# SBT 425: CELL AND TISSUE CULTURE

## Course outline

### course out line

1. History and Development of tissue culture
2. Tissue culture equipment and facilities
3. Sterilization techniques
4. Tissue culture media

## lesson 1

### Introduction

Definition of tissue culture Refers to a technique that allows a whole plant to be produced from minute amount of the plant part (explant e.g immature embryos, shoot tip, root, leaf, ovary ovary, scutellum) or just a single cell on a sterile nutrient medium under laboratory controlled conditions (in vitro) often in small glass or plastic containers.

### Terminologies in tissue culture

• Aseptic---Free of microorganisms.
• Aseptic Technique---Procedures used to prevent the introduction of fungi, bacteria, viruses, *mycoplasma or other microorganisms into cultures.
• Autoclave---A machine capable of sterilizing wet or dry items with steam under pressure. Pressure *cookers are a type of autoclaves.
• Callus---An unorganized, proliferating mass of undifferentiated plant cells or tissues a under the influence of elevatedplant hormone levels.
• Chemically Defined Medium---A nutritive solution for culturing cells in which each component is specifiable and ideally of known chemical structure.
• Clone---Plants produced asexually from a single source plant.
• Clonal Propagation---Asexual reproduction of plants that are considered to be genetically uniform and originated from a single individual or explant.
• Contamination---Being infested with unwanted microorganisms such as bacteria, fungi or viruses
• Culture—A plant growing in vitro.
• Differentiated---Cells that maintain, in culture, all or much of the specialized structure and function typical of the cell type in vivo. Modifications of new cells to form tissues or organs with a specific function.
• Explant---Tissue taken from its original site and transferred to an artificial medium for growth or maintenance. Part of an organism used in "in vitro" culture
• Totipotency - The establishment of missing plant organs or parts; formation of a whole

plant from a few cells or small portion of a plant.

### Basis for the cell culture

In cell and tissue culture the concept of totipotency is used. Plant cells are totipotent if they have the ability to develop into whole plants or plant organs in vitro when given the correct conditions which is a characteristic potrayed by a zygote. Plant tissue culture relies on the fact that many plant cells have the ability to regenerate a whole plant (totipotency). Single cells, plant cells without cell wall(protoplasts), pieces of leaves, or (less commonly) roots can often be used to regenerate a new plant on culture media given the required nutrients and plant hormones. Not all plant cells are totipotent. However, there are a sufficient number of totipotent cells in the plant. Differentiated cells have to be dedifferentiated into callus and redifferentiated back to somatic embryo that will regenerate the entire plant.

Thus totipotency implies that undifferentiated plant cells (meristematic cells) have the ability to display their full genetic potential to form functional plants when cultured in vitro. The major impact of plant tissue culture will be felt in the area of controlled manipulations of plants at the cellular level in ways which have not been possible prior to the introduction of tissue culture.

ITissue culture has applications in research and commerce. In commercial settings, tissue culture is primarily used for plant propagation and it is often referred to as micropropagation. Micropropagation refers to the production of whole plants from cell cultures derived from explants (the initial piece of tissue put into culture); the explants usually consist of tissues that contain or develop into meristem cells.

Plant tissue culture techniques are essential to many types of academic inquiry, as well as to many applied aspects of plant science. In the past, plant tissue culture techniques have been used in academic investigations of totipotency and the roles of hormones in cytodifferentiation and organogenesis. Currently, tissue-cultured plants that have been genetically engineered provide insight into plant molecular biology and gene regulation. Plant tissue culture techniques are also central to innovative areas of applied plant science, including plant biotechnology and agriculture.

For example, selected plants can be cloned and cultured as suspended cells from which plant products can be harvested. In addition, the management of genetically engineered cells to form transgenic whole plants requires tissue culture procedures; tissue culture methods are also required in the formation of somatic haploid embryos from which homozygous plants can be generated. Thus, tissue culture techniques have been, and still are, prominent in academic and applied plant science. The techniques demonstrated in these exercises range from simple ones that can easily be performed by beginning students to those done by botany or physiology students. Experiment 1 and 2 employ plant material derived from aseptic seed germinations, while Experiments 3, 4, and 5 use portions of large intact plants. Experiment 1 demonstrates "in vitro" morphogenesis and totipotency and has been used successfully by beginning classes containing both biology majors and non-majors The remaining experiments are designed for use by more advanced students.

. History and advancement in Development of Tissue culture The history of plant tissue culture can be traced back to near the turn of the 20th century when Gottlieb Haberlandt (1902) reported isolated single palisade cells from leaves/leaf mesophyll tissue cultured in Knop's salt solution enriched with sucrose. But failed to divide. Gottlieb Haberlandt a German botanist was the first to generate tissue from fully differentiated tissue. The first embryo culture, although crude, was done by Hanning in 1904; he cultured nearly mature embryos of certain crucifers and grew them to maturity. The technique was utilised by Laibach in 1925 to recover hybrid progeny from an interspecific cross in Linum. Subsequently, contributions from several workers led to the refinement of this technigue.

White (1934) repeatedly reported continuously growing culture of meristematic cells of tomato from root tip-derived tissues of on medium containing inorganic salts, yeast extract and sucrose and 3 vit B (pyridoxine, thiamine, nicotinic acid) – established the importance of additives.


Techniques for plant tissue culture progressed rapidly during the 1930s due to the discovery of the necessity of B vitamins and auxin for the growth of isolated meristem tissues. Some notable major discoveries included chemical and hormonal control of regeneration by Skoog and their associates on the nutritional requirements of tobacco tissue culture led to not only the discovery of plant growth hormones, kinetin and auxins, but also to the formation of an important plant tissue culture medium, the MS medium (Skoog and Tsui, 1951; Skoog and Miller, 1957; Murashige and Skoog, 1962). Haploid plants from pollen grains were first produced by Maheshwari and Guha in 1964 by culturing anthers of Datura. This marked the beginning of anther culture or pollen culture for the production of haploid plants. The technique was further developed by many workers, more notably by JP. Nitch (1967), C. Nitch and coworkers. These workers showed that isolated microspores of tobacco produce complete plants. Murashige cloned plants in vitro, raised haploid plants from pollen grains and used protoplast fusion to hybridize 2 species of tobacco into one plant contained 4N. Since 1960s, tissue and cell culture has increasingly been used as a tool by plant scientists and biotechnologists. Practical micropropagation and production of virus-free plants (G. Morel, 1960), haploid plants (S. Guha and S.C. Maheshwari, 1964, J. P. Nitsch, 1967), culture and regeneration of protoplasts (E.C. Cocking, 1960), production of secondary metabolites (B. Kaul and E.J. Staba, 1967) and large-scale cell culture in bioreactors (M. Noguchi et al., 1977), to mention just a few landmarks. 1970’s and 1980s marked the beginning of genetic engineering.. The ongoing achievements in in vitro culture of pollen, protoplasts and cell suspensions – and their ability to regenerate whole plants has resulted into a new disciplines of plant science: somatic cell genetics and metabolite production, somatic hybrids, production of haploid plants, selection of variants and mutants, and improved generation of metabolites. All became available for the community of plant scientists and breeders, expanding the diversity, agronomic and commercial value of plants. Yet, much remains to be explored in terms of methodology, procedures and the theories behind the technology

Tissue culture equipment and facilities


Tissue culture equipment and supplies Laminar flow hoods. There are two types of laminar flow hoods, vertical and horizontal. The vertical hood, also known as a biology safety cabinet, is best for working with hazardous organisms since the aerosols that are generated in the hood are filtered out before they are released into the surrounding environment. Horizontal hoods are designed such that the air flows directly at the operator hence they are not useful for working with hazardous organisms but are the best protection for your cultures. Both types of hoods have continuous displacement of air that passes through a HEPA (high efficiency particle) filter that removes particulates from the air. In a vertical hood, the filtered air blows down from the top of the cabinet; in a horizontal hood, the filtered air blows out at the operator in a horizontal fashion.. The hoods are equipped with a short-wave UV light that can be turned on for a few minutes to sterilize the surfaces of the hood, but be aware that only exposed surfaces will be accessible to the UV light. Do not put your hands or face near the hood when the UV light is on as the short wave light can cause skin and eye damage. The hood is used for the transfer of explants and culture, dispensing sterile media.

b. Microscopes. Inverted phase contrast microscopes are used for visualizing the cells. Microscopes should be kept covered and the lights turned down when not in use. Before using the microscope or whenever an objective is changed, check that the phase rings are aligned. c. Vessels. Anchorage dependent cells require a nontoxic, biologically inert, and optically transparent surface that will allow cells to attach and allow movement for growth. The most convenient vessels are specially-treated polystyrene plastic that are supplied sterile and are disposable. These include petri dishes, multi-well plates, microtiter plates, roller bottles, and screwcap flasks. Autoclave/pressure cooker – for sterilizing the media and the vessels Refrigirator – for storing chemicals at low temperature (4oC) such as vitamins, growth regulators and micronutrients Cultures bottles/baby jars/petridishes – for putting a medium used for culturing, transfers of cultures Incubator – used for growth of culture at a specific controlled temperature Freezer – storage of chemicals at -20oC Shaker – for the growth of the suspension cultures Balances – for weighing chemicals used for preparing stock solutions and culture media pH meter – for measuring the pH of the media Thermometer – for measuring temperature in the growth room Dry Sterilization oven – for sterilizing vessels and instruments Microva – for heating agar media Bunsen burner Beakers, Pipettes, micropipettes, volumetric flasks, graduated measuring cylinders

–for measuring stock solutions/chemicals and the media


Parafilm – for sealing petrishes Magnetic stirrer – used when preparing the media Metal trays – for carrying the media, stock solutions and tissue culture tools Aluminium foil – for wrapping instruments and vessel opening when sterilizing Scapels, forceps – for cutting and transferring explants and cultures Hot plate – for melting and dissolving the gelling agents Growth chamber – for growing culture or plants under controlled conditions of light and temperature.

Facilities Culture Room – room where all activities of sterile transfers of explants and cultures are done Media preparation room –Room where the preparation of the culture media is done. Growth room – Room with controlled temperature and light where cultures are incubated or grown Green house – where in vitro regenerated plants are hardened and grown to maturity

## lesson 2

Sterilization (Aseptic) Techniques Used in Tissue Culture Tissue culture requires sterile conditions. The culture room, lamina hood, instruments, vessels, culture media, plant materials used must be sterile. Aim the essence of aseptic technique is to ensure all cell culture procedures are performed to a standard that will prevent contamination from bacteria,viruses and fungi and cross contamination with other cell lines from culture to culture If sterile tissues are available, then the exclusion of microorganisms is accomplished by using sterile instruments and culture media concurrently with standard bacteriological transfer procedures to avoid extraneous contamination.

Aseptic technique is absolutely necessary for the successful establishment and maintenance of plant cell, tissue and organ cultures. The in vitro environment in which the plant material is grown is also ideal for the proliferation of microorganisms. In most cases the microorganisms outgrow the plant tissues,.

The environmental control of air is also of concern because room air may be highly contaminated. Example: Sneezing produces 100,000 - 200,000 aerosol droplets which can then attach to dust particles.

Successful control of contamination depends largely upon the operator’s techniques in aseptic culture. One should always be aware of potential sources of contamination such as dust, hair, hands, and clothes All the materials, e.g., vessels, instruments, medium, plant material, etc., used in culture work must be freed from microbes. This is achieved by one of the following approaches: (i) dry heat treatment, (ii) flame sterilization, (iii) autoclaving, (iv) filter sterilization, (v) wiping with 70% ethanol, and (vi) surface sterilization.

### Sterilization of the culture room

The room should be sterilized with 70% ethanol or methylated spirit 20minutes before culturing.

### Sterilization of the lamina hood

UV Radiation: It is possible to use germicidal lamps to sterilize items in the transfer hood when one is working in the hood. UV lamps should not be used when people are present because the light is damaging to eyes and skin. Plants left under UV lamps will die. Ethanol: The hood should be sterilized with 70% ethanol or methylated spirit 20 minutes before culturing. 70% ethanol is used

### How to mantain sterilization when working in the Transfer Hood/lamina hood:

1. Spray or wipe the inside of the transfer hood using 70% ethanol let the spray dry before commencing work . Wipe up any spills quickly; use 70% EtOH for cleaning. Clean hood surface periodically while working
2. The hood should remain on continuously. If for some reason it has been turned off, turn it on and let it run for at least 15 minutes before using.
3. Sterilize gloves by washing them in 70% ethanol and allowing to air dry for 30 seconds before commencing work.
4. Remove watches, etc., roll up long sleeves, and wash hands thoroughly with soap (preferably bactericidal) and water then Wipe hands and lower arms with 70% EtOH
5. Spray equipment put into the sterile area with 70% ethanol. For example, spray bags of petri dishes, media bottles, pipette aids with 70 % alcohol before you open them and place the desired number of unopened dishes in the sterile area.

 Make sure that everything needed for the work is in the hood and all unnecessary things are removed. Keep as little in the hood as possible. As few things as possible should be stored in the hood. Make sure that materials in use are to the side of your work area, so that airflow from the hood is not blocked. Arrange tools and other items in the hood so that your hands do not have to cross over each other while working. All other items in the hood should be arranged so that your work area is directly in front of you, and between 8 and 10 inches in from the front edge. No materials should be placed between the actual work area and the filter.  Sterilize culture tubes with lids or caps on. When you open a sterile tube, touch only the outside of the cap, and do not set the cap on any laboratory surface. Instead, hold the cap with one or two fingers while you complete the operation, and then replace it on the tube. This technique usually requires some practice, especially if you are simultaneously opening tubes and operating a sterile pipette. After you remove the cap from the test tube, pass the mouth of the tube through a flame. If possible, hold the open tube at an angle. Put only sterile objects into the tube. Complete the operation as quickly as you reasonably can, and then flame the mouth of the tube again. Replace the lid. If you don't have to be careful about the volume you transfer, a pure sterile solution can be transferred to a sterile container or new sterile medium by pouring. For example, we do not measure a specific volume of medium when we pour culture plates, although after you have done it for a while, you become pretty consistent. Remove the cap or lid from the solution to be transferred. Thoroughly flame the mouth of the container, holding it at an angle as you do so. Remove the lid from the target container. Hold the container at an angle. Quickly and neatly pour the contents from the first container into the second. Replace the lid.  If you must transfer an exact volume of liquid, use a sterile pipette or a sterile graduated cylinder. When using a sterile graduated cylinder, complete the transfer as quickly as you reasonably can to minimize the time the sterile liquid is exposed to the air.

1. Don’t touch any surface that is supposed to remain sterile with your hands. Use forceps, etc.
2. Whilst working do not contaminate gloves by touching anything outside the cabinet (especially face and hair). If gloves become contaminated re-sterilized with 70% ethanol as above before proceeding.
3. Plant material should be placed on a sterile surface when manipulating it in the hood. Sterile petri dishes (expensive), sterile paper towels, or sterile paper plates work fine. Pre-sterilized plastic dishes have two sterile surfaces-the inside top and inside bottom.
4. Plastic pipettes are purchased presterilized in individual wrappers. To use a pipette, remove it from its wrapper or container by the end opposite the tip. Do not touch the lower two-thirds of the pipette. Do not allow the pipette to touch any laboratory surface. Insert only the untouched lower portion of the pipette into a sterile container.
5. Discard gloves after handling contaminated cultures and at the end of all cell culture procedures.
6. Know which of your implements, flasks, etc. are sterile and which are not
7. Sterilized items should be used within a short time
8. Movement within and immediately outside the cabinet must not be rapid. Movements in the hood should be limited to a small area. Slow movement will allow the air within the cabinet to circulate properly. Work well back in the transfer hood (behind the line). Especially keep all flasks as far back to the back of the hood as possible.
9. Speech, sneezing and coughing must be directed away from the cabinet so as not to disrupt the airflow.
10. Remove items from the hood as soon as they are no longer needed. All cultures must be sealed before leaving the hood.
11. After completing work disinfect all equipment and material before removing from the cabinet. Spray the work surfaces inside the cabinet with 70% ethanol and wipe dry with tissue. Dispose of tissue by autoclaving.
12. It is pointless to practice good sterile technique in a dirty lab. Special problems are contaminated cultures, dirty dishes and solutions where microorganisms can grow.

Sterilization technique used include: • Ethanol: Instruments (scalpels, forceps) can be sterilized dipping them in 70% or 95% EtOH and then immediately placing them in the flame of an alcohol lamp or gas burner. This can be dangerous if the vessel holding the alcohol tips over and an alcohol fire results. A fairly deep container, like a coplin-staining jar, should be used to hold the ethanol. Use enough ethanol to submerge the business ends of the instruments but not so much that you burn your hands. Some people wear gloves in the hood for certain procedures. If you do this, be very careful not to get them near the flame. • Microwave: Instrument are rubbed using aluminum foil and then sterilized. • Dry heat: Empty glassware (culture vessels, pipettes, etc.), and certain Plasticware (Teflon FEP); instruments like scalpels, forceps, needles, etc. can be sterilized using an oven at 160-180oC for 4 hours.. More recently, glass bead sterilizers (300°C) are being employed for the sterilization these devices use dry heat. Small instruments such as scalpels and forceps are placed into the glass beads and are sterilized within 10-60 seconds. The instruments will cool down to working temperatures with 30-60 seconds. These sterilizers heat to approximately 275-350° C and will destroy bacterial and fungalspores that may be found on your instruments. The instruments simply need to be inserted into the heated glass beads for a period of 10 to 60 sec. The instruments should then be placed on a rack under the hood to cool until needed. These units kill most major classes of fungi, bacteria, and viruses. • Autoclaving: Empty vessels, beakers, graduated cylinders, etc., should be closed with a cap or aluminum foil. Tools should also be wrapped in foil or paper or put in a covered sterilization tray. It is critical that the steam penetrate the items in order for sterilization to be successful. However autoclaving is not advisable for metal instruments because they may rust and become blunt under these conditions. • Flame sterilization: Instruments like scalpels, forceps, etc that have been sterilized in hot dry air should be removed from their wrapping, dipped in 95% ethyl alcohol, and exposed to the heat of a flame. After an instrument has been used, it can again be dipped in ethyl alcohol, re-flamed, and then reused. This technique is called flame sterilization . Flaming instruments prior to use and flaming the opening of receiving vessels prior to transfer is reqiured. Aseptic transfers are more easily performed in a transfer chamber such as a laminar flow hood, which is also preferably equipped with a bunsen burner. • NOTE: Items that come packaged sterile e.g Petri dishes should be examined carefully for damage before use. Surface-sterilizing Plant Materials Plants materials used in tissue culture need to be healthy and actively growing. Stressed plants, particularly water-stressed plants, usually do not grow as tissue cultures. Insect and disease-free greenhouse plants are rendered aseptic more readily, so contamination rate is lower when these plants are used in tissue culture procedures. Seeds that can be easily surface sterilized usually produce contamination-free plants that can be grown under clean greenhouse conditions for later experimental use

If experimental tissues are not aseptic, then surface sterilization procedures specific to the tissues are employed. Common sterilants are ethyl alcohol and/or chlorox Bleach, Calcium hypochlorite, Mercuric chloride, Hydrogen peroxide with an added surfactant. Concentration of sterilants and exposure time are determined empirically.

Steps in sterilization of plant materials (explants) 1. Preparation of Stock Plants Prior good care of stock plants may lessen the amount of contamination that is present on explants. Plants grown in the field are typically more “dirty” than those grown in a greenhouse or growth chamber. Overhead watering increases contamination of initial explants. Likewise, splashing soil on the plant during watering will increase initial contamination. Treatment of stock plants with fungicides and/or bacteriocides is sometimes helpful. It is sometimes possible to harvest shoots and force buds from them in clean conditions. The shoots may be free of contaminants when surface-sterilized in a normal manner. Seeds may be sterilized and germinated in vitro to provide clean material. Covering growing shoots for several days or weeks prior to harvesting tissue for culture may supply cleaner material. Explants or material from which material will be cut can be washed in soapy water and then placed under running water for 1 to 2 hours.

2. Sterilization of explants sterilization using sterilants a) Ethanol Ethanol is a powerful sterilizing agent but also extremely phytotoxic. Therefore, plant material is typically exposed to it for only seconds or minutes depending on the sensitivity of the explants and how difficult it is to disinfect. Explants from Woody and field plants takes longer time. The more tender the tissue, the more it will be damaged by alcohol. Tissues such as dormant buds, seeds, or unopened flower buds can be treated for longer periods of time since the tissue that will be explanted or that will develop is actually within the structure that is being surface-sterilized. Generally 70% ethanol is used prior to treatment with other compounds.

3. Sterilization using bleaching agents a. Sodium Hypochlorite Sodium hypochlorite, usually purchased as laundry bleach, is the most frequent choice for surface sterilization. It is readily available and can be diluted to proper concentrations. Commercial laundry bleach is 5.25% sodium hypochlorite. It is usually diluted to 10% - 20% of the original concentration, resulting in a final concentration of 0.5 - 1.0% sodium hypchlorite. Plant material is usually immersed in this solution for 10 - 20 minutes. A balance between concentration and time must be determined empirically for each type of explant, because of phytotoxicity.

b. Calcium Hypochlorite The concentration of calcium hypochlorite used is 3.25 %. The solution must be filtered prior to use since not the entire compound goes into solution. Calcium hypochlorite may be less injurious to plant tissues than sodium hypochlorite.

c. Mercuric Chloride Mercuric chloride is used only as a last resort. Concentration of 0.2% is used. It is extremely toxic to both plants and humans and must be disposed of with care. Since mercury is so phytotoxic, it is critical that many rinses be used to remove all traces of the mineral from the plant material.

d. Hydrogen Peroxide The concentration of 10 % hydrogen peroxide is used for surface sterilization of plant material. Some researchers have found that hydrogen peroxide is useful for surface-sterilizing material while in the field.

Enhancing Effectiveness of Sterilization Procedure • Surfactant/wetting agent (e.g.Tween 20) is frequently added to the sodium hypochlorite. • The solutions that the explants are in are often shaken or continuously stirred.

4. Rinsing with sterile distilled water After plant material is sterilized with one of the above compounds, it must be rinsed thoroughly with sterile water. Typically three to five separate rinses are done to eliminate the residue of the disinfectant.

Media sterilization Two methods (autoclaving and membrane filtration under positive pressure) are commonly used to sterilize culture media. Autoclaving: Autoclaving is the method most often used for sterilizing culture media In order to be sterilized, the culture media must be held at 121C, 15 psi, (pounds per square inch; 1.06 kg/cm2). It is important that this temperature is attained before timing begins. Therefore time in the autoclave will vary, depending on volume in individual vessels and number of vessels in the autoclave. Most autoclaves automatically adjust time when temperature and psi are set, and include time in the cycle for a slow decrease in pressure. There are tape indicators that can be affixed to vessels, but they may not reflect the temperature of liquid within them.. Culture media, distilled water, and other heat stable mixtures can be autoclaved in glass containers that are sealed with cotton plugs, aluminum foil, or plastic closures. For small volumes of liquids (100 ml or less), the time required for autoclaving is 15-20 min, larger quantities (2-4 liter), 30-40 min is required to complete the cycle. The pressure should not exceed 20 psi, as higher pressures may lead to the decomposition of carbohydrates and other components of a medium. Too high temperatures or too long cycles can also result in changes in properties of the medium. ABA, IAA, IBA, kinetin, pyridoxine, 2-ip and thiamine are usually autoclaved.

A summary of the time required for the of various volumes of the medium at 121oC Volume of the medium per vessel Pre-heating time to reach 121oC Total sterilization time 20-25 9 24 50 11 26 100 13.5 28.5 250 16.5 31.5 500 20 35 1000 25 40 2000 33 48 3000 40 55 4000 48 63

Membrane filtration: Solutions that contain heat-labile components are best sterilized by ultrafiltration. However the process is slower and more costly than autoclaving. Organic compounds such as some growth regulators such as heat labile compounds like GA3, ABA, zeatin, enzymes, amino acids, and vitamins may be degraded during autoclaving. These compounds require filter sterilization through a 0.22 µm membrane. Several manufacturers make nitrocellulose membranes that can be sterilized by autoclaving. They are placed between sections of a filter unit and sterilized as one piece. Other filters (the kind we use) come pre-sterilized. Larger ones can be set over a sterile flask and a vacuum is applied to pull the compound dissolved in liquid through the membrane and into the sterile flask. Smaller membranes fit on the end of a sterile syringe and liquid is pushed through by depressing the top of the syringe. The size of the filter selected depends on the volume of the solution to be sterilized and the components of the solution.

# -=Tissue culture media==

Can be a solid or a liquid culture media There are different types of tissue culture media such as  MS (developed for tobacco),  LS (Linsmaier and Skoogs medium),  Gamborg B5 (developed for soyabean cell suspension cultures),  Schenk and Hildebrandt (developed for callus cultures of monocots and dicots),  N6 (developed for anther culture of rice),  Whites medium (developed for tissue culture of tomato roots),  Wood medium (for trees) and  Nitschs medium for anther culture).

Ms and B5 are the most commonly used media.Virtually all tissue culture media are synthetic or chemically defined; only a few of them use complex organics, e.g., potato extract, as their normal constituents. A synthetic medium consists of only chemically defined compounds. A variety of recipes have been developed since none of them is suitable for either all plant species or for every purpose. Composition of Some Plant Tissue Culture Media - Components of media for the growth of plant callus and suspension cultures can be classified into five groups. The division usually reflects the way in which stock solutions are prepared and stored. These groups are: i) inorganic nutrients (macronutrients and micronutrients)

ii) organic supplement (vitamins)


iii) carbon source

iv) Growth regulators
`

v) gelling agents Media compositions are formulated considering specific requirements of a particular culture system or species. 'For example, some tissues show better response on a solid medium while others prefer a liquid medium or others do better in MS medium than in B5 medium.. Some tissues are grown on simple media containing only inorganic salts and utilizable carbon (sugar) source, but for most others it is essential to supplement the medium with vitamins, amino acids and growth substances. Very often, complex nutritive substances are added to the medium. Such a medium, composed of "chemically defined" compounds, is referred to as a "synthetic" medium. 1. Inorganic nutrients a) Macronutrients – Required in large quantities. The macronutrients include six major elements-nitrogen (N), phosphorus (P), potassium (K), calcium (Ca), magnesium (Mg), and surphur (S)-present as salts that constitute various media. MgSO4 provide Mg & S; KH2PO4 and NaHPO4 provides P; CaCl2.2H2O or Ca(NO3)2. 4H2Oprovide calcium and KCl, KNO3 KH2PO4 provides K; KCl Or CaCl2.2H2O provide chloride. All are essential for plant cell and tissue growth. Culture media should contain at least 25 mM nitrate and potassium. However, considerably better results are obtained if the source for nitrogen in media is contributed by both nitrates and ammonium (2-20 mM) or any other reduced nitrogen source. In case only ammonium is used, there is need to add one or more tricarboxylic acid (TCA) cycle acids (e.g., citrate, succinate, or malate) so that any deleterious effect due to ammonium concentrations in excess of 8 mM in the medium is diluted. When nitrate and ammonia ions are present together in the culture medium, the latter are used more rapidly. b) Micronutrients Are required in lower concentrations. Include boron (H2BO3); Cobalt (COCl2); iron (NaFeEDTA); managanase (MnSO4.H2O)); molybdenum (NaMoO4); copper (CuSO4. H2O) and Zinc (ZnSO4.7H20).