Aseptic Transfer Technique for Plant Tissue Culture | Lab Protocol

Three types of plant growth regulators control organogenesis: auxins, cytokinins, and gibberellins. Cytokinins exist in inactive form and become active when reacting with auxins, promoting cell division (cytokinesis). Auxins regulate root formation, while cytokinins promote shoot development. The ratio determines organ type: high cytokinin/low auxin induces shoot formation; high auxin/low cytokinin induces root formation. Equal concentrations cause general cell division without preferential organ formation. These hormonal interactions are fundamental to directing plant development in tissue culture systems.

Plant growth regulators are chemical compounds that control plant development. Cytokinins promote shoot formation, while auxins promote root formation. The ratio between these two types determines the outcome: high cytokinin-to-auxin ratios produce shoots, high auxin-to-cytokinin ratios produce roots, and equal ratios induce callus formation—a mass of undifferentiated cells. Commonly used cytokinins include TDZ and BAP, while common auxins include NAA and IBA.

Plant growth regulators are small molecules controlling growth. Auxins (indole derivatives) were discovered by Darwin through canary grass experiments and isolated by F.W. Went from oat seedlings. Functions include promoting root formation in stem cuttings, inducing pineapple flowering, preventing early fruit/leaf drop, promoting apical dominance, and inducing parthenocarpy. 2,4-D is a synthetic auxin effective for dicot weed control. Apical dominance occurs when apical buds release auxins inhibiting lateral bud growth; decapitation removes this inhibition for hedge formation. Gibberellins were discovered accidentally from rice blast disease (fungal infection causing 'foolish seedling' syndrome). They are terpenes that promote stem elongation (grape stalks, apples, sugar cane), delay senescence to extend market life, speed up malting in brewing, promote early seed production in conifers, and promote bolting. They are antagonistic to abscisic acid. Cytokinins (adenine derivatives) promote cell division, new leaf/chloroplast formation, lateral/adventitious shoot growth, and delay senescence. They break apical dominance and promote nutrient mobilization. Memory aids help retention: 'Bold Mali Gabbur' for gibberellins (Bold=bolting, Mali=malting), 'Selina Shoots' for cytokinins (Selina=cell division, Shoots=new shoots and chloroplasts).

Plant tissue culture, also known as micropropagation, is a technique that utilizes the principle of cellular totipotency—the ability of a single plant cell to regenerate into a complete organism. The process involves collecting an explant (a small piece of plant tissue), sterilizing it, and placing it on a nutrient medium containing plant hormones (auxins and cytokinins). By manipulating the ratio of these hormones, scientists can induce different developmental stages: equal ratios promote callus formation (undifferentiated dividing cells), excess cytokinin relative to auxin stimulates shoot development, and excess auxin relative to cytokinin encourages root formation. After developing into plantlets, they undergo a hardening process to adapt to outdoor conditions before transplantation. This method allows for rapid, large-scale production of genetically identical plants (clones) while also enabling the propagation of disease-free plants from infected source material.

Callus culture is a plant tissue culture technique where plant explants (such as carrot root pieces) are placed on nutrient medium containing plant growth regulators (auxins, cytokinins, gibberellins) to induce the formation of an unspecialized, unorganized mass of cells called callus; the procedure involves selecting fresh plant material, washing it, surface sterilizing with 5% teepol solution followed by sodium hypochlorite solution, isolating the cambium tissue, transferring it to nutrient medium in a test tube, and incubating at 25°C for 6-8 weeks until callus forms through cell division; callus can be either compact (densely aggregated cells) or friable (loosely associated cells), and it lacks photosynthetic capability since it develops in dark conditions.

Tissue culture plantlets require a gradual acclimation process lasting approximately two weeks to adapt from their high-humidity laboratory environment to greenhouse conditions, during which growers must carefully manage humidity (maintaining vapor pressure deficit below 0.5 kPa), light intensity (recommending 70+ micromoles with attention to radiant heat from HID lights), and substrate moisture to prevent wilting and ensure successful establishment; key challenges include the absence of functional cuticles, underdeveloped root systems, and disorganized chloroplasts in newly transferred plantlets.

This extensive section details the critical humidity gradient approach for acclimating tissue culture plantlets. It covers maintaining 99% humidity for the first 7-10 days, transitioning to 92% humidity for another 7-10 days, then gradually introducing normal room conditions (40-50% humidity). The process includes using heat mats from 8 PM to 8 AM for root development, monitoring temperatures to prevent overheating, and adjusting environmental controls based on season (opening doors in summer, using humidifiers in winter). The goal is to ensure plants can thrive in standard home conditions without special equipment.

To successfully acclimate tissue culture plants to ex-vitro conditions, first transfer them to a highly porous growing medium such as perlite-vermiculite-peat moss mix, then gradually reduce humidity by slowly opening germination dome vents over several days before completely removing the dome and exposing plants to normal environmental conditions; this gradual transition helps plants adapt to lower humidity and direct light without shock.

Acclimation (hardening off) is the final stage in tissue culture where baby plants transition from laboratory conditions to outdoor environments. Plants grown in tissue culture live in near-perfect conditions and cannot survive direct transplantation to soil. The process requires gradual adjustment to ambient conditions. Essential equipment includes seed starter trays with adjustable vents for humidity control. Recommended substrate is a 50% mix of fluorite stratum and perlite, kept moist but not wet. Before transplanting, plants must be thoroughly cleaned in room temperature water to prevent mold growth from residual sugar in tissue culture media.

In places where weather is conducive, the plantlets are hardened for 4 to 6 weeks in mini sand beds with 90 to 95% humidity maintained for the initial 6 to 8 days under diffused light. The humidity is slowly reduced to 70%, light intensity raised to normal, and temperatures maintained at 24 to 26°C by the end of 6 weeks.

Some bacterial contaminants (Bacillus, Erwinia, Pseudomonas) may remain inside plantlets without detection until late in the culture period, causing latent infestation. These require identification procedures in commercial labs. Antibiotics are sometimes used in medium to control bacterial growth, but prolonged use should be avoided. Plant Preservative Mixture (PPM) is a new product that dramatically reduces infestation rates when added to culture medium.

Both antibiotics and Plant Preservative Mixture (PPM) can be effective for controlling contamination in tissue culture, with effectiveness depending on the specific organisms present. PPM is advantageous because it works against both bacteria and fungi simultaneously, providing broad-spectrum protection. Specific antibiotics may be more effective against certain bacterial species—for example, tetracycline may be particularly effective against some bacteria. The choice between antibiotics and PPM depends on the nature of contamination encountered and the specific requirements of the tissue culture protocol.

Latent contamination occurs when microorganisms remain dormant within plant tissues for extended periods before suddenly appearing. After initial appearance of healthy growth, contamination may emerge after 3 weeks or more despite no prior signs. This delayed manifestation makes contamination control challenging. To detect latent contamination, laboratories should test media in incubators for at least one week before use to ensure no hidden contaminants will later compromise cultures.

This video demonstrates how to subculture a Cephalotus plant that has become contaminated and is dying due to lack of media, showing the practical application of Plant Preservative Mixture (PPM), a broad-spectrum biocide/fungicide designed to combat microbial airborne, waterborne, and endogenous contamination in plant tissue culture environments.

Professional tissue culture uses Plant Preservative Mixture (PPM) to maintain sterility in packages. Despite high-tech equipment and protocols, contamination can still occur because contamination agents are invisible. Contamination can spread rapidly within days of package arrival, similar to how bacteria spread in general. The contamination may have occurred during shipping or storage. Plants with visible contamination should not be shipped out. The key is understanding that tissue culture plants are living things with multiple growth points, and some imperfection is normal. The balance between expecting perfect plants and accepting natural plant variability is essential for successful tissue culture.

Meristem culture eliminates viruses through rapid cell division outpacing viral replication, producing disease-free plants for commercial micropropagation. Embryo rescue saves aborting embryos from interspecific crosses through tissue culture, enabling hybrid recovery and dormancy bypass. Hairy root culture uses Agrobacterium rhizogenes to produce highly branched, fast-growing roots for secondary metabolite production and metabolic engineering. Synthetic seeds encapsulate somatic embryos in calcium alginate matrices, providing protection for storage, transport, and direct sowing. These techniques enable clonal propagation, metabolic pathway modification, and long-term plant material preservation.

Meristem culture is a plant tissue culture technique that excises the meristem region (a small region of active cell division less than 0.1 mm) to produce virus-free plants, as the high mitotic rate and absence of differentiated vascular tissues prevent viral contamination; this technique is primarily used for virus elimination, germplasm conservation, and genetic engineering applications.

Meristem culture is a laboratory technique that regenerates healthy, virus-free plants from infected plant material by culturing the apical meristem, which is the only plant tissue capable of being free from viruses; this technique involves disinfecting plant material, dissecting the meristem under sterile conditions, and culturing it on a development medium until plantlets can be acclimatized to greenhouse conditions, effectively eliminating viral contamination in ornamental plants like carnations and research plants like Arabidopsis halleri.

Plant transformation involves introducing foreign DNA into plant cells using methods including Agrobacterium-mediated transformation, particle bombardment (biolistics), and electroporation. Meristem culture is used to produce virus-free plants, as viruses typically do not infect the meristem tissue. The regenerated plants from meristem culture are virus-free and can be used for producing healthy planting material. This technique is widely used in propagation of fruit trees and other economically important crops.

Meristems are plant regions containing undifferentiated cells where growth occurs, classified as apical (at root and shoot tips) and lateral (along stems/roots). Meristem culture, discovered by Morel and Mettner in 1952, uses shoot apical tips as explants to produce virus-free plants through in vitro culturing. The three-stage process involves: Stage 1 - Shoot initiation using cytokinins; Stage 2 - Multiplication with high cytokinin levels for genetic stability; Stage 3 - Root regeneration using auxins. Key components include inorganic nutrients, vitamin-rich organic nutrients, plant growth hormones, and solidifying agents. The practical procedure involves isolating shoot tips, surface sterilization in sodium hypochlorite, rinsing in distilled water, removing outer leaves, transferring to MS medium, and incubating at 25°C with fluorescent light.