GROWTH AND CULTIVATION OF BIOOBJECTS - V. M. Samygin - 2016

CHAPTER 11. PLANT CELL CULTIVATION

In 1922, W. Robbins and Kotte independently demonstrated the feasibility of cultivating ROOT meristem Cells of tomatoes and corn on synthetic nutrient media. These experiments initiated the application of the method of cultivating isolated PLANT CELLS AND Organs. In the 1930s–1960s, thanks to the work of numerous scientists (P. White, R. Gautheret, and others), the number of plant species whose Cells and Tissues were grown in vitro exceeded 150.

The compositions of nutrient media were described, the nutrient requirements of cultures for Vitamins and growth stimulants were determined, and Methods were developed for obtaining and growing large masses of Cell Suspensions, as well as cultivating a single cell isolated from a suspension. Using isolated protoplasts, methods were developed for somatic cell Hybridization via protoplast fusion using polyethylene glycol and the Introduction of viral RNAs, cell Organelles, and bacterial cells into them. Using the meristem culture method, virus-free economically important plants with a high propagation coefficient were obtained.

At present, The Development of methods for submerged cell cultivation, fusion of isolated protoplasts, etc., is actively ongoing. Such studies have led to The Emergence of more productive cell strains adapted to cultivation conditions, which are used to create new forms and varieties of agricultural, medicinal, ornamental, and other plants.

11.1. Methods for Establishing Plant Cell Cultures

The primary type of cultured plant cell is the callus cell. A callus cell, through the division of which callus tissue arises, represents one of the types of Cell Differentiation inherent in higher plants. For a plant, callus is a tissue that arises under exceptional circumstances (usually upon injury) and Functions for a short time. This tissue protects the injured site and accumulates nutrients for anatomical regeneration or the generation of a lost organ. Much less frequently, cultures of plant tumors of various origins are cultivated. Tumor cell cultures, regardless of the cultivation method, differ little morphologically from callus cell cultures. An important physiological difference between them is the hormone independence of tumor cells, which allows them to divide and grow on nutrient media without The addition of phytohormones or their analogues. However, tumor cells lack The ability to give rise to normally organized structures.

To obtain cultured callus cells in vitro, tissue fragments from various organs of higher plants (explants) are placed on an artificial nutrient medium in test tubes, flasks, or Petri dishes. The process of obtaining primary callus and maintaining a subcultured line requires strictly sterile conditions. Cells of specialized tissues explanted onto a nutrient medium must dedifferentiate, that is, lose the structures characteristic of their specific function in the plant and return to the state of a dividing cell. Often, the explant used to obtain callus includes tissues whose cells are differentially differentiated.

In a cell preparing for division, the synthesis of all types of RNA is stimulated, tissue-specific antigen Proteins disappear, and proteins specific to dividing cells and callus tissue appear. This indicates A change in Gene activity and the protein machinery of cells during dedifferentiation.

Callus formation is not associated with traumatic impact in all cases. Callus can arise As a result of the proliferation of internal Tissues of the explant independently of the cut surface. The growing callus ruptures tissue layers and develops On the surface. The primary callus formed on the explants after 4–6 weeks (depending on The rate of cell growth) is transferred to a fresh nutrient medium (subcultured). Differentially differentiated cells (including meristematic ones) undergo a complex process of dedifferentiation in vitro, lose their inherent structural Organization and specific functions, and are induced to divide, forming a primary callus. During subculturing, a strain is formed that is characterized by individual genetic and physiological features.

The Main Components of nutrient media for PLANT CELL AND tissue culture are mineral salts (macro- and microelements), a carbon source (usually sucrose or glucose), vitamins, and growth regulators. Sometimes, complex organic additives (casein hydrolysate or a mixture of Amino Acids, Yeast extract, or extracts from various plant organs) are included in nutrient media.

11.2. Methods for Culturing Callus Tissues

11.2.1. Surface Cultivation

Callus tissue culture is grown superficially on a semi-solid agarized medium (Agar concentration of 0.6–1%), a medium using other gelling polymers, or on filter paper bridges or polyurethane foam disks immersed in a liquid nutrient medium. Callus tissue grown superficially represents an amorphous mass of thin-walled parenchymal cells lacking a strictly defined Anatomical Structure. The color of the mass can be white, yellowish, green, or red; pigmented entirely or in zones. As a rule, in long-term subcultured lines, callus tissues lose pigmentation and become looser. Callus cells grown superficially are often used to maintain collections of various strains, lines, and mutants in a growing state, for Plant Regeneration, and to obtain cell suspensions cultured in a liquid nutrient medium.

11.2.2. Suspension Cultivation

Plant cell cultures grown in a liquid nutrient medium are usually called suspension (submerged) cultures. There are certain challenges in obtaining a culture consisting predominantly of individual cells or small aggregates thereof. To address these challenges, methods for obtaining cell suspensions are adapted for plant cells.

Growing cell suspensions in a liquid nutrient medium has A number of advantages over growing callus cells by surface cultivation. Suspension cultures are more convenient for biochemical and molecular biological experiments—studying enzyme induction and its connection to Cell Cycle events, the expression and repression of specific genes, and the isolation and characterization of mutants.

A suspension culture can be obtained from plant organ fragments (parenchyma disks, etc.), although this route is more labor-intensive and time-consuming. In this case, the explant cells must form a primary callus, and only then will surface callus cells that have entered the liquid medium and multiplied in it give rise to a line capable of growing in suspension. Typically, a callus tissue culture is used to obtain a cell culture. To establish a cell culture, the most viable (proliferating) part of the callus tissue is taken, and its quantity per unit volume of nutrient medium must be 15–20 times greater than during serial cultivation on agar. A nutrient medium of the same composition as for surface cultivation can be used.

The formation of a primary plant cell suspension can result from three processes:

- The breakdown of callus tissue into cells and small cell aggregates at the time of introduction into the liquid nutrient medium;

- the detachment of cells and cell aggregates from The surface of tissue pieces During the first subcultures;

- Cell Division and growth, and the breakdown of proliferating cell aggregates into smaller aggregates and cells.

Prior to subculturing, the primary suspension is fractionated according to sedimentation velocity using a special cylinder (the upper fraction is taken), or filtered through 1–2 layers of gauze, nylon, or metal sieves to remove large dense pieces of callus tissue, explant remnants, and very large aggregates.

For submerged cultivation of plant cells, methods developed for microbiological purposes are applied. Closed or open systems in batch or continuous modes are used. However, during submerged plant cell growth, the turbidostat principle is practically never applied. One of the reasons for this is the destruction of some cells when routed to the optical device. The cultivation of plant cell suspensions in continuous culture setups According to the chemostat principle is used both to study the METABOLISM of cells stably maintained in different Phases of the CELL CYCLE AND for the industrial cultivation of cell biomass to obtain economically important products.

The most studied and widespread mode of submerged cultivation of cell suspensions at present is the closed batch system. In this case, rollers, shakers (usually circular), fermenters with mechanical and magnetic stirrers, or bubble-Column fermenters, where aeration and agitation are carried out by an air flow, are used to aerate and agitate the suspension. Growth criteria in the cultivation cycle include an increase in cell number, as well as their fresh and dry mass. The model growth curve has a typical S-shape, on which the following are distinguished: I — lag phase; II — exponential phase; III — linear growth phase; IV — decelerating growth phase; V — stationary phase; VI — death phase (Fig. 11).

Class="center">Fig. 11. Growth curve of cell suspensions in a closed batch system

In the latent phase (lag phase), no visible growth of the inoculum is observed. At the same time, high Respiration intensity, maximum energy level values, intensive synthesis of DNA, RNA, proteins, and other cellular components are noted, along with a low mitotic index. The duration of this phase depends on the quantity and physiological state of the inoculum, as well as the cultivation conditions. The exponential phase is characterized by maximum growth rates and levels of mitotic activity, as well as the predominance of small meristematic-type cells. Throughout the linear phase, the growth rate remains constant. The deceleration phase is associated with substrate limitation and inhibition by metabolic products. The stationary phase is characterized by a low rate of cell degradation, which is balanced by cell division. In the cell death phase, the specific growth rate takes on a negative value.

The shape of real growth curves can differ significantly from the model regarding the duration of phases. The processes of the growth cycle depend on the plant species, The amount of inoculated material, and cultivation conditions (COMPOSITION OF THE nutrient medium, Temperature, initial pH, composition of the gas phase, and agitation speed). Primary and secondary metabolism in cultured plant cells are species-specific, depend on the type of Differentiation of the initial plant cells, and are regulated by growth conditions. Genetic variation in cells resulting from their cultivation outside the Organism leads to the emergence of genetically and phenotypically distinct cell lines from the primary callus tissue. This allows for the Selection or experimental creation of lines that retain the biosynthetic systems characteristic of the original plant, as well as lines that synthesize entirely new substances.

11.3. Cultivation of Single Cells

The sources of single cells include cell suspensions in a liquid nutrient medium, tissue maceration (maceration: softening; the Separation of cells resulting from the dissolution of the intercellular substance), and isolated protoplasts after they have regenerated their cell walls. Furthermore, isolated protoplasts represent ideal single cells.

The cultivation of single cells makes it possible to obtain clones and investigate genetic and physiological stability or variation during the propagation of clonal material. Single cells are essential for the clonal selection of mutant, hybrid, and transformed lines. Typically, marker genes are introduced into these cells or marker traits are engineered to provide selective conditions for screening.

Macerated plant tissue cells serve as an excellent model for the comparative study of physiological processes in tissues versus single cells. At the same time, when these cells are cultured on a medium that stimulates cell division, they differentiate and form callus cell colonies. Special macerating enzymatic preparations are used to obtain individual macerated cells.

Single cells can also be obtained from suspensions using a micromanipulator, a flow cytometer, or through serial dilutions.

The induction of single-cell divisions is possible through The Use of a very rich nutrient medium, with the volume of the medium kept to a minimum. However, even when all these conditions are met, the percentage of divided cells remains very low. The "feeder layer" or "nurse tissue" methods are more effective. To create a "feeder layer," a cell suspension of the same plant species is used. The Cell suspension must be in the early stage of the growth cycle. The callus culture serving as the "nurse tissue" must also be in an active growth phase. Once the colony grown from a single cell reaches a size of 0.5–1 mm, it can be transferred for further growth onto an agar-solidified nutrient medium—either directly or via a filter placed on the surface of the nutrient agar. The use of a "feeder layer," "nurse tissue," or a minimal volume of medium in which a single cell is placed is associated with a phenomenon known as the "conditioning factor effect." Despite numerous attempts to determine The Nature of the substances inducing single-cell division and The Mechanism of the conditioning factor, this issue remains ultimately unresolved.

11.4. Plant Protoplasts

11.4.1. Methods for Plant Protoplast Isolation

Plant protoplasts are membrane-bound cytoplasmic entities containing intracellular organelles that exhibit structural integrity and the ability to carry out active metabolism, Biosynthesis, and energy transformation. The isolation of plant protoplasts was first performed in 1892 by G. Klercker.

The simplest, although time-consuming and labor-intensive, methods for obtaining plant protoplasts are mechanical methods. In this approach, a piece of plant tissue is placed in a concentrated sucrose solution and kept there until the protoplasts shrink and pull away from the cell walls; the epidermis is then carefully cut, releasing the protoplasts into the medium. However, even when using modified mechanical methods, only a limited number of protoplasts can be obtained.

A fundamentally different method for obtaining isolated protoplasts is the enzymatic method, which utilizes Enzymes to remove The Cell wall. Compared to the mechanical method, enzymatic isolation offers distinct advantages. This approach allows for a relatively rapid and simultaneous yield of A large number of protoplasts that have not undergone severe osmotic shrinkage, thereby leaving the cells more intact.

Three MAIN TYPES OF enzyme preparations are used to remove the cell walls: cellulases, hemicellulases, and pectinases, most commonly of fungal or bacterial origin. As a result of enzymatic Treatment, a mixture containing protoplasts, cellular debris, and whole cells is formed. To separate the protoplasts from impurities, the suspension is filtered through nylon meshes and then subjected to gentle centrifugation.

In addition to plant tissue fragments, cell suspensions and callus cultures serve as sources for protoplast isolation. While isolating protoplasts is a relatively straightforward and technically well-established Procedure, the challenge lies in obtaining viable protoplasts and ensuring their subsequent cultivation. The success of the process depends on numerous factors: the composition and quality of the enzymes, the pH of the medium, the choice of osmoticum, the physiological state of the plant material, and the methods used for isolation and cultivation. Standardized growing conditions for the source plants or cells, determining the optimal age of the plant or organ for isolation, as well as proper temperature, lighting, and Nutrition are crucial for consistently obtaining high yields of protoplasts. For obtaining protoplasts from suspension cultures, the late logarithmic growth phase is optimal, as the cell walls are most susceptible to enzymatic degradation and the protoplasts exhibit maximum viability.

11.4.2. Cultivation of Plant Protoplasts

For protoplast cultivation, the liquid drop method can be used, in which a protoplast suspension (as droplets in a liquid medium) is placed in plastic Petri dishes. This method ensures good gas exchange and diffusion of excreted products into the solution, while allowing fresh solution to be added at the required concentration. However, in this setup, protoplasts tend to aggregate in the center of each droplet, producing significant amounts of phenolic or other toxic compounds. This method is also inconvenient when tracking the development of individual protoplast colonies is required.

Another common approach is the agar culture method, in which a specific volume of protoplast suspension in a liquid nutrient medium is poured into a plastic Petri dish, and an equal volume of a similar medium containing 1% agar-agar (melted and cooled to 45°C) is added. The dishes are sealed with Parafilm and cultured at 28°C. The protoplasts are fixed in a single position and spatially separated from one another. A major advantage of this method is that it allows researchers to observe all Stages of development of a specific protoplast: Cell wall formation, cell division, and Plant GROWTH AND DEVELOPMENT. The drawback is potential mechanical damage to the protoplasts during mixing with the warm agar-agar.

A variation of this technique involves the use of "nurse" protoplasts or cells exposed to X-ray or gamma irradiation, which are mixed with viable protoplasts. The irradiation dose is selected such that the cells lose their capacity for division while continuing to support and stimulate the growth of other cells. This method enables the cultivation of protoplast suspensions at lower densities than those typically required for growth.

A similar approach is the co-culture method, used for recalcitrant protoplasts. Such protoplasts are cultivated alongside rapidly growing protoplast lines. The success of this cultivation relies on active growth factors secreted by the fast-growing species.

A convenient variation of the liquid drop method is the cultivation of single protoplasts in a micro-volume (up to 1 µL), known as micro-isolation. In these micro-droplets, The ratio of cell volume to nutrient medium volume is comparable to that of a culture with a density of about 1,000 cells/mL.

In terms of nutritional requirements, isolated protoplasts resemble whole cells; consequently, their nutrient media are similar to those used for cell cultures. Since the primary challenge in cultivating plant objects on artificial media is establishing adequate buffering capacity, conjugated salt pairs—combining chemically and physiologically acidic and basic salts—are typically employed. The main conjugated pairs include salts containing nitrogen and phosphorus. The media incorporate iron in chelated form and sucrose as a carbon source. The addition of Cytokinins to the medium is effective for inducing protoplast division. Cultivation temperature is species-specific and varies over a wide range. The temperature regime must be strictly maintained, as protoplasts are sensitive even to minor deviations from the optimum. Optimal light intensities can range from complete darkness to bright light. Protoplast plating density is another critical factor in cultivation: at low densities, protoplasts often fail to divide, whereas extremely high densities lead to complications in later stages due to the accumulation of toxic Metabolic waste products in the growth medium. The optimal protoplast density in culture ranges from 10³ to 10⁵ cells/mL.

Thus, by employing diverse methods, accounting for the Specific characteristics of the source material, and strictly maintaining all required conditions, the cultivation of plant protoplasts can be successfully achieved, leading to the regeneration of whole plants from single protoplasts. The development of plant cell culture techniques, alongside the isolation and cultivation of plant protoplasts, has laid the foundation for Modern Methods of plant bioengineering with predefined traits.