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Important structures in plant development are buds, shoots, , leaf, and ; produce these tissues and structures throughout their life from Review. located at the tips of organs, or between mature tissues. Thus, a living plant always has embryonic tissues. By contrast, an animal embryo will very early produce all of the body parts that it will ever have in its life. When the animal is born (or hatches from its egg), it has all its body parts and from that point will only grow larger and more mature. However, both plants and animals pass through a phylotypic stage that evolved independently and that causes a developmental constraint limiting morphological diversification.
According to plant physiology A. Carl Leopold, the properties of organization seen in a plant are emergence which are more than the sum of the individual parts. "The assembly of these tissues and functions into an integrated multicellular organism yields not only the characteristics of the separate parts and processes but also quite a new set of characteristics which would not have been predictable on the basis of examination of the separate parts."
Additionally, mature plant parts, including leaves, stems, roots, petioles, and flower segments, can also serve as viable explants for organ formation under suitable conditions. Plant regeneration occurs through the formation of callus, an undifferentiated mass of cells that later gives rise to new organs. Callus formation can be induced from various explants, such as Cotyledon, Hypocotyl, Plant stem, Leaf, shoot apices, roots, Inflorescence, and floral structures, when cultured under controlled conditions.
Generally, explants containing actively dividing cells are more effective for callus initiation, as they have a higher capacity for cellular reprogramming. Immature tissues tend to be more adaptable for regeneration compared to mature tissues due to their increased developmental plasticity. The size and shape of the explant also influence the success of culture establishment, as larger or more structurally favorable explants may enhance the chances of survival and growth. Callus development is primarily triggered by wounding and the presence of plant hormones, which may be naturally present in the tissue or supplemented in the growth medium to stimulate cellular activity and organ formation.
The interaction between auxins and cytokinins in regulating organogenesis is well-established, though responses vary by species. Some plants, such as tobacco, can spontaneously form shoot buds without exogenous growth regulators, while others like Scurrula pulverulenta, Lettuce, and Brassica juncea strictly require hormonal supplementation. In B. juncea cotyledon cultures, benzylaminopurine (BAP) alone induces shoot formation from petiole tissue, similar to Pinus radiata where cytokinin alone suffices for shoot induction.
Research indicates that endogenous hormone concentrations, rather than exogenous application levels, ultimately determine organogenic differentiation. Among the various Cytokinin (2iP, BAP, thidiazuron, kinetin, and zeatin) used for shoot induction, BAP has demonstrated superior efficacy and widespread application. Auxin similarly influence organogenic pathways, with 2,4-D commonly used for callus induction in cereals, though organogenesis typically requires transfer to media containing IAA or NAA or lacking 2,4-D entirely. The auxin-to-cytokinin ratio largely determines which organs develop.
Gibberellic acid (GA3) contributes to cell elongation and meristemoid formation, while unconventional compounds like tri-iodobenzoic acid (TIBA), abscisic acid (ABA), kanamycin, and auxin inhibitors have proven effective for recalcitrant species. Natural additives like ginseng powder can enhance regeneration frequency in certain cultures. Since ethylene typically suppresses shoot differentiation, inhibitors of ethylene synthesis such as aminoethoxyvinylglycine (AVG) and silver nitrate (AgNO3) are often employed to promote organogenesis, with documented success in wheat, Nicotiana, and Common sunflower cultures.
Agar is not an essential component of the culture medium, but quality and quantity of agar is an important factor that may determine a role in organogenesis. Commercially available agar may contain impurities. With a high concentration of agar, the nutrient medium becomes hard and does not allow the diffusion of nutrients to the growing tissue. It influences the organogenesis process by producing adventitious roots, unwanted callus at the base, or senescence of the foliage. The pH is another important factor that may affect organogenesis route. The pH of the culture medium is adjusted to between 5.6 and 5.8 before sterilization. Medium pH facilitates or inhibits nutrient availability in the medium; for example, ammonium uptake in vitro occurs at a stable pH of 5.5 (Thorpe et al., 2008).
Similar seasonal dependency is observed in Chlorophytum borivillianum, a medicinally valuable species that shows markedly enhanced in vitro tuber formation during monsoon seasons compared to other times of year. This seasonal variation in morphogenic potential likely reflects differences in the physiological state of the source plant, including endogenous hormone levels, carbohydrate reserves, and metabolic activity that fluctuate throughout the annual growth cycle.
The regulatory effect of different wavelengths demonstrates how light quality can selectively control morphogenic outcomes. Artificial fluorescent lighting produces variable responses depending on the species, promoting root formation in some cultures while inhibiting it in others. Some species exhibit specialized light requirements, as observed in Pea (garden pea), where shoot bud initiation occurs optimally in darkness before exposure to light stimulates further development.
For most tissue culture applications, standard lighting protocols typically recommend illumination of approximately 2,000-3,000 lux intensity with a 16-hour photoperiod. However, certain species demonstrate exceptional light intensity requirements, exemplified by Nicotiana tabacum (tobacco) callus cultures, which require substantially higher light intensities of 10,000-15,000 lux to induce shoot bud formation or somatic embryogenesis.
Geophytic species from temperate regions typically require lower temperature regimes than the standard protocol. Notable examples include bulbous plants such as Galanthus (snowdrop) which exhibits optimal growth at approximately 15°C, while certain cultivars of Narcissus (daffodil) and Allium (ornamental onion) demonstrate enhanced regeneration efficiency at around 18°C.
Conversely, species of tropical origin generally require elevated temperatures for optimal growth and organogenesis in culture. Date palm cultures thrive at 27°C, while Monstera deliciosa (Swiss cheese plant) exhibits peak regenerative performance at 30°C. These temperature requirements reflect evolutionary adaptations to the plants' native environmental conditions.
The conclusion of the induction phase is marked by a cell or group of cells committing to either shoot or root formation. This determination is tested by transferring the tissue from a growth regulator-supplemented medium to a basal medium containing essential minerals, vitamins, and a carbon source but no plant growth regulators. At this stage, the tissue completes the induction process and becomes fully determined to its developmental fate.
A key concept in this process is canalization, which refers to the ability of a developmental pathway to consistently produce a standard phenotype despite potential genetic or environmental variations. If explants are removed from a shoot-inducing medium before full canalization occurs, shoot formation is significantly reduced, and root development becomes the dominant outcome. This phenomenon highlights the morphogenic plasticity of plant tissues in vitro, demonstrating their ability to adjust to external conditions and developmental cues.
The sequential development of organogenesis can be observed in species such as Pinus oocarpa Schiede, where shoot buds are regenerated directly from cotyledons through direct organogenesis. However, the specific developmental patterns may vary across different plant species grown in vitro. The progression of organ formation includes distinct morphological changes, beginning with alterations in surface texture, the emergence of meristemoids, and the expansion of the Meristem region either vertically or horizontally. This is followed by the protrusion of the meristematic region beyond the epidermal layer, the formation of a structured meristem with visible leaf primordia, and eventually, the full development of an adventitious bud.
A notable characteristic of in vitro organogenic cultures is the simultaneous formation of multiple meristemoids on a single explant, with varying degrees of differentiation. Within the same explant, buds may exist in different developmental stages, ranging from early initiation to fully developed structures. Once the elongated shoots surpass a length of 1 cm, they are transferred to either in vitro or ex vitro rooting substrates, allowing for the completion of plantlet regeneration and the establishment of a fully formed plant.
However, there are some limitations to organogenesis. Somaclonal variation, which can result in unwanted genetic diversity, is a potential issue, particularly in the indirect organogenesis process. Additionally, this technique may not be suitable for recalcitrant plant species, which are those that do not respond well to in vitro culture or regeneration protocols. These limitations highlight the need for ongoing research and optimization of methods for different plant species to overcome these challenges in plant propagation and conservation.
Plant growth and development are mediated by specific and plant growth regulators (PGRs) (Ross et al. 1983).Ross, S.D.; Pharis, R.P.; Binder, W.D. 1983. Growth regulators and conifers: their physiology and potential uses in forestry. p. 35–78 in Nickell, L.G. (Ed.), Plant growth regulating chemicals. Vol. 2, CRC Press, Boca Raton FL. Endogenous hormone levels are influenced by plant age, cold hardiness, dormancy, and other metabolic conditions; Photoperiodism, drought, temperature, and other external environmental conditions; and exogenous sources of PGRs, e.g., externally applied and of Rhizosphere origin.
There is variation among the parts of a mature plant resulting from the relative position where the organ is produced. For example, along a new branch the leaves may vary in a consistent pattern along the branch. The form of leaves produced near the base of the branch differs from leaves produced at the tip of the plant, and this difference is consistent from branch to branch on a given plant and in a given species.
The way in which new structures mature as they are produced may be affected by the point in the plants life when they begin to develop, as well as by the environment to which the structures are exposed. Temperature has a multiplicity of effects on plants depending on a variety of factors, including the size and condition of the plant and the temperature and duration of exposure. The smaller and more Succulent plant the plant, the greater the susceptibility to damage or death from temperatures that are too high or too low. Temperature affects the rate of biochemical and physiological processes, rates generally (within limits) increasing with temperature.
Juvenility or heteroblasty is when the organs and tissues produced by a young plant, such as a seedling, are often different from those that are produced by the same plant when it is older. For example, young trees will produce longer, leaner branches that grow upwards more than the branches they will produce as a fully grown tree. In addition, leaves produced during early growth tend to be larger, thinner, and more irregular than leaves on the adult plant. Specimens of juvenile plants may look so completely different from adult plants of the same species that egg-laying insects do not recognize the plant as food for their young. The transition from early to late growth forms is sometimes called vegetative phase change.
Adventitious roots and buds usually develop near the existing vascular tissues so that they can connect to the xylem and phloem. However, the exact location varies greatly. In young stems, adventitious roots often form from parenchyma between the . In stems with secondary growth, adventitious roots often originate in phloem parenchyma near the vascular cambium. In stem cuttings, adventitious roots sometimes also originate in the callus cells that form at the cut surface. Leaf cuttings of the Crassula form adventitious roots in the epidermis.McVeigh, I. 1938. Regeneration in Crassula multicava. American Journal of Botany 25: 7-11. [1]
Adventitious buds are often formed after the stem is wounded or pruning. The adventitious buds help to replace lost branches. Adventitious buds and shoots also may develop on mature tree trunks when a shaded trunk is exposed to bright sunlight because surrounding trees are cut down. Redwood ( Sequoia sempervirens) trees often develop many adventitious buds on their lower trunks. If the main trunk dies, a new one often sprouts from one of the adventitious buds. Small pieces of redwood trunk are sold as souvenirs termed redwood burls. They are placed in a pan of water, and the adventitious buds sprout to form shoots.
Some plants normally develop adventitious buds on their roots, which can extend quite a distance from the plant. Shoots that develop from adventitious buds on roots are termed Basal shoot. They are a type of natural vegetative reproduction in many species, e.g. many grasses, quaking aspen and Canada thistle. The Pando quaking aspen grew from one trunk to 47,000 trunks via adventitious bud formation on a single root system.
Some leaves develop adventitious buds, which then form adventitious roots, as part of vegetative reproduction; e.g. piggyback plant ( Tolmiea menziesii) and mother-of-thousands ( Kalanchoe daigremontiana). The adventitious plantlets then drop off the parent plant and develop as separate Cloning of the parent.
Coppicing is the practice of cutting tree stems to the ground to promote rapid growth of adventitious shoots. It is traditionally used to produce poles, fence material or firewood. It is also practiced for biomass crops grown for fuel, such as Populus or willow.
The ability of plant stems to form adventitious roots is utilised in commercial propagation by cuttings. Understanding of the physiological mechanisms behind adventitious rooting has allowed some progress to be made in improving the rooting of cuttings by the application of synthetic auxins as rooting powders and by the use of selective basal wounding. Further progress can be made in future years by applying research into other regulatory mechanisms to commercial propagation and by the comparative analysis of molecular and ecophysiological control of adventitious rooting in 'hard to root' vs. 'easy to root' species.
Adventitious roots and buds are very important when people propagate plants via cuttings, layering, tissue culture. Plant hormones, termed , are often applied to stem, shoot or leaf cuttings to promote adventitious root formation, e.g., African violet and sedum leaves and shoots of poinsettia and coleus. Propagation via root cuttings requires adventitious bud formation, e.g., in horseradish and apple. In layering, adventitious roots are formed on aerial stems before the stem section is removed to make a new plant. Large houseplants are often propagated by air layering. Adventitious roots and buds must develop in tissue culture propagation of plants.
An external stimulus is required in order to trigger the differentiation of the meristem into a flower meristem. This stimulus will activate mitosis cell division in the meristem, particularly on its sides where new primordium are formed. This same stimulus will also cause the meristem to follow a developmental pattern that will lead to the growth of floral meristems as opposed to vegetative meristems. The main difference between these two types of meristem, apart from the obvious disparity between the objective organ, is the verticillate (or whorled) phyllotaxis, that is, the absence of plant stem elongation among the successive whorls or verticils of the primordium. These verticils follow an acropetal development, giving rise to , , and . Another difference from vegetative axillary meristems is that the floral meristem is «determined», which means that, once differentiated, its cells will no longer cell cycle.
The identity of the organs present in the four floral verticils is a consequence of the interaction of at least three types of gene products, each with distinct functions. According to the ABC model, functions A and C are required in order to determine the identity of the verticils of the perianth and the reproductive verticils, respectively. These functions are exclusive and the absence of one of them means that the other will determine the identity of all the floral verticils. The B function allows the differentiation of petals from sepals in the secondary verticil, as well as the differentiation of the stamen from the carpel on the tertiary verticil.
To determine pathway regulation, P. hybrida Mitchell flowers were used in a petal-specific microarray to compare the flowers that were just about to produce the scent, to the P. hybrida cultivar W138 flowers that produce few volatile benzenoids. cDNAs of genes of both plants were sequenced. The results demonstrated that there is a transcription factor upregulated in the Mitchell flowers, but not in the W138 flowers lacking the floral aroma. This gene was named ODORANT1 (ODO1). To determine expression of ODO1 throughout the day, Northern blot was done. The gel showed that ODO1 transcript levels began increasing between 1300 and 1600 h, peaked at 2200 h and were lowest at 1000 h. These ODO1 transcript levels directly correspond to the timeline of volatile benzenoid emission. Additionally, the gel supported the previous finding that W138 non-fragrant flowers have only one-tenth the ODO1 transcript levels of the Mitchell flowers. Thus, the amount of ODO1 made corresponds to the amount of volatile benzenoid emitted, indicating that ODO1 regulates benzenoid biosynthesis.
Additional genes contributing to the biosynthesis of major scent compounds are OOMT1 and OOMT2. OOMT1 and OOMT2 help to synthesize orcinol O-methyltransferases (OOMT), which catalyze the last two steps of the DMT pathway, creating 3,5-dimethoxytoluene (DMT). DMT is a scent compound produced by many different roses yet, some rose varieties, like Rose gallica and Damask rose Rose damascene, do not emit DMT. It has been suggested that these varieties do not make DMT because they do not have the OOMT genes. However, following an immunolocalization experiment, OOMT was found in the petal epidermis. To study this further, rose petals were subjected to ultracentrifugation. Supernatants and pellets were inspected by western blot. Detection of OOMT protein at 150,000g in the supernatant and the pellet allowed for researchers to conclude that OOMT protein is tightly associated with petal epidermis membranes. Such experiments determined that OOMT genes do exist within Rosa gallica and Damask rose Rosa damascene varieties, but the OOMT genes are not expressed in the flower tissues where DMT is made.
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