1. Investigation: Light is Necessary for Photosynthesis
Procedure: Place a potted plant in darkness for 24 hours to destarch it. Test one leaf for starch to confirm destarching is complete.
Cover part of both sides of a leaf using black tape or aluminum foil. Leave the plant in sunlight for 5 hours.
Remove the leaf and foil, then test the leaf for starch using iodine solution.
Expected Results: Areas that received light turn blue-black with iodine, showing photosynthesis occurred. Covered parts remain brown, indicating no photosynthesis took place.
Conclusion: The results demonstrate that light is necessary for photosynthesis.
2. Investigation: Carbon Dioxide is Necessary for Photosynthesis
Procedure: Place the plant in darkness for 24 hours to destarch it. Select two leaves on the plant.
Place one leaf in a flask containing potassium hydroxide solution (absorbs CO₂). Place another leaf in a flask containing sodium bicarbonate solution (produces extra CO₂).
Keep both leaves attached to the plant and place in sunlight for 5 hours. Remove both leaves and test for starch.
Expected Results: The leaf from potassium hydroxide flask remains brown (no starch present). The leaf from sodium bicarbonate flask turns blue-black (starch present).
Conclusion: The results indicate that carbon dioxide is necessary for photosynthesis.
3. Investigation: Chlorophyll is Necessary for Photosynthesis
Procedure: Take a potted plant with variegated leaves and place in darkness for 24 hours to destarch. Place the plant in sunlight for 5 hours.
Remove one variegated leaf and test for starch using iodine solution.
Expected Results: Green parts of the leaf turn blue-black, indicating starch presence and photosynthesis. White parts turn brown, showing no starch formation due to lack of chlorophyll.
Conclusion: The results demonstrate that chlorophyll is necessary for photosynthesis.
4. Testing for Starch in a Leaf
Procedure: Pluck a fresh leaf and place in boiling water for few minutes to kill cells and break the cuticle. Boil the leaf in ethanol using a water bath to remove chlorophyll (ethanol is flammable).
Place the leaf in warm water briefly to wash off ethanol and soften the brittle leaf. Spread the leaf on a white tile and add few drops of iodine solution.
Expected Results: Dark blue color indicates starch presence in the leaf. Brown color (iodine’s natural color) indicates absence of starch.
5. Investigation: Green Leaves Contain Different Pigments
Procedure: Collect and crush fresh green leaves using pestle and mortar. Place crushed leaves in a small beaker and add ethanol to half-fill.
Create a water bath by placing the small beaker in a larger beaker with water. Heat until alcohol turns green, showing dissolved pigments.
Filter the green alcohol mixture into another beaker. Cut filter paper strip (3cm × 10cm) and attach to glass rod with tape.
Suspend the paper strip so its tip touches the colored ethanol for 30 minutes.
Expected Results: Three distinct colors appear on the strip from bottom: green (chlorophyll), orange (carotene), and yellow (xanthophyll).
Conclusion: Fresh green leaves contain different types of pigments as demonstrated by chromatography.
6. Estimating Grasshopper Population in School Playground
Procedure: Use a sweep net to capture grasshoppers, mark and count them, then release back. After one day, capture grasshoppers again from the same area.
Count total grasshoppers in second catch and count marked individuals separately.
Calculation: Population = (First catch × Second catch) ÷ Marked individuals in second catch
7. Estimating Population Density of Grasshoppers
Procedure: Follow the same capture-mark-recapture method as above. Calculate total population using the formula: (First catch × Second catch) ÷ Marked individuals in second catch.
Measure the total area of the school playground in square meters.
Calculation: Population density = Total population ÷ Area of playground
8. Estimating Population of Tridax procumbens
Procedure: Identify the field containing the plant and estimate its total area in square meters. Throw a quadrat randomly into the selected area multiple times.
Count and record Tridax procumbens individuals in each quadrat throw. Calculate the average number of organisms per quadrat.
Calculation: Total population = (Average organisms per quadrat × Total field area) ÷ Quadrat area
Note: Population density = Total population ÷ Total field area
9. Investigation: Effect of Wind Speed on Transpiration Rate
Procedure: Cut two leafy shoots and fix them to potometers, ensuring all cutting and fixing is done underwater. This prevents air bubbles from being trapped in the apparatus which would affect accurate readings.
Place both shoots in identical environmental conditions with similar temperature and lighting. Position a fan close to one shoot and switch it on to increase air movement around the leaves.
The fan creates air currents that reduce humidity around the leaf surface by blowing away water vapor. Leave both setups for several hours and record transpiration rates from the potometer readings.
Expected Results: The shoot exposed to the fan shows significantly higher transpiration rates compared to the control shoot. The moving air prevents water vapor accumulation around the leaf surface, maintaining a steep concentration gradient.
This lower humidity around the leaves accelerates the diffusion of water vapor from the leaf interior to the external environment.
Conclusion: Wind speed directly affects transpiration rate – higher wind speeds increase transpiration by reducing humidity around leaves.
10. Investigation: Effect of Humidity on Transpiration Rate
Procedure: Cut and fix two leafy shoots to potometers, ensuring underwater cutting and fixing to prevent air bubble formation. Cover one shoot completely with transparent plastic paper, creating a sealed environment around the leaves.
Leave the second shoot uncovered as a control. Place both setups in identical environmental conditions with same temperature and light intensity.
Monitor both setups for several hours and record transpiration rate measurements from the potometer scales.
Expected Results: The covered shoot demonstrates significantly lower transpiration rates than the uncovered control shoot. The plastic covering traps water vapor evaporating from the leaves, creating high humidity conditions around the leaf surface.
This high humidity reduces the concentration gradient between the leaf interior and external environment. The uncovered shoot maintains normal transpiration due to natural air circulation removing water vapor.
Conclusion: Humidity inversely affects transpiration rate – high humidity reduces transpiration while low humidity increases it.
11. Investigation: Effect of Temperature on Transpiration Rate
Procedure: Cut and fix two identical leafy shoots to potometers using underwater cutting and fixing techniques. Position one shoot in a shaded area with cooler temperatures while placing the second shoot under direct hot sunlight.
Ensure both locations have similar humidity and wind conditions to isolate temperature as the only variable. Monitor and record transpiration rates from both potometers over several hours.
The temperature difference between shade and direct sunlight creates distinct thermal environments for comparison.
Expected Results: The shoot in direct sunlight registers markedly higher transpiration rates than the shaded shoot. Higher temperatures increase the kinetic energy of water molecules in leaf cells, accelerating evaporation processes.
Heat also increases the water-holding capacity of air around the leaves, creating steeper concentration gradients. The shaded shoot maintains lower, more consistent transpiration rates due to cooler temperatures.
Conclusion: Temperature positively correlates with transpiration rate – higher temperatures significantly increase water loss from plant leaves.
12. Investigation: Effect of Light Intensity on Transpiration Rate
Procedure: Cut and fix two leafy shoots to potometers using proper underwater techniques. Set up both shoots in a controlled room environment with an electric bulb as the light source.
Position one shoot directly under the bulb to receive maximum light intensity. Place the second shoot at a greater distance from the bulb to receive reduced light intensity while maintaining the same room conditions.
Allow both setups to run for several hours and carefully record transpiration rate measurements.
Expected Results: The shoot under direct light shows higher transpiration rates compared to the shoot with lower light exposure. Higher light intensity triggers greater stomatal opening as plants prepare for increased photosynthetic activity.
More open stomata create larger surface areas for water vapor diffusion, accelerating transpiration. The shoot with reduced light maintains more closed stomata, resulting in lower transpiration rates.
Conclusion: Light intensity directly influences transpiration rate through its effect on stomatal behavior – brighter light increases transpiration.
13. Investigation: Effect of Temperature on Pepsin Digestion of Milk
Procedure: Prepare three labeled test tubes (A, B, C) and add equal samples of coagulated milk to each. Add 1cm³ of pepsin enzyme solution to all three test tubes to ensure equal enzyme concentration.
Place test tube A in a water bath maintained at 37°C, simulating optimal human body temperature conditions. Place test tube B in cold water or refrigerate it to create low-temperature conditions.
Boil the contents of test tube C to create high-temperature conditions that may denature the enzyme. After appropriate incubation periods, test samples from each tube for remaining protein content.
Expected Results: Test tube A shows complete protein digestion with negative protein tests, demonstrating pepsin’s optimal activity at body temperature. Test tube B retains high protein levels as low temperatures slow enzyme activity and reduce molecular collision rates.
Test tube C also shows high protein levels because excessive heat denatures pepsin’s protein structure, permanently destroying its catalytic activity. The enzyme cannot function after structural denaturation occurs.
Conclusion: Temperature critically affects pepsin activity – optimal temperatures enable digestion while extreme temperatures inhibit or destroy enzyme function.
14. Investigation: Effect of pH on Amylase Digestion of Starch
Procedure: Label three test tubes A, B, and C, then add 1cm³ of starch solution to each tube. Create different pH environments by adding 3 drops of hydrochloric acid to tube B (acidic conditions) and several drops of sodium hydroxide to tube C (alkaline conditions).
Leave tube A without pH modifiers as a neutral control. Add 1cm³ of amylase enzyme solution to tubes B and C only, maintaining the control without enzyme.
Allow all setups to incubate for 20 minutes at room temperature, then test each tube’s contents for remaining starch using iodine solution.
Expected Results: Test tube A turns blue-black, confirming starch presence since no enzyme was added. Test tube B also turns blue-black despite enzyme presence because amylase cannot function in highly acidic conditions.
Test tube C shows brown coloration, indicating successful starch breakdown because amylase functions optimally in alkaline pH conditions. The enzyme’s active site maintains proper configuration in alkaline environments.
Conclusion: pH significantly affects amylase enzyme activity – alkaline conditions promote starch digestion while acidic conditions inhibit it.
15. Investigation: Effect of Particle Size on Enzyme Activity
Procedure: Cut three identical liver pieces into 2cm × 2cm × 2cm cubes and prepare three test tubes with 10ml hydrogen peroxide each. Label the tubes A, B, and C, then prepare different particle sizes from the liver cubes.
Keep one cube whole for tube A. Cut the second cube into four equal pieces for tube B. Crush the third cube using pestle and mortar until it forms fine particles for tube C.
Prepare a stopwatch and systematically add each liver preparation to its respective tube. Record the time required for foam production to reach the test tube mouth in each case.
Expected Results: Test tube A with the whole liver cube produces foam slowly, taking the longest time to reach the tube mouth. Test tube B with cut pieces produces foam faster than A due to increased surface area exposure.
Test tube C with crushed liver produces foam most rapidly, reaching the tube mouth fastest. The catalase enzyme in liver tissues has maximum exposure to hydrogen peroxide when particle size is smallest, creating optimal reaction conditions.
Conclusion: Particle size inversely affects enzyme activity – smaller particles provide greater surface area, significantly increasing reaction rates.
16. Investigation: Presence of Catalase Enzyme in Liver Tissue
Procedure: Cut a fresh liver piece and crush it thoroughly into small fragments to maximize surface area exposure. Prepare two test tubes with 50ml of hydrogen peroxide solution each for comparative analysis.
Add the crushed liver pieces to only one test tube, leaving the second tube as a control containing only hydrogen peroxide. Observe both setups carefully for several minutes, noting any visible reactions or changes.
The setup allows direct comparison between enzyme-catalyzed and non-catalyzed hydrogen peroxide breakdown.
Expected Results: The test tube containing liver pieces shows immediate bubbling and vigorous foam formation that rises within the tube. This reaction occurs because catalase enzyme in liver tissue rapidly breaks down hydrogen peroxide into water and oxygen gas.
The oxygen gas release creates visible bubbling and foam formation. The control tube with only hydrogen peroxide shows no bubbling or foam, confirming that the reaction requires the enzyme present in liver tissue.
Conclusion: Liver tissue contains catalase enzyme, which effectively breaks down hydrogen peroxide into harmless water and oxygen products.