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Tissue Culture
Tissue Culture
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Plant tissue culture

  1. 1. INTRODUCTION TO PLANT TISSUE CULTURE:  It is the process of producing plants from tissue of the desired plant in an artificial nutrient medium under controlled environment.  The plants so grown would be exactly similar to the mother plant in all aspects.  The science of plant tissue culture takes its roots from path breaking research in botany like discovery of cell followed by propounding of cell theory. In 1839, Schleiden and Schwann proposed that cell is the basic unit of organisms. They visualized that cell is capable of autonomy and therefore it should be possible for each cell if given an environment to regenerate into whole plant. Based on this premise, in 1902, a German physiologist, Gottlieb Haberlandt developed the concept of in vitro cell culture. He isolated single fully differentiated individual plant cells from different plant species like palisade cells from leaves of Laminum purpureum, glandular hair of Pulmonaria and pith cells from petioles of Eicchornia crassiples etc and was first to culture them in Knop’s salt solution enriched with glucose. In his cultures, cells increased in size, accumulated starch but failed to divide. Therefore, Haberlandt’s prediction failed that the cultured plant cells could grow, divide and develop into embryo and then to whole plant. This potential of a cell is known as totipotency, a term coined by Steward in 1968.
  2. 2. STERILIZATION TECHNIQUES:  Sterilization Methods Used in Tissue Culture Laboratory - All the materials, e.g., vessels, instruments, medium, plant material, etc., used in culture work must be freed from microbes. This is achieved by one of the following approaches: (i) dry heat treatment, (ii) flame sterilization, (iii) autoclaving, (iv) filter sterilization, (v) wiping with 70% ethanol, and (vi) surface sterilization.
  3. 3. EXPERIMENTS: Protocol.1- Tissue Culture Media Preparation PRINCIPLE: Murashige and Skoog medium or (MSO or MS0 (MS-zero)) is a plant growth medium used in the laboratories for cultivation of plant cell culture. MSO was invented by plant scientists Toshio Murashige and Folke K. Skoog during Murashige's search for a new plant growth regulator. It is the most commonly used medium in plant tissue culture experiments. A series of experiments demonstrated that varying the levels of these nutrients enhanced growth substantially over existing formulations. It was determined that nitrogen in particular enhanced growth of tobacco in tissue culture.
  4. 4. Protocol-2- Explant Preparation and Surface Sterilization PRINCIPLE: Surface sterilization treatments applied on the explants were: Dipping in Ethyl alcohol 70% for 5 minutes (mins). (T1) Dipping in Chlorox 3% (commercial sodium hypochlorite, active ingredients 5.2%) for 20 mins. (plus Tween 20). (T2) Dipping in undiluted Chlorox for 20 mins. (plus Tween 20). (T3) Dipping in Ethyl alcohol 70% for 5 mins. then in Chlorox 3% (plus Tween 20) for 20 mins. (T4) dipping in Ethyl alcohol 70% for 5 mins. then in Mercuric chloride 0.3% for 5 mins. (T5) After each treatment explants were rinsed three times in autoclaved distilled water.
  5. 5. Protocol-3-EMBRYO CULTURE: PRINCIPLE: Organogenesis is the formation of individual organs such as shoots and roots either directly on the explant in which pre-formed meristem are lacking or the meristems develop de novo from callus. Although callus is an actively growing undifferentiated mass of cells, differentiation can take place at random, but may be associated with centres of morphogenesis, which can give rise to organs such as shoots, roots and embryos. To induce shoot organogenesis from the callus, concentration of growth regulators is varied in the medium. Cytokinins such as BAP and kinetin promote shoot organogenesis. RESULT:
  6. 6. Protocol-4- Culture of Anther for Production of Androgenic Haploids PRINCIPLE: Anther and Microscope Culture - One of the very popular methods for production of haploids is through culturing anthers or microspores on artificial culture medium. This leads to the growth of microspores into saprophytes. After the initial reports of successful production of haploids from anther culture in Datura (Guha and Maheshwari, 1966, 1967), haploids have been obtained in more than 150 species belonging to 23 families of angiosperms (Maheshwari et a1., 1980). These include a wide variety of economically important species. More often, anthers rather than microspores are cultured, since the extraction and culture methods for microspores differ and have been successful only in a few species (Datura inoxia, Nicotiana sylvestris, N. tabacum, Oryza sativa, etc.).
  7. 7. Protocol-5-Meristem culture PRINCIPLE: Organogenesis is the formation of individual organs such as shoots and roots either directly on the explant in which pre-formed meristem are lacking or the meristems develop de novo from callus. Although callus is an actively growing undifferentiated mass of cells, differentiation can take place at random, but may be associated with centres of morphogenesis, which can give rise to organs such as shoots, roots and embryos. To induce shoot organogenesis from the callus, concentration of growth regulators is varied in the medium. Cytokinins such as BAP and kinetin promote shoot organogenesis.
  8. 8. Protocol-6- Meristem tip culture for production of Virus –free Plants  Theme: Shoot apical meristem lies in the 'shoot tip' beyond the youngest leaf or first leaf primordium ; it measures upto about 100 µm in diameter and 250 µm in length. Thus a shoot-tip of 100-500 11m would contain 1-3 leaf primordia in addition to the apical meristem.In practice, shoot-tips of up to 1 mm are used when the objective is virus elimination. Shoot-tip culture is widely used for rapid clonal propagation for which much larger, e.g., 5-10 mm, explants are used. Therefore, most cases of meristem culture are essentially shoot-tip cultures. Nodal explants of various sizes are also commonly employed for rapid clonal propagation. When the objective is vegetative propagation, the size of shoot-tip used for culture is not important. The upper few millimeters (ca.5-6mm) in a shoot apex is considered to be free from virus in those plants which are systematically infected. Due to active cell division (faster than virus multiplication), absence of vascular connection and high auxin concentration the shoot meristem remains virus free. If the meristem tip is used as an explant for propagation, virus free plats can be obtained.
  9. 9. Protocol-7- Induction of Somatic Embryogenesis (Monocot and Dicot System)  In somatic embryogenesis the embryo arises from somatic cells, tissue or organs under in vitro conditions. Somatic embryo is a bipolar structure and has no vascular connection with the maternal cultured explant. The somatic embryos are functionally equivalent to zygotic embryos but the process of embryogeny is different. It induces four developmental phases i.e.0, 1, 2, and 3 phases. Phase 0: In phase 0 the competent single cells (state 0 cells) form embryogenic cell clusters (state 1 cells) in the presence of auxin. During this phase the cell cluster formed from single cells gains the ability to develop into embryos when auxin (a PGR used to induce somatic embryogenesis) is removed from the medium giving rise to state 1 clusters.  Phase 1: Phase 1 is induced by transfer of state 1 cell clusters on to auxin free medium. In this phase the cell cluster proliferate slowly and undifferentiately.  Phase 2: In phase 2, rapid cell division occurs in certain parts of cell clusters leading to formation of globular embryos. Phase 3: In phase 3, globular embryos develop into plantlets via heart shaped, torpedo shaped and cotyledonary stage embryos. Somatic embryos could be induced either directly from the explant tissue in the absence of callus formation (direct somatic embryogenesis) or via the callus from the explant (Indirect somatic embryogenesis). Embryogenic cells are small, isodiametric in shape, filled with dense cytoplasm and have a conspicuous nucleus. In comparison to this, on-embryogenic cells are relatively large, vacuolated and lack dense cytoplasm.
  10. 10. Protocol-8- Protoplast Isolation, Culture and Regeneration PRINCIPLE: Mechanical Method of Isolation of Protoplast - In mechanical method, cells are kept in a suitable plasmolyticum (in plasmolysed cells, protoplasts shrink away from cell wall) and cut with a fine knife, so that protoplasts are released from cells cut through the cell wall, when the tissue is again deplasmolysed. This method is suitable for isolation of protoplasts from vacuolated cells (e.g. onion bulbs, scales, radish roots). However, this method gives poor yield of protoplasts and is not suitable for isolating protoplast from meristematic and less vacuolated cells. The mechanical method, though, was used as early as 1892, is now only rarely used for isolation of protoplasts.
  11. 11. Protocol-9- Microculture Chamber Technique for Single Cell Isolation PRINCIPLE:  Single Cell Culture - Establishment of a single cell culture provides an excellent opportunity to investigate the properties and potentialities of plant cells. Such studies contribute to our understanding of the interrelationships and complementary influences of cells in multicellular organisms. Several workers have successfully isolated single cell division and even raised complete plants from single cell cultures. Using cell cultures in studies designed to describe the pathways of cellular metabolism was another aspect that initially attracted the attention of plant biologists. It was soon realized that single cell systems have great potential for crop improvement.  The microculture chamber technique was first developed by Jones et al. (1960) and later was used by Vasil and Hildebrandt (1965) after some modifications. This method consists of culturing 30-50μ/ of medium containing one or more protoplasts on a microscope slide enclosed by a cover glass resting on two other cover glasses placed on either side of the drop. The cultures are sealed with sterile paraffin oil and incubated in light at 23-25º C.
  12. 12. Protocol-10- Encapsulation of somatic embryos / shoot buds for Production of Synthetic seeds PRINCIPLE: Synthetic seeds are prepared by encapsulating the somatic embryos obtained from tissue culture in a protective jelly capsule, which is usually prepared with sodium alginate. From the synthetic seeds whole plant can be recovered under in vitro, greenhouse and field conditions. Alternatively, shoot buds, conservation and maintenance of rare and threatened species. Synthetic Seeds - In the conventional plant tissue culture for clonal propagation, storage and transportation of propagules for transplantation is a major problem. To overcome this problem, in recent years the concept of synthetic or artificial seeds has become popular, where somatic embryos are encapsulated in a suitable matrix (e.g. sodium alginate), along with substances like mycorrhizae, insecticides, fungicides and herbicides. In. India, this technique of synthetic seeds is being standardized and practiced for sandalwood and mulberry at BARC (Bombay) under the leadership of Dr. P.S. Rao. Synthetic seeds have many advantages including the following: (i) they can be stored upto a year without loss of viability; (ii) they are easy to handle, and useful as units of delivery; (iii) they can be directly sown in the soil like natural seeds and do not need hardening in greenhouse. The only limitation of synthetic seeds, is the high cost of their production. However, this may go down in future, so that these synthetic seeds will become popular at the commercial scale in due course of time.
  13. 13. Protocol-11- Establishment of Cell Suspension Culture PRINCIPLE:  Suspension Culture Growth and Subculture - Cell suspensions are clonally maintained by the routine transfer (subculture) of cells in the early stationary phase to a fresh medium. During the incubation period the biomass of the suspension cultures increases due to cell division and cell enlargement. This continues for a limited period since the viability of cells in suspension after the stationary phase decreases due to the exhaustion of some factors or the accumulation of toxic substances in the medium. At this stage an aliquot of the cell suspension with uniformly dispersed free cells and cell aggregates is transferred to afresh liquid medium of the original composition. The timing of a subculture is very important.  Types of Suspension Cultures - There are mainly two types of suspension cultures, batch cultures and continuous cultures. Batch cultures are maintained by propagating a small aliquot of the inoculum in the moving liquid medium and transferring it to a fresh medium at regular intervals. Generally cell suspensions are grown in flasks (100-250 ml) containing 20-75 ml of the culture medium: The biomass growth in batch cultures follows a fixed pattern.
  14. 14. Protocol-12- Culture of single Cells (Bergmann’s cell plating technique)  PRINCIPLE: PRINCIPLE:  In this technique, free cell are suspended in a liquid medium (if cell aggregates are there, the culture is filtered), and a culture medium with agar (0.6-1%) is cooled and maintained at 35ºC in a water bath. Equal volumes of these liquid and agar media are mixed and rapidly spread in a Petri dish, so that cells are evenly distributed in a thin layer, after solidification. The Petri dishes are sealed with parafilm and examined with inverted microscope to mark single cells (marking is done on outer surface of the dish). The Plates are incubated in dark at 25°C and cell colonies developing from marked single cells are used to obtain single cell cultures. Various other methods (e.g. filter paper raft technique; microchamber technique) have also been developed to grow individual cells (Bhojwani and Razdan, 1983) In this technique, free cell are suspended in a liquid medium (if cell aggregates are there, the culture is filtered), and a culture medium with agar (0.6-1%) is cooled and maintained at 35ºC in a water bath. Equal volumes of these liquid and agar media are mixed and rapidly spread in a Petri dish, so that cells are evenly distributed in a thin layer, after solidification. The Petri dishes are sealed with parafilm and examined with inverted microscope to mark single cells (marking is done on outer surface of the dish). The Plates are incubated in dark at 25°C and cell colonies developing from marked single cells are used to obtain single cell cultures. Various other methods (e.g. filter paper raft technique; microchamber technique) have also been developed to grow individual cells (Bhojwani and Razdan, 1983)
  15. 15. Protocol-13- Maintenance of cell line PRINCIPLE: Maintenance of Cultures Cell Lines - When a primary culture produced on a substrate or in suspension has increased to the extent that all the available substrate is occupied or the medium largely consumed, there arises the need to subculture it. From a very heterogeneous primary culture containing many types of cells derived from the original tissue, during sub culturing (passages or transfer) a more homogeneous 'cell line' emerges. The culture now called a cell line can be propagated, characterized and stored. The potential increase in cell number and uniformity of cells, open up a much wider range of possibilities. The term' cell line implies the presence of several cell lineages either similar or distinct. Among these cell lineages, if a particular cell lineage has specific properties, which are identified in bulk of the cells of that culture, it is described then as a 'cell strain'. A "cell line" or 'cell strain' may be finite or continuous, depending upon whether it has limited culture life spans or it is immortal in culture. Finite cell lines grow upto 20-80 population doublings before I extinction. Some commonly used cell lines and cell strains and a comparison of the properties of mite and continuous cell lines is presented

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