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STUDENT OBJECTIVE

Students study the structure of monocot and dicot leaves and determine the absorption spectrum of chlorophyll. The rate of oxygen released during photosynthesis in Elodea is measured at various distances from a light source. Students graphically determine the light intensity at which production of O₂ equals the consumption of O₂. Cross sections of C₃ leaves are studied.

EQUIPMENT	AMOUNT
	(Class of 24 with 8 groups)
Compound microscope Spectrophotometers (400 nm-700 nm) Blender Light source setup (200 or 300 watt indoor/outdoor flood bulb, desk lamp)	1/student 1/group 1/lab 1/group
MATERIALS
Leaf structure: Prepared slides (can be shared to save costs) Dicot leaf, cross section Corn leaf (CBS#30-3496) Examples of simple and compound leaves of monocots and dicots Leaves for hand sectioning Chlorophyll absorption spectrum test: (can be demonstration) 500 ml separatory funnel and ring stand 125 ml Erlenmeyer flask Cheesecloth, 20 cm x 20 cm squares Ring stand with two clamps 100 ml graduated cylinder Jar for solvent wastes Oxygen measurement: 800 ml beaker Apparatus for oxygen measurement Test tube, 20 mm x 150 to 200 mm, pipette with bend as in figure 22.7 (1 ml * 0.01) One hole-#3 stopper Syringe (1 ml), #18 needle (assembled as shown in laboratory manual) Ring stand with two clamps Meter sticks	1/student 1/student     1/group 1/group 4/group 1/group 1/group 1/lab   1/group       2/group 1/group 5/lab
Elodea (5"—6" piece) (CBS#15-7340)* Chromatography tank or fish bowl (for heat filter) Razor blades	1/group 1/group 6/lab

*Please refer to the Appendix for name and address of supplier.

SOLUTIONS

Chlorophyll extract (spinach, parsley)
Petroleum ether
80% methanol (CH₃OH)
10% sodium chloride (NaCl)
Acetone (CH₃COCH₃)
0.1 M sodium bicarbonate (NaHCO₃)
Sodium sulfate (Na₂SO₄) anhydrous
Magnesium oxide (MgO) powder

PREPARATION

One Week before Lab

1. Purchase fresh spinach and weigh out 10 grams per lab (can be frozen).
2. Put approximately 3 g sodium sulfate in a small vial with screw top (for removal of water from spinach extract). If not anhydrous (powdery), heat in crucible over bunsen burner before packaging.
3. Methanol preparation:

   80% methanol 80 ml methanol/20 ml distilled water

4. Sodium chloride preparation:

   10% NaCl 50 g NaCl/450 ml distilled water

5. Obtain Elodea and put it under a strong light in a large aerated aquarium. (Plants must be healthy to have noticeable O₂ release for student setups.)
6. Sodium bicarbonate preparation:

   0.1 M NaHCO₃ 8.4 g NaHCO₃/cool tap water to make 1 liter

   Dissolve salt in tap water. If high levels of Cl₂ in the water affect the plant, aerate the water for eight hours before preparing the solution.

7. Collect samples of monocot and dicot leaves for demonstration table in the laboratory.

Day of Lab

1. Chlorophyll extract preparation:
   CAUTION! Acetone and petroleum ether are highly flammable. No flames, heaters, or smoking during this procedure. Students can prepare the extract in lab or instructor can prepare extract before or during lab and students start at the alternative procedure in #2 below.
   
   
   1. Add 20 g frozen spinach to a blender with 100 ml water and 0.2 g MgO; blend until homogenized. (A mortar and pestle can be used if a blender is not available.) Filter through several layers of cheesecloth. Add acetone to the filtrate to bring the extract solution to a final volume of 200 ml. This extract can be capped and stored in refrigerator for use later in the day or can be used immediately in the following procedure. Never blend spinach in acetone; sparks from blender may ignite acetone.
   2. Pour extract into a separatory funnel and add 200 ml of petroleum ether. Shake vigorously; release the pressure that builds up by raising the glass stopper occasionally. Let sit until two layers form. Drain and discard the lower aqueous solution. If layers don’t separate, add some 10% NaCl. Chlorophyll and carotenoids are soluble in nonpolar petroleum ether and will transfer to the upper ether phase.
   3. Add 120 ml of 80% methanol and shake. (Open the stopper or stopcock to release the pressure once during the shaking.) Add 10% NaCl if layers don’t separate. This separates slightly polar carotenoids from chlorophyll. Drain and discard the lower aqueous phase.
   4. The resulting ether phase should be washed twice with 60 ml of 10% NaCl to remove the methanol and acetone. Drain and discard the aqueous phase.
   5. To remove the water from the ether phase, add the vial of anhydrous Na₂SO₄ and swirl to mix.
   6. From the top of the separatory funnel, pour off the resulting clear, green solution to be used in the determination of the absorption spectrum for chlorophyll.
   7. Chlorophyll extracts used by students can be collected after the lab and stored in the freezer for backup use in future laboratories. Extract will keep indefinitely. However, the spectrum should be checked to determine if evaporation has occurred. (Absorbance should be 1.0 to 1.5 at 425 nm.) Addition of petroleum ether will dilute the extract to the proper concentration needed for use in determining the spectrum.
   8. Dispose of petroleum ether according to local safety/environmental standards.
2. Alternative procedure for providing student with chlorophyll for the determination of its absorption spectrum:
   Before the lab, prepare 120 ml chlorophyll extract according to the above-described method. Store in freezer at —20^oC. The morning of the lab, pour 8 ml of extract into each cuvette and plug with a shortened "00" cork (not rubber) stopper. (The stopper should remain in the tube throughout the lab, therefore, it should be modified so that the lid of the spectrophotometer closes easily but securely.) For the blank, pour 8 ml petroleum ether into another cuvette and cap. If the solutions are not capped, the concentration can be altered by evaporation. Add petroleum ether if evaporation occurs.

NOTES

1. Spectrophotometers should have a broad spectrum photo tube installed. Otherwise, readings above 625 nm may not be possible.
2. Apparatus assembly: Using a hot flame, bend a pipette into a right angle 3" from the top (wide) end. Rest the hot tube on a flat surface until it is cool. Place a drop of glycerin in the hole of the stopper and insert the small bent end of the pipette into the hole. Push until 3/8" of tube projects from the stopper. Heavy gloves will protect the hands if the glass breaks while twisting and pushing the tube. Push the needle of a syringe through the stopper parallel to the hole. Wipe off all traces of glycerin before using.
3. Razor blades must be sharp and free of oil. A straight fresh cut of the Elodea just prior to performing the experiment is important.
4. Oxygen production is dependent on the health of the Elodea. Select dark green, large-leaf sprigs and store in well-aerated aquarium with a high-intensity light source. The plants may be reused in later exercises.
5. Sprigs of Elodea should be placed in the tube upside down with cut end of stem up.

CLASSROOM SUGGESTIONS

1. Students should observe two peaks in the absorption spectrum of chlorophyll (425 nm and 660 nm).
2. To assist the students in determining the light color in table 22.1, a piece of paper can be inserted into an empty cuvette.
3. When assembling the apparatus to measure oxygen production, students should halfway withdraw the plunger of the syringe before the final insertion of the stopper. This will give them space to adjust the levels of the solution before proceeding with the test.
4. If the apparatus for measuring oxygen production is not assembled securely, leaks will develop during the tests. With the Elodea in the tube, fill the tube to the top with the bicarbonate solution. Insert the stopper assemblage to displace the excess solution. Remove the assemblage and dry the top inside of the tube and stopper. Carefully reinsert the stopper and gently twist to insure a tight fit. Too much pressure will jam the assemblage and there is risk of the pipette snapping in the hand and/or the tube top cracking.
5. A minimum homework assignment would be to hand in the graphs on oxygen production by Elodea at different light intensities and the derivative graph of rate of oxygen production versus distance between lamp and plant. The latter defines the light compensation point.
6. An excellent interactive photosynthesis simulation program for Mac and Windows is available from Sunburst Educational Software. It allows students to investigate the effects of light intensity, light spectrum, carbon dioxide concentration, and moisture on photosynthetic rate as measured by a number of parameters. More information is available on the WWW at http://www.nysunburst.com.
7. Check out the links for this lab topic at http://auth.mhhe.com/biosci/genbio/dolphin/ You will find useful materials for developing your lab introduction or summary, and in some cases, you may want to tell students to connect to a particular site for further information.

ANSWERS TO CRITICAL THINKING QUESTIONS

1. Carotenoids, including xanthophyll and carotene, are red, orange, or yellow pigments found in all chloroplasts. They absorb light from below a wavelength of 400nm up to approximately 540nm. Therefore, they can absorb wavelengths of light than chlorophyll cannot, thus broadening the spectrum of light useable by the plant for photosynthesis.
2. Water, itself, absorbs light. The longer wavelengths are absorbed first. Therefore, at depth, the only light penetrating is in the blue-green spectrum. This is the spectrum of light absorbed by the phycobilins, which transfers the energy to chlorophyll for photosynthesis. (Remember, the color we perceive is the light wavelength being reflected, not absorbed.)
3. Basic leaf structure would be the same with epidermis and mesophyll. Differences would be in adaptations to water conservation. Plants from dry environments would be expected to have waxy cuticles, reduced or modified stomate structure, and possibly thicker leaves. Metabolism could also be different with dry adapted plants have some form of C₄ photosynthesis instead of C₃.

SUPPLEMENTAL MATERIALS