidants, higher sucrose levels, silicon tubes, etc. The technique has great potential for improvement and large scale multiplication of fruit plants. It has been used on commercial scale for production of virus-free plants in fruit crops and viroid free plants. Micrografting has also been used in prediction of incompatibility between the grafting partners, histological studies, disease indexing, production of disease-free plants particularly resistant to soil borne pathogens and multiplication of difficult to root plants.

Stages of micrografting

Micro-propagation protocol for scion as well as rootstock needs to be standardized separately before performing the micrografting operation under in vitroconditions. Thus, micrografting can be divided into three main stages:

Establishment and multiplication of scion

Shoot or meristem tips intended for grafting can be taken from actively growing shoots in greenhouse, chambers, field or in vitro. Generally, apical shoot tips or nodal cuttings are used as explants for the establishment of in vitro cultures. Following establishment, microshoots are transferred to shoot proliferation medium where shoot number increases by the development of new axillary shoots. Microshoots of desired thickness, age and length are used as scions for in vitro grafting operations.

Establishment and multiplication of rootstock

Rootstocks used for micrografting are in vitro or in vivo germinated seedlings and rooted or unrooted micropropagated shoots. When seedling rootstocks are used and all stages of grafting are conducted in vitro, seeds are surface sterilized and germinated aseptically in vessels containing nutrient salts. The seedlings may be supported on agar medium. Seedlings can also be on a porous substrate, such as sterile vermiculite, which allows the growth of a branched root system. Preparation of rootstock and scion for micrografting Micrografting is affected by cutting off the top of the seedling rootstocks usually just above the cotyledons or top of the micropropagated shoot and placing small shoot apices of scion onto the exposed surface of decapitated rootstock in such a way that the cambium layer or vascular ring of the cut surfaces coincides with each other. This is called surface placement method. Wedge or cleft grafting is performed, incase thickness of rootstock and scion material is large enough to allow making of wedge on the scion material. Firm contact between rootstock and scion is extremely important at the graft junction for proper union of partners and callus formation. Several techniques have been developed for holding grafts together until fusion takes place such as translucent silicon tubing, elastic strip, filter paper bridge, and glass tubing, nylon bands, aluminum foil tubes, dual layer apparatus of aluminum foil and absorbent paper. When grafts are successful, rootstock and scion grow together to produce a plant. It is usually necessary to examine freshly grafted seedlings on a regular basis and remove any adventitious shoot arising on or below the graft union.

Applications of micrografting

§  Virus and viroid elimination

§  Production of plants resistant to pests and diseases

§  Assessment of graft incompatibility

§  Improvement of plant regeneration

§  Mass multiplication

§  Indexing viral diseases

§  Safe germplasm exchange

Production of Disease-Free Plants:

Many plant species are infected with pathogens — viruses, bacteria, fungi, mycoplasma and nematodes that cause systemic diseases. Although these diseases do not always result in the death of plants, they reduce the quality and yield of plants. The plants infected with bacteria and

fungi frequently respond to chemical treatment by bactericides and fungicides. However, it is very difficult to cure the virus-infected plants. Further, viral disease is easily transferred in seed- propagated as well as vegetatively propagated plant species. Plant breeders are always interested to develop disease-free plants, particularly viral disease-free plants. This has become a reality through tissue cultures.

Apical Meristems with Low Concentration of Viruses:

In general, the apical meristems of the pathogen infected and disease harbouring plants are either free or carry a low concentration of viruses, for the following reasons:

1. Absence of vascular tissue in the meristems through which viruses readily move in the plant body.
2. Rapidly dividing meristematic cells with high metabolic activity do not allow viruses to multiply.
3. Virus replication is inhibited by a high concentration of endogenous auxin in shoot apices.

Tissue culture techniques employing meristem-tips are successfully used for the production of disease-free plants, caused by several pathogens — viruses, bacteria, fungi, mycoplasmas.

Methods to Eliminate Viruses in Plants:

In general, plants are infected with many viruses; the nature of some of them may be unknown. The usage virus-free plant implies that the given plant is free from all the viruses, although this may not be always true. The commonly used methods for virus elimination in plants are listed below, and briefly described next.

        I.            Heat treatment of plant

     II.            Meristem-tip culture

  III.            Chemical treatment of media

  IV.            Other in vitro methods

Heat Treatment (Thermotherapy) of Plants:

In the early days, before the advent of meristem cultures, in vivo eradication of viruses from plants was achieved by heat treatment of whole plants. The underlying principle is that many viruses in plant tissues are either partially or completely inactivated at higher temperatures with minimal injury to the host plant. Thermotherapy (at temperatures 35-40°C) was carried out by using hot water or hot air for elimination viruses from growing shoots and buds.

 There are two limitations of viral elimination by heat treatment:

1. Most of the viruses are not sensitive to heat treatment.

2. Many plant species do not survive after thermotherapy.

With the above disadvantages, heat treatment has not become popular for virus elimination.

Meristem-Tip Culture:

A general description of the methodology adopted for meristem and shoot tip cultures has been described. For viral elimination, the size of the meristem used in cultures is very critical. This is due to the fact that most of the viruses exist by establishing a gradient in plant tissues.

 In general, the regeneration of virus-free plants through cultures is inversely proportional to the size of the meristem used. The meristem-tip explant used for viral elimination cultures is too small. A stereoscopic microscope is usually employed for this purpose.

Meristem-tip cultures are influenced by the following factors:

1. Physiological condition of the explant — actively growing buds are more effective.
2. Thermotherapy prior to meristem-tip culture — for certain plants (possessing viruses in the meristematic regions), heat treatment is first given and then the meristem-tips are isolated and cultured.
3. Culture medium —MS medium with low concentrations of auxins and cytokinins is ideal.

Chemical Treatment of Media:

Some workers have attempted to eradicate viruses from infected plants by chemical treatment of the tissue culture media. The commonly used chemicals are growth substances (e.g. cytokinins) and antimetabolites (e.g thiouracil, acetyl salicylic acid).

There are however, conflicting reports on the elimination viruses by chemical treatment of the media. For instance, addition of cytokinin suppressed the multiplication of certain viruses while for some other viruses, it actually stimulated.

Other in Vitro Methods:

Besides meristem-tip culture, other in vitro methods are also used for raising virus-free plants. In this regard callus cultures have been successful to some extent. The callus derived from the infected tissue does not carry the pathogens throughout the cells. In fact, the uneven distribution of tobacco mosaic virus in tobacco leaves was exploited to develop virus-free plants of tobacco. Somatic cell hybridization, gene transformation and somaclonal variations also useful to raise disease-free plants.

Elimination of Pathogens Other than Viruses:

Besides the elimination of viruses, meristem-tip cultures and callus cultures are also useful for eradication bacteria, fungi and mycoplasmas. Some examples are given

1. The fungus Fusarium roseum has been successfully eliminated through meristem cultures from carnation plants.

2. Certain bacteria (Pseudomonas carophylli, Pectobacterium parthenii) are eradicated from carnation plants by using meristem cultures.

Merits and Demerits of Disease-Free Plant Production:

Among the culture techniques, meristem-tip culture is the most reliable method for virus and other pathogen elimination. This, however, requires good knowledge of plant pathology and tissue culture. Virus-free plants exhibit increased growth and vigour of plants, higher yield (e.g. potato), increased flower size (e.g. Chrysanthemum), and improved rooting of stem cuttings (e.g. Pelargonium) Virus-free plants are more susceptible to the same virus when exposed again. This is the major limitation. Reinfection of disease-free plants can be minimized with good knowledge of greenhouse maintenance.

Callus and Suspension Cultures

Callus is an unorganized, proliferative mass of differentiated plant cells, and usually occurs naturally as wound response. Tissues and cells cultured on an agar-gelled medium form an unorganised mass of cells is also called callus. It can be induced through culture of plant tissue on a medium usually containing relatively high levels of auxin, especially 2,4-D.

However, because of the phase of disorganization that occurs, plants regenerating from callus, can be prone to genetic change.Callus cultures need to be sub-cultured every 3-5 weeks in view of cell growth, nutrient depletion and medium drying. Therefore, calluses are easy to maintain and are the most widely used. When explants are cultured on a suitable PGR(s) combination, many of its cells undergo division. Even mature and certain differentiated, e.g., parenchyma and often colenchyma, cells undergo changes to become meristematic; this is called dedifferentiation.

Dedifferentiation involves, among other things, renewed and enhanced RNA and protein syntheses leading to the formation of new cellular components needed for meristematic activity. Initially, cell divisions are confined to the cut ends, but subsequently it covers the entire explant. The resulting cell mass is ordinarily unorganised, but it often consists of several cell types including fibers, and vascular elements.

Suspension Cultures:

Tissues and cells cultured in a liquid medium produce a suspension of single cells and cells clumps of few to many cells; these are called suspension cultures. Liquid cultures must be constantly agitated, generally by a gyratory shaker at 100-250 rpm (revolution per minute), to facilitate aeration and dissociation of cell clumps into smaller pieces.

Suspension cultures grow much faster than callus cultures, need to be sub-cultured about every week, allow a more accurate determination of the nutritional requirements of cells and are the only system amenable to scaling up for a large-scale production of cells and even somatic embryos (SEs). The suspension cultures are broadly grouped as follows: (1) batch cultures, (2) continuous cultures, and (3) immobilized cell cultures.

Batch Cultures:

In a batch culture, the same medium and all the cells produced are retained in the culture vessel, e.g., culture flasks (100-250 ml), fermenters (variable size), etc. The cell number or biomass of a batch culture exhibits a typical sigmoidal curve, having a lag phase during which the cell number or biomass remains unchanged, followed by a logarithmic (log) phase when there is a rapid increase in cell number and, finally, ending in a stationary phase during which cell number does not change.

The lag phase duration depends mainly on inoculum size and growth phase of the culture from which the inoculum is taken. The log phase lasts about 3-4 cell generations (a cell generation is the time taken for doubling of cell number), and the duration of a cell generation may vary from 22-48 hr, depending mainly on the plant species. The stationary phase is forced on the culture by depletion of the nutrients and possibly due to an accumulation of cellular wastes. If the culture is kept in stationary phase for a prolonged period, the cells may die.

 

Initiation of callus and suspension cultures

Batch cultures are maintained by sub-culturing. They are used for initiation of cell suspensions, which may be used for cloning, cell selection or as seed cultures for scaling up or for continuous cultures.

They are, however, unsuitable for studies on cell growth and metabolism because there is a constant change in cell density and nutritional status of the medium. But batch cultures are much more convenient than continuous cultures and, as a result, are routinely used.

A Model curve for cell number in a batch culture

Continuous Cultures:

In a continuous culture, the cell population is maintained in a steady state by regularly replacing a portion of the used or spent medium by fresh medium. Such culture systems are of either (1) closed or (2) open type. In a closed continuous culture, cells are separated from the used medium taken out for replacement, and added back to the culture so that cell biomass keeps on increasing. In contrast, both cells and the used medium are taken out from open continuous cultures and replaced by equal volume of fresh medium. The replacement volume is so adjusted that cultures remain at submaximal growth indefinitely.

The open cultures are of either turbidostat or chemostat types. In a turbidostat, cells are allowed to grow upto a preselected turbidity (usually, measured as OD) when a predetermined volume of the culture is replaced by fresh normal culture medium. But in a chemostat, a chosen nutrient is kept in a concentration so that it is depleted very rapidly to become growth limiting, while other nutrients are still in concentrations higher than required. In such a situation, any addition of the growth-limiting nutrient is reflected in cell growth. Chemostats are ideal for the determination of effects of individual nutrients on cell growth and metabolism.

Immobilized Cell Cultures:

Plant cells and cell groups may be encapsulated in a suitable material, e.g., agarose and calcium alginate gels, or entrapped in membranes or stainless steel screens. The gel beads containing cells may be packed in a suitable column or, alternatively, cells may be packed in a column of a membrane or wire cloth.

Liquid medium is continuously run through the column to provide nutrients and aeration to cells. Immobilization of cells changes their cellular physiology in comparison to suspension culture cells; this offers several advantages for their use in biochemical production, but they are usually not used for other studies.

Subculture:

After a period of time, it becomes necessary to transfer organs and tissues to fresh media chiefly due to nutrient depletion and medium drying. This is particularly true of tissue and cell cultures where a portion of tissue is used to inoculate new culture tubes or flasks; this is known as sub-culturing. In general, callus cultures are sub-cultured every 4-6 weeks, while suspension cultures need to be sub-cultured every 3-14 days. Plant cell and tissue cultures may be maintained indefinitely by serial sub-culturing.

In case of suspension cultures, sub-culturing should be done about or somewhat prior to the time of their maximum growth. The inoculums volume should be 20-25% of the fresh medium volume; in any case, the initial cell density of the fresh culture (just after inoculation) should be around 5 x 104 cells m1-1 or higher otherwise the cells may fail to divide.

Estimation of Growth:

Cell number is the most informative measure of cell growth. This measurement is applicable to only suspension cultures, and even their cell aggregates must be treated, e.g., with pectinase, to dissociate them into single cells before counting the cell number in a haemocytometer.

Therefore, cell number is estimated only where information obtained justifies the efforts. In contrast, packed cell volume of suspension cultures is easily determined by pipetting a known volume into a 15 ml graduated centrifuge tube, spinning at 200 ×g for 5 min and reading the volume of cell pellet, which is expressed as ml cells/1 of culture.

 Culture fresh and dry weights are the most commonly used measures of growth of both suspension and callus cultures. In case of callus cultures, the cell mass is placed on a preweighed dry filter paper or nylon filter and weighed to determine fresh weight.

Cells from suspension cultures are filtered onto a filter paper or nylon filter, washed with distilled water, excess water removed under vacuum and weighed along with the filter; the filter is preweighed in wet condition. For dry weight determination, the cells and the filter are dried in an oven at 60°C for 12 hr and weighed; the filter is pre-weighed in dry condition. Cell fresh and dry weights may either be expressed as per ml (suspension culture) or per culture.

Nuclear Cytology:

Callus and suspension cultures show both numerical (polyploidy and aneuploidy) and structural (deletions, translocations, etc.) chromosome changes. The frequency of these changes tends to increase with the duration of in vitro culture so that some cultures may become predominantly or even completely polyploid or aneuploid.

Explants contain endopolyploid cells, which may give rise to a portion of the polyploid cells in cultures. But most polyploid cells appear to originate through endoreduplication (additional rounds of DNA replication without intervening cell division) although selection for such cells cannot be ruled out.

Aneuploid cells originate mainly due to anaphase irregularities like unequal chromatid separation, lagging chromatids or chromosomes, anaphase bridges giving rise to breakage-fusion-bridge cycle, chromosome fragmentation, etc.

The cytogenetic status of cultured cells is influenced by several factors of the culture system, e.g., GR concentrations and combination, culture age, liquid or agar medium, subculture interval, sucrose concentration, etc. Suspension cultures of many diploid species show a selection for diploid cells so that they remain predominantly diploid for long periods, e.g., in case of Vicia hajastana and Haplopappus gracilis cultures remained predominantly diploid for over 300 days.

Secondary metabolite production through Cell suspension cultures

Plant cell and tissue cultures can be established routinely under sterile conditions from explants, such as plant leaves, stems, roots, and meristems for multiplication and extraction of secondary metabolites. Strain improvement, methods for the selection of high-producing cell lines, and medium optimizations can lead to an enhancement in secondary metabolite production.

The capacity for plant cell, tissue, and organ cultures to produce and accumulate many of the same valuable chemical compounds as the parent plant in nature has been recognized almost since the inception of in vitro technology. The strong and growing demand in today's marketplace for natural, renewable products has refocused attention on in vitro plant materials as potential factories for secondary phytochemical products and has paved the way for new research exploring secondary product expression in vitro. There is a series of distinct advantages to producing a valuable secondary product in plant cell culture, rather than in vivo in the whole crop plant.

These include the following:

• Production can be more reliable, simpler, and more predictable.
• Isolation of the phytochemical can be rapid and efficient, when compared with extraction from complex whole plants
• Compounds produced in vitro can directly parallel compounds in the whole plant.
• Interfering compounds that occur in the field-grown plant can be avoided in cell cultures.
• Tissue and cell cultures can yield a source of defined standard phytochemicals in large volumes.
• Tissue and cell cultures are a potential model to test elicitation.
• Cell cultures can be radiolabeled, such that the accumulated secondary products, when provided as feed to laboratory animals, can be traced metabolically.

While research to date has succeeded in producing a wide range of valuable secondary phytochemicals in unorganized callus or suspension cultures, in some cases production requires more differentiated micro plant or organ cultures. This situation often occurs when the metabolite of interest is only produced in specialized plant tissues or glands in the parent plant. A prime example is ginseng (Panax ginseng). Because saponin and other valuable metabolites are specifically produced in ginseng roots, root culture is required in vitro. Similarly, herbal plants such as Hypericum perforatum (St. John's wort), which accumulates the hypericins and hyperforins in foliar glands, have not demonstrated the ability to accumulate phytochemicals in undifferentiated cells. As another example, biosynthesis of lysine to anabasine occurs in tobacco (Nicotiana tabacum) roots, followed by the conversion of anabasine to nicotine in leaves. Callus and shoot cultures of tobacco can produce only trace amounts of nicotine because they lack the organ-specific compound anabasine. In other cases, at least some degree of differentiation in a cell culture must occur before a product can be synthesized (e.g., vincristine or vinblastine from Catharanthus roseus). Reliance of a plant on a specialized structure for production of a secondary metabolite, in some cases, is a mechanism for keeping a potentially toxic compound sequestered. Intensive activities have been centered on production of natural drugs or chemoprotective compounds from plant cell culture by one or more of the following strategies:

Accumulation of secondary metabolites in plant cell cultures for plant cell culture techniques to become economically viable, it is important to develop methods that would allow for consistent generation of high yields of products from cultured cells. Careful selection of productive cells and cultural conditions resulted in accumulation of several products in higher levels in cultured cells.

In order to obtain yields in high concentrations for commercial exploitation, efforts have focused on the stimulation of biosynthetic activities of cultured cells using various methods - Culture productivity is critical to the practical application of plant cell culture technology to production of plant-specific bioactive metabolites. Until now, various strategies have been developed to improve the production of secondary metabolites using plant cell cultures. The tissue culture cells typically accumulate large amounts of secondary compounds only under specific conditions. That means maximization of the production and accumulation of secondary metabolites by plant tissue cultured cells requires

1. manipulating the parameters of the environment and medium,
2. selecting high yielding cell clones,
3. precursor feeding, and
4. elicitation.

Optimization of cultural conditions:

Number of chemical and physical factors like media components, phytohormones, pH, temperature, aeration, agitation, light affecting production of secondary metabolites has been extensively studied. Several products were found to be accumulating in cultured cells at a higher level than those in native plants through optimization of cultural conditions. Manipulation of physical aspects and nutritional elements in a culture is perhaps the most fundamental approach for optimization of culture productivity. For example, ginsenosides by Panax ginseng, rosmarinic acid by Coleus bluemei, shikonin by Lithospermum erythrorhizon, ubiquinone-10 by Nicotiana tabacum, berberin by Coptis japonica, were accumulated in much higher levels in cultured cells than in the intact plants.

Selection of high-producing strains:

Plant cell cultures represent a heterogeneous population in which physiological characteristics of individual plant cells are different. Synthesis of several products in high amounts using selection and screening of plant cell cultures have been already described by many workers. Cell cloning methods provide a promising way of selecting cell lines yielding increased levels of product.

Precursor feeding:

Exogenous supply of a biosynthetic precursor to culture medium may also increase the yield of the desired product. This approach is useful when the precursors are inexpensive. The concept is based on the idea that any compound, which is an intermediate, in or at the beginning of a secondary metabolite biosynthetic route, stands a good chance of increasing the yield of the final product. Attempts to induce or increase the production of plant secondary metabolites, by supplying precursor or intermediate compounds, have been effective in many cases. For example, amino acids have been added to cell suspension culture media for production of tropane alkaloids, indole alkaloids etc. Addition of phenylalanine to Salvia officinalis cell suspension cultures stimulated the production of rosmarinic acid. Addition of the same precursor resulted stimulation of taxol production in Taxus cultures. Feeding ferulic acid to cultures of Vanilla planifolia resulted in increase in vanillin accumulation. Furthermore, addition of leucine, led to enhancement of volatile monoterpenes in cultures of Perilla frutiscens, where as addition of geraniol to rose cell cultures led to accumulation of nerol and citronellol.

Elicitation:

Plants produce secondary metabolites in nature as a defense mechanism against attack by pathogens. Elicitors are signals triggering the formation of secondary metabolites. Use of elicitors of plant defense mechanisms, i.e. elicitation, has been one of the most effective strategies for improving the productivity of bioactive secondary metabolites. Biotic and abiotic elicitors which are classified on their origin are used to stimulate secondary metabolite formation in plant cell cultures, thereby reducing the process time to attain high product concentrations. Production of many valuable secondary metabolites using various elicitors were also reported

Conclusions

The use of plant cell culture for the production of chemicals and pharmaceuticals has made great strides building on advances in plant science. The increased use of genetic tools and an emerging picture of the structure and regulation of pathways for secondary metabolism will provide the basis for the production of commercially acceptable levels of product. The increased appeal of natural products for medicinal purposes coupled with the low product yields and supply concerns of plant harvesation has renewed interest in large-scale plant cell culture technology. Knowledge of biosynthetic pathways of desired compounds in plants as well as in cultures is often still in its infancy, and consequently, strategies are needed to develop an information based on a cellular and molecular level. Because of the complex and incompletely understood nature of plant cells in in vitro cultures, case-by-case studies have been used to explain the problems occurring in the production of secondary metabolites from cultured plant cells. These results show that plant cell culture systems have potential for commercial exploitation of secondary metabolites. The introduction of the techniques of molecular biology, so as to produce transgenic cultures and to effect the expression and regulation of biosynthetic pathways, is also likely to be a significant step towards making cell cultures more generally applicable to the commercial production of secondary metabolites.

Protoplast isolation, culture and regeneration

Protoplasts are naked plant cells without the cell wall, but they possess plasma membrane and all other cellular components. They represent the functional plant cells but for the lack of the barrier, cell wall.  Protoplasts of   different   species   can be fused to generate a hybrid and this process   is   referred   to   as   somatic   hybridization (or   protoplast   fusion).   Cybridization   is   the phenomenon of fusion of a normal protoplast with an enucleated (without nucleus) protoplast that results in the formation of a cybrid or cytoplast (cytoplasmic hybrids).

Historical developments:

The term protoplast was introduced in 1880 by Hanstein. The first isolation of protoplasts was achieved by Klercker (1892) employing a mechanical method. A real beginning in protoplast research was made in 1960 by Cocking who used an enzymatic method for the removal of cell wall.

Rakabe and his associates (1971) were successful to achieve the regeneration of whole tobacco plant from protoplasts. Rapid progress occurred after 1980 in protoplast fusion to improve plant genetic material, and the development of transgenic plants.

Importance of Protoplasts and Their Cultures:

The isolation, culture and fusion of protoplasts is a fascinating field in plant research. Protoplast isolation and their cultures provide millions of single cells (comparable to microbial cells) for a variety of studies.

Protoplasts have a wide range of applications; some of them are listed below:

1. The protoplast in culture can be regenerated into a whole plant.
2. Hybrids can be developed from protoplast fusion.
3. It is easy to perform single cell cloning with protoplasts.
4. Genetic transformations can be achieved through genetic engineering of protoplast DNA.
5. Protoplasts are excellent materials for ultra-structural studies.
6. Isolation of cell organelles and chromosomes is easy from protoplasts.
7. Protoplasts are useful for membrane studies (transport and uptake processes).
8. Isolation of mutants from protoplast cultures is easy.

Isolation of Protoplasts:

Protoplasts are isolated by two techniques

1)      Mechanical method

2)      Enzymatic method

Mechanical Method:

Protoplast isolation by mechanical method is a crude and tedious procedure. This results in the isolation of a very small number of protoplasts.

The technique involves the following stages:

1. A small piece of epidermis from a plant is selected.
2. The cells are subjected to plasmolysis. This causes protoplasts to shrink away from the cell walls.
3. The tissue is dissected to release the protoplasts.

Mechanical method for protoplast isolation is no more in use because of the following limitations:

1. Yield of protoplasts and their viability is low.
2. It is restricted to certain tissues with vacuolated cells.
3. The method is laborious and tedious.

However, some workers prefer mechanical methods if the cell wall degrading enzymes (of enzymatic method) cause deleterious effects to protoplasts.

Enzymatic Method:

Enzymatic method is a very widely used technique for the isolation of protoplasts. The advantages of enzymatic method include good yield of viable cells, and minimal or no damage to the protoplasts.

Sources of protoplasts:

Protoplasts can be isolated from a wide variety of tissues and organs that include leaves, roots, shoot apices, fruits, embryos and microspores. Among these, the mesophyll tissue of fully expanded leaves of young plants or new shoots are most frequently used. In addition, callus and suspension cultures also serve as good sources for protoplast isolation.

Enzymes for protoplast isolation:

Different enzyme preparations are available in the market but the idea is to combine one middle lamella dissolving and one cell wall digesting enzymes in proper composition to achieve maximum protoplasts release from one gm. material.

Following enzymes are used:

1. Macerozymes R-10
2. Cellulase – Onozuka R-10
3. Hemicellulose
4. Pectinase
5. Drieselase

A combination of these enzymes in a concentration of 0.5-2% is used. In many cases only macerozyme and cellulase are sufficient to obtain protoplasts in significant number. The enzyme solution (pH 5.5) is prepared in 10-15% sorbitol or mannitol containing small amount of CaCl2 (7 mM) for membrane stability.

This solution is sterilized through a membrane filter (cold sterilization) and leaf or callus tissues are placed in it. Petri-plates containing tissue and enzyme mixture are sealed with parafilm and incubated for 4-12 hours (sometimes 0.5 to 20 hrs.) on a rocking shaker at 24-26 °C.

After incubation, solution is filtered through a wire or nylon mesh (50-100 µm) to remove debris (undigested cells, tissues, broken cells etc.), transferred into screw capped small centrifuge tubes (sterilized), and centrifuged at 100 g. The protoplasts formed a pellet while the debris in the supernatant is carefully removed.

Fresh sterilized sorbitol solution (no enzyme) is added to tube and centrifuged. By repeating the process 2 to 3 times, protoplasts are cleaned (debris is removed). If 20% sucrose solution is used, protoplasts will float and debris will settle during centrifugation at 200g for 1 min.

Floating protoplasts are carefully removed with the help of sterilized pipette and bulked together for further use. Protoplasts are counted by haemocytometer and then diluted to proper strength (number per ml) in the culture medium containing osmoticum.

The enzymes that can digest the cell walls are required for protoplast isolation. Chemically, the plant cell wall is mainly composed of cellulose, hemicellulose and pectin which can be respectively degraded by the enzymes cellulose, hemicellulose and pectinase. In fact, the various enzymes for protoplast isolation are commercially available. The enzymes are usually used at a pH 4.5 to 6.0, temperature 25-30°C with a wide variation in incubation period that may range from half an hour to 20 hours.

The enzymatic isolation of protoplasts can be carried out by two approaches:

1. Two step or sequential method:

The tissue is first treated with pectinase (macerozyme) to separate cells by degrading middle lamella. These free cells are then exposed to cellulose to release protoplasts. Pectinase breaks up the cell aggregates into individual cells while cellulose removes the cell wall proper.

2. One step or simultaneous method:

This is the preferred method for protoplast isolation. It involves the simultaneous use of both the enzymes — macerozyme and cellulose.

Isolation of protoplasts from leaves:

Leaves are most commonly used, for protoplast isolation, since it is possible to isolate uniform cells in large numbers.

The procedure broadly involves the following steps:

1. Sterilization of leaves.
2. Removal of epidermal cell layer.
3. Treatment with enzymes.
4. Isolation of protoplasts.

Besides leaves, callus cultures and cell suspension cultures can also be used for the isolation of protoplasts. For this purpose, young and actively growing cells are preferred.

Purification of protoplasts:

The enzyme digested plant cells, besides protoplasts contain undigested cells, broken protoplasts and undigested tissues. The cell clumps and undigested tissues can be removed by filtration. This is followed by centrifugation and washings of the protoplasts. After centrifugation, the protoplasts are recovered above Percoll.

Viability of protoplasts:

It is essential to ensure that the isolated protoplasts are healthy and viable so that they are capable of undergoing sustained cell divisions and regeneration.

There are several methods to assess the protoplast viability:

1. Fluorescein diacetate (FDA) staining method—The dye accumulates inside viable protoplasts which can be detected by fluorescence microscopy.
2. Phenosafranine stain is selectively taken up by dead protoplasts (turn red) while the viable cells remain unstained.
3. Exclusion of Evans blue dye by intact membranes.
4. Measurement of cell wall formation—Calcofluor white (CFW) stain binds to the newly formed cell walls which emit fluorescence.
5. Oxygen uptake by protoplasts can be measured by oxygen electrode.
6. Photosynthetic activity of protoplasts.
7. The ability of protoplasts to undergo continuous mitotic divisions (this is a direct measure).

Culture of Protoplasts:

The very first step in protoplast culture is the development of a cell wall around the membrane of the protoplast. This is followed by the cell divisions that give rise to a small colony. With suitable manipulations of nutritional and physiological conditions, the cell colonies may be grown continuously as cultures or regenerated to whole plants. Protoplasts are cultured either in semisolid agar or liquid medium. Sometimes, protoplasts are first allowed to develop cell wall in liquid medium, and then transferred to agar medium.

Agar culture:

Agarose is the most frequently used agar to solidify the culture media. The concentration of the agar should be such that it forms a soft agar gel when mixed with the protoplast suspension. The plating of protoplasts is carried out by Bergmann’s cell plating technique .In agar cultures, the protoplasts remain in a fixed position, divide and form cell clones. The advantage with agar culture is that clumping of protoplasts is avoided.

Liquid culture:

Liquid culture is the preferred method for protoplast cultivation for the following reasons:

1. It is easy to dilute and transfer.
2. Density of the cells can be manipulated as desired.
3. For some plant species, the cells cannot divide in agar medium, therefore liquid medium is the only choice.
4. Osmotic pressure of liquid medium can be altered as desired.

Culture Media:

The culture media with regard to nutritional components and osmoticum are briefly described.

Nutritional components:

In general, the nutritional requirements of protoplasts are similar to those of cultured plant cells (callus and suspension cultures). Mostly, MS and B5 media with suitable modifications are used.

Some of the special features of protoplast culture media are listed below:

1. The medium should be devoid of ammonium, and the quantities of iron and zinc should be less.
2. The concentration of calcium should be 2-4-times higher than used for cell cultures. This is needed for membrane stability.
3. High auxin/kinetin ratio is suitable to induce cell divisions while high kinetin/auxin ratio is required for regeneration.
4. Glucose is the preferred carbon source by protoplasts although a combination of sugars (glucose and sucrose) can be used.
5. The vitamins used for protoplast cultures are the same as used in standard tissue culture media.

Osmoticum and osmotic pressure:

Osmoticum broadly refers to the reagents/ chemicals that are added to increase the osmotic pressure of a liquid. The isolation and culture of protoplasts require osmotic protection until they develop a strong cell wall. In fact, if the freshly isolated protoplasts are directly added to the normal culture medium, they will burst. Thus, addition of an osmoticum is essential for both isolation and culture media of protoplast to prevent their rupture. The osmotica are of two types — non-ionic and ionic.

Non-ionic osmotica:

The non-ionic substances most commonly used are soluble carbohydrates such as mannitol, sorbitol, glucose, fructose, galactose and sucrose. Mannitol, being metabolically inert, is most frequently used.

Ionic osmotica:

Potassium chloride, calcium chloride and magnesium phosphate are the ionic substances in use to maintain osmotic pressure. When the protoplasts are transferred to a culture medium, the use of metabolically active osmotic stabilizers (e.g., glucose, sucrose) along with metabolically inert osmotic stabilizers (mannitol) is advantageous. As the growth of protoplasts and cell wall regeneration occurs, the metabolically active compounds are utilized, and this results in the reduced osmotic pressure so that proper osmolarity is maintained.

Culture Methods:

The culture techniques of protoplasts are almost the same that are used for cell culture with suitable modifications. Some important aspects are briefly given.

Feeder layer technique:

For culture of protoplasts at low density feeder layer technique is preferred. This method is also important for selection of specific mutant or hybrid cells on plates. The technique consists of exposing protoplast cell suspensions to X-rays (to inhibit cell division with good metabolic activity) and then plating them on agar plates.

Co-culture of protoplasts:

Protoplasts of two different plant species (one slow growing and another fast growing) can be co- cultured. This type of culture is advantageous since the growing species provide the growth factors and other chemicals which help in the generation of cell wall and cell division.

The co-culture method is generally used if the two types of protoplasts are morphologically distinct.

Micro drop culture:

Specially designed dishes namely cuprak dishes with outer and inner chambers are used for micro drop culture. The inner chamber carries several wells wherein the individual protoplasts in droplets of nutrient medium can be added. The outer chamber is filled with water to maintain humidity. This method allows the culture of fewer protoplasts for droplet of the medium.

Regeneration of Protoplasts:

Protoplast regeneration which may also be regarded as protoplast development occurs in two stages:

1. Formation of cell wall.
2. Development of callus/whole plant.

Formation of cell wall:

The process of cell wall formation in cultured protoplasts starts within a few hours after isolation that may take two to several days under suitable conditions. As the cell wall development occurs, the protoplasts lose their characteristic spherical shape. The newly developed cell wall by protoplasts can be identified by using calcofluor white fluorescent stain.

The freshly formed cell wall is composed of loosely bound micro fibrils which get organized to form a typical cell wall. This process of cell wall development requires continuous supply of nutrients, particularly a readily metabolised carbon source (e.g. sucrose). Cell wall development is found to be improper in the presence of ionic osmotic stabilizers in the medium. The protoplasts with proper cell wall development undergo normal cell division. On the other hand, protoplasts with poorly regenerated cell wall show budding and fail to undergo normal mitosis.

Development of Callus/whole Plant:

As the cell wall formation around protoplasts is complete, the cells increase in size, and the first division generally occurs within 2-7 days. Subsequent divisions result in small colonies, and by the end of third week, visible colonies (macroscopic colonies) are formed. These colonies are then transferred to an osmotic-free (mannitol or sorbitol-free) medium for further development to form callus. With induction and appropriate manipulations, the callus can undergo organogenic or embryo genic differentiation to finally form the whole plant. A general view of the protoplast isolation, culture and regeneration is represented in Fig given below.

Plant regeneration can be done from the callus obtained either from protoplasts or from the culture of plant organs. There are however, certain differences in these two calluses. The callus derived from plant organs carries preformed buds or organized structures, while the callus from protoplast culture does not have such structures.

Sub-Protoplasts:

The fragments derived from protoplasts that do not contain all the contents of plant cells are referred to as sub-protoplasts. It is possible to experimentally induce fragmentation of protoplasts to form sub-protoplasts. This can be done by application of different centrifugal forces created by discontinuous gradients during centrifugation. Exposure of protoplasts to cytochalasin B in association with centrifugation is a better approach for fragmentation of protoplasts.

There are three types of sub-protoplasts:

1. Mini-protoplasts:

These are also called as karyoplasts and contain the nucleus. Mini-protoplasts can divide and are capable of regeneration into plants.

2. Cytoplasts:

These are sub-protoplasts containing the original cytoplasmic material (in part or full) but lack nucleus. Thus, cytoplasts are nuclear-free sub-protoplasts which cannot divide, but they can be used for cybridization.

3. Micro-protoplasts:

This term was suggested for sub-protoplasts that contain not all but a few chromosomes.

Somatic Hybridization: Aspects, Applications and Limitations

The conventional method to improve the characteristics of cultivated plants, for years, has been sexual hybridization. The major limitation of sexual hybridization is that it can be performed within a plant species or very closely related species. This restricts the improvements that can be done in plants.

The species barriers for plant improvement encountered in sexual hybridization can be overcome by somatic cell fusion that can form viable hybrids. Somatic hybridization broadly involves in vitro fusion of isolated protoplasts to form a hybrid cell and its subsequent development to form a hybrid plant.

Plant protoplasts are of immense utility in somatic plant cell genetic manipulations and improvement of crops. Thus, protoplasts provide a novel opportunity to create cells with new genetic constitution. And protoplast fusion is a wonderful approach to overcome sexual incompatibility between different species of plants. More details on the applications of somatic hybridization are given later.

Somatic hubridization involves the following aspects:

1. Fusion of protoplasts
2. Selection of hybrid cells
3. Identification of hybrid plants.

A. Fusion of Protoplasts:

As the isolated protoplasts are devoid of cell walls, there in vitro fusion becomes relatively easy. There are no barriers of incompatibility (at interspecific, inter-generic or even at inter-kingdom levels) for the protoplast fusion. Protoplast fusion that involves mixing of protoplasts of two different genomes can be achieved by spontaneous, mechanical, or induced fusion methods.

Spontaneous fusion:

Cell fusion is a natural process as is observed in case of egg fertilization. During the course of enzymatic degradation of cell walls, some of the adjoining protoplasts may fuse to form homokaryocytes (homokaryons). These fused cells may sometimes contain high number of nuclei (2-40).

This is mainly because of expansion and subsequent coalescence of plasmodermal connections between cells. The frequency of homokaryon formation was found to be high in protoplasts isolated from dividing cultured cells. Spontaneously fused protoplasts, however, cannot regenerate into whole plants, except undergoing a few cell divisions.

Mechanical fusion:

The protoplasts can be pushed together mechanically to fuse. Protoplasts of Lilium and Trillium in enzyme solutions can be fused by gentle trapping in a depression slide. Mechanical fusion may damage protoplasts by causing injuries.

Induced fusion:

Freshly isolated protoplasts can be fused by induction. There are several fusion-inducing agents which are collectively referred to as fusogens e.g. NaN03, high pH/Ca2+, polyethylene glycol, polyvinyl alcohol, lysozyme, concavalin A, dextran, dextran sulfate, fatty acids and esters, electro fusion. Some of the fusogens and their use in induced fusion are described.

A diagrammatic representation of protoplast fusion is given below

Treatment with sodium nitrate:

The isolated protoplasts are exposed to a mixture of 5.5% NaNO3 in 10% sucrose solution. Incubation is carried out for 5 minutes at 35°C, followed by centrifugation (200 x g for 5 min). The protoplast pellet is kept in a water bath at 30°C for about 30 minutes, during which period protoplast fusion occurs. NaNO3 treatment results in a low frequency of heterokaryon formation, particularly when mesophyll protoplasts are fused.

High pH and high Ca2+ ion treatment:

This method was first used for the fusion of tobacco protoplasts, and is now in use for other plants also. The method consists of incubating protoplasts in a solution of 0.4 M mannitol containing 0.05 M CaCI2 at pH 10.5 (glycine-NaOH buffer) and temperature 3 7°C for 30-40 minutes. The protoplasts form aggregates, and fusion usually occurs within 10 minutes. By this method, 20-50% of the protoplasts are involved in fusion.

Polyethylene glycol (PEG) treatment:

This has become the method of choice, due to its high success rate, for the fusion of protoplasts from many plant species. The isolated protoplasts in culture medium (1 ml) are mixed with equal volume (1 ml) of 28-56% PEG (mol. wt. 1500-6000 Daltons) in a tube. PEG enhances fusion of protoplasts in several species. This tube is shaken and then allowed to settle.

The settled protoplasts are washed several times with culture medium.

PEG treatment method is widely used protoplast fusion as it has several advantages:

1. It results in a reproducible high-frequency of heterokaryon formation.
2. Low toxicity to cells.
3. Reduced formation of bi-nucleate heterokaryons.
4. PEG-induced fusion is non-specific and therefore can be used for a wide range of plants.

Electro-fusion:

In this method, electrical field is used for protoplast fusion. When the protoplasts are placed in a culture vessel fitted with micro- electrodes and an electrical shock is applied, protoplasts are induced to fuse. Electro-fusion technique is simple, quick and efficient and hence preferred by many workers.

Further, the cells formed due to electro-fusion do not show cytotoxic responses as is the case with the use of fusogens (including PEG). The major limitation of this method is the requirement of specialized and costly equipment.

Mechanism of fusion:

The fusion of protoplasts involves three phases agglutination, plasma membrane fusion and formation of heterokaryons.

1. Agglutination (adhesion):

When two protoplasts are in close contact with each other, adhesion occurs. Agglutination can be induced by fusogens e.g. PEG, high pH and high Ca2+.

2. Plasma membrane fusion:

Protoplast membranes get fused at localized sites at the points of adhesion. This leads to the formation of cytoplasmic bridges between protoplasts. The plasma membrane fusion can be increased by high pH and high Ca2+, high temperature and PEC, as explained below.

a)      High pH and high Ca2+ ions neutralize the surface charges on the protoplasts. This allows closer contact and membrane fusion between agglutinated protoplasts.

b)      High temperature helps in the intermingling of lipid molecules of agglutinated protoplast membranes so that membrane fusion occurs.

c)      PEG causes rapid agglutination and formation of clumps of protoplasts. This results in the formation of tight adhesions of membranes and consequently their fusion.

3. Formation of heterokaryons:

The fused protoplasts get rounded as a result of cytoplasmic bridges leading to the formation of spherical homokaryon or heterokaryon.

B. Selection of Hybrid Cells:

About 20-25% of the protoplasts are actually involved in the fusion. After the fusion process, the protoplast population consists of a heterogenous mixture of un-fused chloroplasts, homokaryons and heterokaryons as shown below. It is therefore necessary to select the hybrid cells (heterokaryons). The commonly used methods employed for the selection of hybrid cells are biochemical, visual and cytometric methods.

Biochemical methods:

The biochemical methods for selection of hybrid cells are based on the use of biochemical compounds in the medium (selection medium). These compounds help to sort out the hybrid and parental cells based on their differences in the expression of characters.

Drug sensitivity and auxotrophic mutant selection methods are described below:

1. Drug sensitivity:

This method is useful for the selection hybrids of two plant species, if one of them is sensitive to a drug. Protoplasts of Petunia hybride (species A) can form macroscopic callus on MS medium, but are sensitive to (inhibited by) actinomycin D. Petunia parodii protoplasts (species B) form small colonies, but are resistant to actinomycin D.

When these two species are fused, the fused protoplasts derive both the characters — formation of macroscopic colonies and resistance to actinomycin D on MS medium. This helps in the selection of hybrids. The parental protoplasts of both the species fail to grow. Protoplasts of P. parodii form very small colonies while that of P. hybrida are inhibited by actinomycin D.

 

 Drug sensitivity technique was originally developed by Power et al (1976) for the selection of hybrids of Petunia sp. A similar procedure is in use for the selection of other somatic hybrids e.g., hybrids between Nicotiana Silvestre’s and Nicotiana knightiana.

2. Auxotrophic mutants:

Auxotroph’s are mutants that cannot grow on a minimal medium and therefore require specific compounds to be added to the medium. Nitrate reductase deficient mutants of tobacco (N. tabacum) are known. The parental protoplasts of such species cannot grow with nitrate as the sole source of nitrogen while the hybrids can grow.

Two species of nitrate reductase deficiency— one due to lack of apoenzyme (nia-type mutant) and the other due to lack of molybdenum cofactor (cnx- type mutant) are known. The parental protoplasts cannot grow on nitrate medium while the hybrid protoplasts can grow (Fig.44.7).

 

The selection of auxotrophic mutants is possible only if the hybrid cells can grow on a minimal medium. Another limitation of the technique is the paucity of higher plant auxotroph’s.

Visual methods:

Visual selection of hybrid cells, although tedious is very efficient. In some of the somatic hybridization experiments, chloroplast deficient (albino or non-green) protoplasts of one parent are fused with green protoplasts of another parent.

This facilitates the visual identification of haterokaryons under light microscope. The heterokaryons are bigger and green in colour while the parental protoplasts are either small or colourless. Further identification of these heterokaryons has to be carried out to develop the specific hybrid plant. There are two approaches in this direction — growth on selection medium, and mechanical isolation.

1. Visual selection coupled with differential media growth:

There exist certain natural differences in the sensitivity of protoplasts to the nutrients of a given medium. Thus, some media can selectively support the development of hybrids but not the parental protoplasts. A diagrammatic representation of visual selection coupled with the growth of heterokaryons on a selection medium is given below.

 2. Mechanical isolation:

The visually identified heterokaryons under the microscope can be isolated by mechanical means. This involves the use of a special pipette namely Drummond pipette. The so isolated heterokaryons can be cloned to finally produce somatic hybrid plants. The major limitation of this method is that each type of hybrid cell requires a special culture medium for its growth. This can be overcome by employing micro drop culture of single cells using feeder layers.

Cytometric methods:

Some workers use flow cytometry and fluorescent-activated cell sorting techniques for the analysis of plant protoplasts while their viability is maintained. The same techniques can also be applied for sorting and selection of heterokaryons. The hybrid cells derived from such selections have proved useful for the development of certain somatic hybrid plants.

C. Identification of Hybrid (Cells) Plants:

The development of hybrid cells followed by the generation of hybrid plants requires a clear proof of genetic contribution from both the parental protoplasts. The hybridity must be established only from euploid and not from aneuploid hybrids. Some of the commonly used approaches for the identification of hybrid plants are briefly described.

Morphology of hybrid plants:

Morphological features of hybrid plants which usually are intermediate between two parents can be identified. For this purpose, the vegetative and floral characters are considered. These include leaf shape, leaf area, root morphology, flower shape, its structure, size and colour, and seed capsule morphology.

The somatic hybrids such as pomatoes and topatoes which are the fused products of potato and tomato show abnormal morphology, and thus can be identified. Although the genetic basis of the morphological characters has not been clearly known, intermediate morphological features suggest that the traits are under the control of multiple genes. It is preferable to support hybrid morphological characters with evidence of genetic data.

Isoenzyme analysis of hybrid plants:

The multiple forms of an enzyme catalysing the same reaction are referred to as isoenzymes. Electrophoretic patterns of isoenzymes have been widely used to verify hybridity. Somatic hybrids possess specific isoenzymes (of certain enzymes) of one or the other parent or both the parents simultaneously.

There are many enzymes possessing unique isoenzymes that can be used for the identification of somatic hybrids e.g. amylase, esterase, aspartate aminotransferase, phosphodiesterase, isoperoxidase, and hydrogenases (of alcohol, lactate, malate). If the enzyme is dimeric (having two subunits), somatic hybrids usually contain an isoenzyme with an intermediate mobility properties. The isoenzymes are often variable within the same plant. Therefore, it is necessary to use the same enzyme from each plant (parents and somatic hybrids), from a specific tissue with the same age.

Chromosomal constitution:

The number of chromosomes present in the hybrid cells can be directly counted. This provides information on the ploidy state of the cells. The somatic hybrids are expected to possess chromosomes that are equal to the total number of chromosomes originally present in the parental protoplasts. Sometimes, the hybrids are found to contain more chromosomes than the total of both the parents. The presence of chromosomal markers is greatly useful for the genetic analysis of hybrid cells.

Molecular techniques:

Many recent developments in molecular biology have improved the understanding of genetic constitution of somatic plant hybrids.

Some of them are listed below:

1. Differences in the restriction patterns of chloroplast and mitochondrial DNAs.
2. Molecular markers such as RFLP, AFLP, RAPD and microsatellites.
3. PCR technology.

Chromosome Number in Somatic Hybrids:

The chromosome number in the somatic hybrids is generally more than the total number of both of the parental protoplasts.

However, wide variations are reported which may be due to the following reasons:

1. Fusion of more than two protoplasts.
2. Irregularities in mitotic cell divisions.
3. In fusogen or electro-induced fusions, about one third of the fusions occur between morethan two protoplasts.
4. Differences in the status of protoplasts (actively dividing or quiescent) from the two species of plants result in formation of asymmetric hybrids.
5. Asymmetric hybrids may be due to unequal replication of DNA in the fusing protoplasts.
6. Protoplast isolation and culture may also lead to somaclonal variations, and thus variations in chromosome number.

A selected list of interspecific hybrids produced through protoplast fusion along with the number of chromosomes in the hybrids is given below

Symmetric and asymmetric hybrids:

If the chromosome number in the hybrid is the sum of the chromosomes of the two parental protoplasts, the hybrid is said to be symmetric. Symmetric hybrids between incompatible species are usually sterile. This may be due to production of 3n hybrids by fusing 2n of one species with n of another species.

Asymmetric hybrids have abnormal or wide variations in the chromosome number than the exact total of two species. These hybrids are usually formatted with full somatic complement of one parental species while all or nearly all of the chromosomes of other parental species are lost during mitotic divisions. Asymmetric hybrids may be regarded as cybrids but for the introgressed genes.

Cybrids:

The cytoplasmic hybrids where the nucleus is derived from only one parent and the cytoplasm is derived from both the parents are referred to as cybrids. The phenomenon of formation of cybrids is regarded as cybridization. Normally, cybrids are produced when protoplasts from two phytogenetically distinct species are fused. Genetically, cybrids are hybrids only for cytoplasmic traits.

Hybrids and Somatic Incompatibility:

Many a times, production of full-pledged hybrids through fusion of protoplasts of distantly related higher plant species is rather difficult due to instability of the two dissimilar genomes in a common cytoplasm. This phenomenon is referred to as somatic incompatibility. Hybrids formed despite somatic incompatibility may exhibit structural and developmental abnormalities. Several generations may be required to eliminate the undesirable genes. Due to this limitation in somatic hybridization, cybridization involving protoplast fusion for partial genome transfer is gaining importance in recent years.

Methodology of Cybridization:

A diagrammatic representation of the formation of hybrids and cybrids is given in below

As the formation of heterokaryon occurs during hybridization, the nuclei can be stimulated to segregate so that one protoplast contributes to the cytoplasm while the other contributes nucleus alone (or both nucleus and cytoplasm). In this way cybridization can be achieved.

Some of the approaches of cybridization are given here under:

1. The protoplasts of cytoplasm donor species are irradiated with X-rays or ? -rays. This treatment renders the protoplasts inactive and non-dividing, but they are efficient donors of cytoplasmic constituents when fused with recipient protoplasts.
2. Normal protoplasts can be directly fused with enucleated protoplasts. Enucleated protoplasts can be isolated by high-speed centrifugation.
3. Protoplasts are inactivated by metabolic inhibitors such as iodoacetate. In practice, iodoacetate treated protoplasts are fused with X-rays irradiated protoplasts for more efficient formation of cybrids.
4. It is possible to suppress nuclear division in some protoplasts and fuse them with normal protoplasts.

Genetic recombination in Asexual or Sterile Plants:

There are many plants that cannot reproduce sexually. Somatic hybridization is a novel approach through which two parental genomes of a sexual or sterile plants can be brought together. Thus, by fusing parental protoplasts, fertile diploids and polyploidy can be produced.

Overcoming Barriers of Sexual Incompatibility:

Sexual crossing between two different species (interspecific) and two different genus (inter-generic) is impossible by conventional breeding methods. Somatic hybridization overcomes the sexual incompatibility barriers.

Two examples are given hereunder:

1. Fusion between protoplasts of potato (Solanum tuberosum) and tomato (Lycopersicon esculentum) has created pomato (Solanopersicon, a new genus).
2. Interspecific fusion of four different species of rice (Oryza brachyantha, O. elchngeri, O. officinalis and O. perrieri) could be done to improve the crop.

A list of selected examples of somatic hybrids developed by interspecific protoplast fusion is given in the following table

A Novel Approach for Gene Transfer:

Somatic hybridization has made it possible to transfer several desirable genetic characters among the plants

Applications of Cybrids:

Cybridization is a wonderful technique wherein the desired cytoplasm can be transferred in a single step. Cybrids are important for the transfer of cytoplasmic male sterility (CMS), antibiotic and herbicide resistance in agriculturally useful plants. Some of the genetic traits in certain plants are cytoplasmically controlled. This includes some types of male sterility, resistance to certain antibiotics and herbicides.

Cybridization has been successfully used to transfer CMS in rice. Cybrids of Brassica raphanus that contain nucleus of B. napus, chloroplasts of atrazinc resistant B. campestris and male sterility from Raphanus sativas have been developed.

Applications of Somatic Hybridization:

Somatic hybridization has opened new possibilities for the in vitro genetic manipulation of plants to improve the crops.

Some of the practical applications are briefly given:

1. Disease resistance:

Several interspecific and inter-generic hybrids with disease resistance have been created. Many disease resistance genes (e.g., tobacco mosaic virus, potato virus X, club rot disease) could be successfully transferred from one species to another. For example, resistance has been introduced in tomato against diseases such as TMV, spotted wilt virus and insect pests.

2. Environmental tolerance:

The genes responsible for the tolerance of cold, frost and salt could be successfully introduced through somatic hybridization, e.g., introduction of cold tolerance gene in tomato.

3. Quality characters:

Somatic hybrids for the production of high nicotine content, and low erucic acid have been developed.

A selected list of genetic traits transferred through protoplast fusion in crop plant species

4. Cytoplasmic male sterility:

A modification of hybridization in the form of cybridization has made it possible to transfer cytoplasmic male sterility.

Other Application of Somatic Hybridization:

1. Somatic hybridization has helped to study the cytoplasmic genes and their functions. In fact, the information is successfully used in plant breeding programmes.

2. Protoplast fusion will help in the combination of mitochondria and chloroplasts to result in a unique nuclear-cytoplasmic genetic combination.

3. Somatic hybridization can be done in plants that are still in juvenile phase.

4. Protoplast transformation (with traits like nitrogen fixation by incorporating exogenous DNA) followed by somatic hybridization will yield innovative plants.

Limitations of Somatic Hybridization:

Although somatic hybridization is a novel approach in plant biotechnology, there are several problems and limitations.

The success of the technique largely depends on overcoming these limitations, some of

which are listed below:

1. Somatic, hybridization does not always produce plants that give fertile and visible seeds.
2. Regenerated plants obtained from somatic hybridization are often variable due to somaclonal variations, chromosomal elimination, organelle segregation etc.
3. Protoplast culture is frequently associated with genetic instability.
4. Protoplast fusion between different species/genus is easy, but the production of viable somatic hybrids is not possible in all instances.
5.  Some of the somatic hybrids, particularly when produced by the fusion of taxonomically different partners, are unbalanced and not viable.
6. There are limitations in the selection methods of hybrids, as many of them are not efficient.
7. There is no certainty as regards the expression of any specific character in somatic hybridization.
8. Somatic hybridization between two diploids results in the formation of an amphidiploid which is not favourable. For this reason, haploid protoplasts are recommended in somatic hybridization.