Colorado Agriscience Curriculum - Download as DOC

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					Colorado Agriscience Curriculum
Section: Unit 4: Plant Reproduction Plant Reproduction & Genetics Cloning African Violets Lab

Lesson Number 7:

Colorado Agricultural Education Standards: AS 11/12.4 The student will demonstrate an understanding of physiological processes in agriculturally important plants HRT 11/12.2 The learner will understand and describe the anatomy of horticulture plants. Colorado Science Standards: SCI 1.1 Asking questions and stating hypotheses, using prior scientific knowledge to help guide their development SCI 3.1 Students know and understand the characteristics of living things, the diversity of life, and how living things interact with each other and with the environment. Using examples to explain the relationship of structure and function in organisms. Describing the pattern and process of reproduction and development in several organisms.

SCI 3.3.5 SCI 3.3.6

Student Learning Objectives (Enablers) As a result of this lesson, the student will … Objective 1: Successfully clone African Violets using tissue culture.

Time: Instruction time for this lesson: Two Class periods of 55 minutes. Source: AgriScience: Fundamentals and Application (pp. 350-354) Tools, Equipment, and Supplies 60” x 80” sheets of 2,3, or 4 mil clear plastic agar plates Petri dishes Cotton balls Hand lens or dissecting microscope Scalpel or razor blade Alcohol Bulldog Clips (2-3” long)

Frames for hanging folders African Violets Test tubes Agar (medium) scissors spray bottles Spinach leaves 70% ethanol

Distelled water 250ml Erlenmeyer flask 20 x 150 mm test tubes 6” scalpel handle with # 11 Autoclave or Pressure Cooker

100ml beaker 500ml beaker curved 8” forceps 200ml beaker

Interest Approach – Day 1 Plant cloning by tissue culture is one of the most widely used biotechnologies. Most potatoes and many houseplants are propagated by cloning. Cloning generates multiple, genetically identical offspring from the nonsexual tissues of a parent plant. In theory, cloning is simple; Cut a leaf off a plant, disinfect it, cut into fragments, then plant the fragment in nutrient agar. This may take thirty minutes. In practice, contaminants from the air, hands, and tools quickly take over. Instead of healthy clones, you get colorful molds and bacteria. You can minimize contamination by using a simple hood and aseptic handling techniques. Cloning African violets in the classroom is a long-term project, but it can be done within a few class periods. The first stage takes about 30 minutes. The violets can be transplanted to a minigreenhouse 6 to 8 weeks later. In another few weeks, the plant will be ready for repotting. Are you ready to begin?

Summary of Content and Teaching Strategies
Objective 1. Successfully clone African Violets using tissue culture.

Aseptic handling is critical for successful plant tissue culture. The culture vessel is a battleground between rapidly growing microbes and slowly regenerating plant fragments. Five simple techniques of aseptic handling will minimize transfer of contaminants, and thus growth of molds and bacteria. These techniques are: 1. Wash hands thoroughly and scrub nails using regular soap and paper towels. Do not touch your face or other objects or put your hands in your pockets. Such practices put contaminants back on your hands and can recontamination your plants, equipment, work area, or medium. 2. Keep your hands from passing over open vessels. 3. Touch vessels far from the rim, neck, and similar areas. Keep the caps on when not in use. 4. Grasp tools as far from the working ends as possible. 5. Use sterile materials and equipment.

Now lets try a couple of practice activities to get the hang of cloning. 1. Make thumbprints on bacteriologic nutrient agar plates before and after hand washing. Incubate overnight in a warm (not greater then 99 degrees) place. Bacterial colonies will appear on both plates. These bacteria are normal for us but will hinder cloning. 2. Study violet leaf fragments using a hand lens or dissecting microscope. Note the hairs protruding from the leaf’s upper surface. Now look at your fingers. Note the many ridges. Where do contaminants hide in each case? How does washing the hand and leaf change how each looks? 3. Practice aseptic handling of tools. How do you pass scissors at home? How would you do this aseptically? Why might a scalpel be better for cloning work than a single-edge razor when both cut just fine? 4. Conduct a dry run of the cloning procedure using a spinach leaf. A spinach leaf bruises as easily as a violet leaf, so it readily shows how gently it has been handled. Constructing a Hood for cloning. Materials: 60” x 80” sheet of 2, 3, 4 mil clear plastic (painter’s tarp), bulldog clips (2” to 3” in length, and support frame for hanging file folders. Assembly 1. Place a file folder frame on the table with its arms facing you. 2. Fold the plastic sheet so it is two layers thick. 3. Drape the folded sheet with the fold line in front, so it overhangs the arms by about 2 inches. 4. Clamp the sheeting to the top of the arms to form a flat roof. 5. Straighten the sheeting to minimize creases. 6. Spray the inside of the hood and the work surface with 70 percent ethanol. Dry only the work surface, not the plastic. (Save this step for the first item on day 2) The hood will look like a lean-to with a short curtain valance in front. This fits well on a student’s desk. The overhanging plastic on the sides and back creates a larger work area than just the frame. When they are working they need to stand over the hood. Do not breathe onto the cloning materials or work area.

Day 2: Complete step six from yesterday. Material Preparation: To sterilize materials 1. Mix and sterilize plant nutrient agar. You man either use the recipe from the last lesson or another type is Murashige African violet/gloxinia multiplication medium, available from Carolina Biological Supply Co. One pack makes 30 to 40 plates. Stir one package into 1 liter distilled water. Add 30g sucrose and 15 g agar. The agar will not dissolve until it is heated. Loosely cap the flask with aluminum foil and autoclave or other wise heat for 15 minutes. Cool in hot tap water of about 122 degrees. Pour 25 to 30 ml agar solution into each plastic Petri dish (20 x 100 mm size). Gel at room temperature, then store in a refrigerator. 2. Sterilize a 100 ml beaker to hold disinfected leaves. Cap with a 4-inch square of heavyduty aluminum foil. 3. Sterilize 150 ml tap water in a foil-capped, 250 ml Erlenmeyer flask. 4. Wrap the glass Petri plate (cover and bottom assembled) in a double layer of white Tshirt rag, then autoclave (pressure cook). The inside of the cover will be used as the cutting surface, and the rim of the bottom will be used to support the tools as they drain. 5. Assemble but do not sterilize: a. 500 ml beaker for waste liquid b. 20 x 150 mm test tube filled to the brim with 70% ethanol (support in a 250 ml Erlenmeyer Flask) c. Curved 8-inch forceps and a 6-inch scalpel handle with # 11 blade d. 200 ml beaker containing 100 ml of 70% ethanol to dip leaf e. Single-edge razor to cut the leaf from the plant f. 100 ml beaker to hold the leaf during disinfection 6. Prepare the disinfectant. Mix a 20% solution of liquid chlorine bleach and add on drop of Joy detergent per 500 ml. Swirl gently to mix; too many bubbles will inhibit wetting of the leaf surface. (Remember those little leaf hairs) 7. Soak the forceps and scalpel in the ethanol tube for at least 5 minutes before use. Do not store in the ethanol because the blade will rust. 8. Place in the hood: a. Plant nutrient agar plate b. Sterile beaker c. Sterile water flask d. Sterile glass Petri dish e. 500 ml waste beaker f. Test tube of ethanol g. Forceps and scalpel h. 200 ml beaker with ethanol

Cloning Procedure: 1. Cut a young leaf so the petiole (stem) remains attached. 2. Put the leaf and petiole in the non-sterile beaker with disinfectant to remove dirt, mites, or other vermin. Leave it there for 10 minutes, but swirl it occasionally. 3. Working under the plastic cover, use the alcohol-soaked forceps to transfer the leaf, by the petiole, to the small sterile beaker. Pour the disinfectant into the waste beaker and remove the non-sterile beaker. 4. Rinse the leaf with 50 ml of sterile water. Swirl and pour the water into the waste beaker. Gently hold the leaf in the small beaker with the forceps. 5. Wash again. 6. Disinfect the leaf by dipping the leaf in the ethanol leaf soak, count to 10, and remove the leaf. Meanwhile, open the glass Petri plate. Use the bottom plate to drain the forceps and the blade. (Re-soak the forceps before cutting) 7. Cut the leaf into fragments. Use the lid of the glass Petri plate as the operating table. a. Cut off the petiole. Do not plant it. b. Cut down the midrib firmly c. Cut across each half into four pieces, begin sure to cut through a branching vein 8. Plant each fragment in the plant nutrient agar so each piece is in, not just on, the agar. Plant in a spoke arrangement to minimize spread of contaminants. 9. Put the lid back on and cover the plastic Petri plate with plastic cling wrap to keep moisture in. 10. Store in the dark for about 1 week, then place where the fragments have a daily light cycle and 68 to 77 degree temperatures. 11. Observe weekly. If part of a plate becomes contaminated, transfer healthy fragments to a fresh plate. Even with a commercial hood with sterile air, only about 50% of the plates remain completely free of contaminants. 12. Plantlets will appear on some fragments after about 8 weeks. When plantlet leaves grow to about 0.5 cm, aseptically remove the fragment. Separate its plantlets and return the plate to incubate. Gently cut the plantlets, being sure to have some root and some shoot. Place the individual plantlets in growing medium to develop under aseptic conditions. 13. After about a month, the plants should be large enough to transplant into a pot in sterile potting medium 14. Plants should be planted with at least two per pot and fertilized with African violet food. 15. A mini-greenhouse for maintaining high humidity around fragile plants can be made by setting the plants in a plastic sandwich bag and adding enough water to maintain some moisture on the inside of the bag.


				
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