This part of the course comprises three practical periods. The experiments will be carried out in rotation by groups as specified in the table below. Each student is allocated to the AM (morning session) or PM (afternoon session) and for all the practicals you will work in groups which will be assigned. A seating chart will be posted in the laboratory.

           You are required to write practical reports for all the experiments. Each report should be handed in to your demonstrator one week after the results for that experiment have been completed. Practical reports will contribute to the final assessment of this course. A ‘zero’ mark will be assigned to those who failed to hand in their practical reports.

Practicals:  1. Photosynthesis & Hill Reaction

2. Water & Salt Relations

3. Water Potential & Transpiration

A.   Demonstration of the Hill reaction

            Illuminated chloroplasts will, in the presence of a suitable hydrogen acceptor, evolve oxygen from water. The Hill reaction may be represented thus:-

4H₂O light 4 [H] + 4 [OH chloroplasts

Hydrogen acceptor + 4 [H] ----------- > Reduced hydrogen acceptor

4 [OH] ------------------- > 2 H₂O + O₂

            In the green plant the usual acceptor is the coenzyme nicotinamide adenine dinucleotide phosphate (NADP). In this experiment you will add an artificial electron acceptor to the suspension of chloroplasts: 2,6-dichlorophenolindolphenol (DPIP). The reaction will be followed by observing the loss of the blue colour of DPIP as it is reduced.

DPIP oxidized + 2 [H]           light                 DP1P.H₂

    chloroplasts
(blue)                                                         (colourless)

Chloroplast Isolation

             The isolation procedure should be carried out at 0 to 4°C. In practice use pre-chilled solutions and containers. Grind to a pulp about 5 g of Chinese spinach leaves (midribs and petioles removed) in a mortar, using initially 10 ml of phosphate buffer extraction medium (0.05 M phosphate buffer pH 6.9 with 0.4 M sucrose and 0.01 M KCl). Add more buffer till 40 ml have been added. Filter the pulp through a layer of muslin and centrifuge the extract at 1000 g for 10 minutes. Pour off the supernatant and resuspend the chloroplast pellet in 10 ml of extraction medium. Keep the chloroplast suspension chilled on ice.

Experiment Procedure

            To each of 3 clean dry test tubes add 5 ml of phosphate buffer, and 1 ml of DPIP. Label the tubes. Take 1 ml portion of the chloroplast preparation and boil in a water bath for 1 minute. Switch on the spectrophotometer at least 5 minutes before taking readings. At zero time add 0.1ml of chloroplast suspension to tubes 1 and 2, and to tube 3 add 0.1 ml of the boiled chloroplast suspension. Mix the contents in each tube and measure the absorbance of each tube at 600 nm against a phosphate buffer blank. Place tubes 1 and 3, 30 cm away from a light bulb and take readings on both of these tubes every minute for the first 5 minutes and then every 5 minutes for a further 25 minutes. Read tube 2 (kept dark by aluminium foil) at the same time intervals. Plot absorbance reading against time. What can you deduce from these results?

             Observe some isolated chloroplasts under the microscope.

             You will be provided with an extract of spinach leaves.

             Spot strips of chromatography paper with the concentrated extract at a position 2 cm from the bottom of the paper strip using a fine capillary tube. Keep the spot to within 0.5 cm in diameter and concentrated (about 20 drops are usually necessary). Blowing on the spot between applications will increase evaporation of the solvent and help to keep the spot small. Suspend the paper strip in a boiling tube containing 1 to 2 ml of the solvent (petroleum ether: acetone mixture, 9 : 1) and stopper. Do not allow the pigment spot to dip into the solvent. Observe over a 10 to 20 minute period.

             How many pigments can you separate? What are their Rf values?

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A. The Münch pressure flow model

             Select a strip of dialysis tubing (about 18 cm long) which has been pre-soaked in water. Twist one end, bend it back, and secure the bend between your fingers. Insert the tapered end of a one-hole rubber stopper into the open end of the dialysis sac and tie the tubing tightly to the stopper so that there are no leaks. Fill the tubing to the top with a 5% (w/v) starch solution. Insert one end of a U-shaped piece of glass tubing through the hole in the topper and into the starch solution. Make sure that the glass tubing fits tightly.

             Select another piece of dialysis tubing and tie off one end as before. In the same manner as before stopper the open end of the dialysis tubing. Fill the tubing to the top with sucrose and I2KI solution. Insert the free end of the U-shaped tube through the stopper and into the sucrose- I2KI solution. Make sure there are no leaks in either of the sacs at each end of the U-shaped tube.

              Support the entire system with a ring stand and clamp so that each sac is immersed completely in a separate beaker of water. Observe the flow of sucrose I2KI solution through the tube. You can try to find out how different concentrations of sucrose- I2KI solutions affect the flow rate.

             1) What is the influence of turgor pressure on the flow of I2KI reagent and sucrose from one sac to the other?

             2) How is the system studied in this experiment analogous to the translocation system of a living plant?

             3) Why is the model studied in this experiment useful as a means of explaining the translocation of osmotically active organic substances within plants?

             4) What are some of the objections to the mechanism of organic solute translocation as  being totally analogous to the Münch pressure flow model?

B. Determination of water potential by the falling drop method

             The water potential of any plant tissue is a measure of the driving force which causes water to move into that tissue. It is possible to determine water potential of tissue by immersing finely cut pieces of the plant in solutions of different osmotic potentials and noting whether the tissue takes up or releases water to the solution. We do this by comparing the densities of solutions in which plants have been immersed with the same solution which has had no plant material added. In the case where there is neither uptake nor release of water, the water potential of the tissue will then be equal to the osmotic potential of the solution.

             You are given a series of test tubes and specimen tubes containing sucrose solution of the following molarities: 0.10, 0.15, 0.20, 0.25, 0.30, 0.35, 0.40, 0.45, 0.50, 0.55; the test series containing 3 ml of solution (in test tubes). the control series about 10 ml of solution (in specimen tubes). Cut some 0.5 ml of the plant tissue provided into small slices and place equal amounts in each of the test series sucrose solutions. Label and stopper the tubes and leave to stand for about an hour, shaking gently from time to time. Then add one drop of methylene blue solution (0.2% w/v in water) to each test tube in the test series and shake. To see if there has been any change in the density of the test series take a small quantity of the bathing solution in a micropipette and release it carefully within the respective control solution series. Watch the subsequent movement of the drop; if it rises the coloured solution has become less dense, i.e. the tissue has lost water to the solution. If the drop falls, the reverse has occurred. The water potential of the tissue will lie between the osmotic potentials of the solutions in which the drop changes direction.

The relation between the concentrations of the sucrose solution and their respective osmotic potentials are given here as:

[Sucrose]        0.10    0.15     0.20   0.25     0.30     0.35    0.40     0.45     0.50     0.55     0.60

O. P. MPa     -0.26  -0.40   -0.53   -0.67   -0.81   -0.96    -1.11   -1.27    -1.43   -1.60   -1.70

A. Determination of water potential by the weighing/measuring method

            The water potential of a cell is a measure of the tendency for water to enter the cell when it is placed in water. The determination rests upon the fact that if the tissue has a lower water potential than distilled water, it will take up water when placed in the latter and hence will increase in weight and volume. On the other hand, if the tissue is placed in a solution which has a lower water potential, then the tissue will lose water and decrease in weight and volume. By using a graded series of sucrose solutions the osmotic potential of that sucrose solution which results in no increase or decrease in the mass or volume of the tissue can be determined as equal to the water potential of the tissue.

            Using a cork borer, punch out 30 cylinders of about 4.0 cm in length from the potatoes provided. Immediately after obtaining a cylinder, trim both ends with vertical cuts to eliminate the suberized peripheral layers. Blot lightly with a paper towel; temporarily store the cylinders in a closed moist chamber lined with moist paper towels.

            After all cylinders have been cut, weight them in groups of three to the nearest 0.001 g. Also measure the length of each cylinder accurately and record the mean length for each group. Immediately after measurement, put each group of cylinders in beakers containing 100 ml of sucrose solutions covering the range 0.15 M to 0.6 M in 0.05 M increments (i.e., you have 10 different concentrations). Cover each beaker with a petri dish; set aside for 2 hours at room temp.

            After the prescribed time, remove the cylinders, one group at a time, from the solutions. Quickly blot the cylinders with a paper towel and measure the final fresh weight and length of cylinders for each group.

            Plot a graph of loss or gain in weight/length against sucrose concentration and determine the molarity of the sucrose solution in which the tissue shows no change in weight/length. Read off the osmotic potential of this sucrose solution from the table given in the schedule for Practical 2b.

            Fill a glass bulb with water and seal the top with parafilm. Remove a leaf from the Coleus plant provided by cutting through the petiole IN WATER, without wetting the leaf. Pierce a small hole in the parafilm; insert leaf such that the cut end of the petiole is immersed in water. Weight the leaf and bulb in a fine balance to 0.0001 g. Work quickly and record the weight. Then place the bulb and leaf under illumination (60W bench lamp) at a distance of about 30 cm; allow leaf to transpire 5 minutes. Weigh the bulb-plus-leaf again. Continue this procedure for 30 minutes in order to obtain a fairly steady rate of transpiration.

            Using a similar procedure and illumination, determine the transpiration rates of the leaf when exposed to room-temp. and warm air currents. Use the blower provided, at a distance which will give the plants wind but not blow them over.

            After the experiment, find the area of the leaf by tracing its outline on a piece of mm² graph paper. Work out the transpiration rates under the various conditions in terms of weight of water transpired per cm². of leaf surface (one side) per minute.

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