African violets are one of the most beautiful flowering plants native to eastern tropical Africa. They are famous among indoor plants, as they can thrive in artificial light. These plants require a good amount of water for their growth. So, the soil, where African violets are grown, should be porous to allow water to pass through easily.
These plants can adjust well to warmer indoor temperatures and dry air. They require light exposure but not direct sunlight. Moreover, the best temperature for these plants will be 60 ℉ at night and 80-85 ℉ during the day.
When growing the African plants inside your home or office, it is recommended to keep the soil moist, and to avoid pouring water directly on the leaves, which causes disfiguring, light-colored spots, or rings.
This article poses a description of African violets and the procedure of organogenesis in these plants using leaf lamina explant.
Saintpaulia ionantha is commonly known as African Violet. The name Saintpauliais named in honor of Baron Walter von Saint Paul-Illaire, who discovered the plant in German East Africa (now Tanzania) in 1892.
The plant is small perennial herbs that grow 6-15 cm in height. The leaves are thick, hairy, petiolate, ovate, and have a fleshy texture. The flowers are bilaterally symmetric with five-lobed velvety corolla. They grow in clusters on peduncles and have capsule-shaped seeds. Beside violet color, they are also available in pink, fuchsia, and white.
|Figure: The single flower of African violet.|
Note the five velvety petals of the flowers.
Organogenesis of Saintpaulia ionantha
The process of organogenesis is a tissue culture technique in which roots, shoots, leaves, etc. of plants are regenerated in laboratory conditions. In this article, a procedure of inducing organogenesis is shown in the leaf lamina explants of Saintpaulia ionantha.
To learn more about organogenesis you can read our previous blog post, “Organogenesis in Plants."
- Leaves excised from a mature plant of African violet (200 cm3 aqueous solution (10% v/v) and commercial bleach containing a final concentration of approximately 0.5% (v/v) NaOCI. Add a few drops of a liquid detergent as a wetting agent.
- 250-cm3 beakers.
- Stainless steel forceps. Before sterilization, place forceps in the test tube and wrap with foil.
- Petri dishes, culture tubes, Whatman filter paper, Erlenmeyer flasks, MS medium, scalpel.
- 80% and 70% ethanol and distilled water.
- Prepare 1 L of MS medium supplemented with myoinositol (100 mg/I), nicotinic acid (0.5 mg/I), pyridoxine-HCI (0.5 Mg/l), thiamine-HCI (0.4 mg/I), NAA (0.1 mg/I), BAP (5.0 mg/I), and adenine sulfate (80 mg/I).
- Adjust the pH of the medium to 5.7.
- Keep two 100 cm3 aliquots of the medium and then add sucrose (3.0% w/v) and agar (0.7% w/v) in each aliquot.
- Excise many healthy leaves, both young and old, and wash the lamina briefly.
- Sterilize the lamina in 70% ethanol and rinse in a double-distilled 250 ml beaker then immerse the blades in hypochlorite solution for 10 minutes. Then, three times rinse the blades in distilled water.
- Transfer lamina to a sterile petri dish containing filter papers and prepare the explants using forceps and scalpel.
- Slice the blade into rectangles approximately 10-12 mm on a side, ensuring that each explant contains a portion of the midvein of the leaf.
- Place the explants individually in culture tubes in an upright position with one-quarter of the explant embedded in the agar medium.
- Place the cultures in a chamber maintained at 25°C with 16-hr photoperiods furnished by a combination of Gro-Lux and cool-white fluorescent tubes. The light intensity should be approximately 1,000-1,500 lux.
The shoots will appear in 2-4 weeks and after 6-8 days of culture, subdivide the proliferated shoots. Rooting is promoted by transferring the shoots to a medium that is devoid of plant hormones and has a sucrose concentration of approximately 1.6% (w/v).
- Dodds H. John and Roberts W. Lorin (1985). Experiments in Plant tissue Culture, 2nd ed. Cambridge University Press, Cambridge, NY, USA.
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