Biology Practical Experiment Procedures - Color Coded Summary

Color Key:

● Red = Independent Variable (the factor you deliberately change)

● Blue = Dependent Variable (what you measure as the result)

● Purple = Controlled Variable (conditions you keep constant)

● Green = Safety Steps (precautions and safety procedures)

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Practical 5.1: Investigating Mitosis in Root Tips

  1. Place onion root tips in a boiling tube containing dilute hydrochloric acid
  2. Cover the tube with a cap and leave for several minutes to soften the tissue
  3. Pour out the acid and transfer root tips to a watchglass
  4. Rinse the root tips thoroughly under running water to remove all acid
  5. Use tweezers to carefully separate the root tip from the main root section
  6. Place the root tip on a clean microscope slide
  7. Add a drop of water or mounting medium to the slide
  8. Gently place a cover slip over the root tip
  9. Press down firmly on the cover slip to squash the tissue (use lens paper to absorb excess liquid)
  10. Observe under the low-power objective lens first to locate mitotic cells
  11. Switch to the high-power objective lens to examine individual cells
  12. Identify different mitotic stages: prophase, metaphase, anaphase, and telophase
  13. Look for chromosomes, spindle fibers, and nuclear envelope changes
  14. Count cells in different stages and calculate the percentage in the mitotic arc
  15. Record observations with labeled drawings showing each distinct stage

Practical 6.1: Extracting DNA from Onion Bulb Cells

  1. Cut onion bulbs into small pieces using a scalpel or knife
  2. Transfer the cut onion pieces into a clean beaker
  3. Add approximately 100 mL of detergent solution to the beaker
  4. Add salt water to the beaker with the onion pieces and detergent
  5. Place the beaker in a water bath at 60°C for 15 minutes
  6. Allow the mixture to cool to room temperature by stirring gently
  7. Pour the cooled mixture into a clean beaker through a funnel or strainer
  8. Carefully add ice-cold ethanol layer by tilting the beaker and pouring along the side
  9. Allow the ethanol to sit on top of the mixture without stirring (watch for white precipitate forming)
  10. Observe the stringy white DNA precipitate forming at the interface between liquids
  11. Use a glass rod or stirring stick to wind the DNA strands gently onto the rod
  12. Carefully remove the rod with extracted DNA from the beaker
  13. Hold the rod with extracted DNA over a watch glass to collect any drips
  14. Observe the visible white DNA threads hanging from the rod
  15. Document the appearance and amount of extracted DNA material

Practical 7.1: Rates of Diffusion in 'Cells' of Different Sizes

  1. Obtain several agar jelly cubes in a petri dish from the stock solution
  2. Cut or identify cubes of at least 4-5 different sizes (varying heights from 1 cm to 5 cm)
  3. Prepare a dye solution by adding methylene blue to a beaker of water
  4. Record the starting time on a clock or timer
  5. Place the first (smallest) agar cube into the dye solution
  6. Place each remaining cube into the dye solution at 1-minute intervals
  7. Observe color changes as the dye penetrates each cube over time
  8. At regular time intervals (every 2-3 minutes), remove one cube from the solution
  9. Rinse the cube gently under running water to remove surface dye
  10. Measure the depth of dye penetration into the cube using a ruler in millimeters
  11. Cut the cube in half to measure internal penetration more accurately if needed
  12. Record measurements for each cube size in a data table
  13. Calculate the total surface area for each cube using the formula SA = 6s²
  14. Calculate the volume for each cube using the formula V = s³
  15. Determine the surface area-to-volume ratio (SA:V) for each cube
  16. Plot a graph showing diffusion time against SA:V ratio
  17. Analyze the relationship and explain why smaller cells exchange substances more efficiently

Practical 7.4: Investigating the Rate of Transpiration of a Leafy Shoot Using a Potometer

  1. Attach a capillary tubing syringe filled with distilled water to the potometer apparatus
  2. Ensure an airtight seal is created between the tubing and the potometer
  3. Submerge the leafy shoot end in a beaker of distilled water
  4. Cut the stem at a 45-degree angle under water using sharp scissors
  5. Quickly transfer the cut stem to the potometer, inserting it into the capillary tubing
  6. Ensure the xylem vessels fill with water and no air bubbles are trapped
  7. Submerge the entire potometer apparatus in a beaker of water
  8. Remove the leafy shoot from the beaker and ensure the cut end seals against the capillary
  9. Remove the leafy shoot and attach the capillary tubing vertically using a clamp
  10. Place a drop of colored oil into the tubing to act as a visible meniscus marker
  11. Record the initial position of the oil drop using a ruler as the reference point
  12. Measure the distance moved by the water meniscus at regular time intervals (every 5-10 minutes)
  13. Record environmental conditions: temperature, humidity, and light intensity
  14. Calculate the rate of transpiration using: r = change in length (cm) × cross-sectional area of capillary
  15. Repeat the experiment with different variables (different leaves, temperatures, or humidity levels)
  16. Compare results to identify factors affecting transpiration rates

Practical 16.1: Investigating the Effect of Temperature on Dehydrogenase Activity in Yeast

  1. Use syringes to place 10 mL of yeast suspension into a clean boiling tube
  2. Add 1 mL of distilled water to each tube as a dilution
  3. Prepare a water bath at room temperature (approximately 20°C)
  4. Record the exact temperature of the water bath using a thermometer
  5. Place the yeast tubes into the water bath for several minutes to allow equilibration
  6. Add 1 mL of methylene blue substrate solution to each tube
  7. Mix the tube contents gently by swirling (do not shake vigorously)
  8. Measure the initial color intensity using a colorimeter or spectrophotometer
  9. Record the time taken for the dye color to change from blue to clear
  10. Repeat steps 1-9 at different temperatures: 30°C, 40°C, 50°C, and 60°C
  11. Use a thermometer to monitor and maintain each water bath at the target temperature
  12. For each temperature, record the time taken for color change and enzyme activity level
  13. Create a data table showing time and activity for each temperature tested
  14. Plot results on a graph with temperature on the x-axis and enzyme activity on the y-axis
  15. Identify the optimal temperature showing maximum dehydrogenase enzyme activity
  16. Explain the effects of temperature on enzyme function and protein denaturation

Practical 16.2: Investigating the Rate of Respiration of Small Organisms Using a Simple Respirometer

  1. Select several small living organisms (maggots, small worms, or germinating seeds)
  2. Place the organisms in the bottom of a sealed syringe or closed container
  3. Add absorbent material (calcium hydroxide solution or soda lime) to trap carbon dioxide produced
  4. Ensure the container is completely sealed with no air leaks
  5. Attach a water-filled capillary tube to the sealed container using a connector
  6. Record the initial position of the water meniscus in the capillary tube
  7. As organisms respire, oxygen is consumed from the enclosed space
  8. Water rises up the capillary tube as atmospheric pressure pushes it inward
  9. Measure the position of the water meniscus at regular time intervals (every 5 minutes)
  10. Record all measurements in a data table with time and distance values
  11. Calculate the volume of oxygen used: Volume = distance moved × cross-sectional area of capillary
  12. Set up a control container with identical setup but no organisms
  13. Record control measurements to account for any physical changes unrelated to respiration
  14. Subtract control values from experimental values to get net respiration rate
  15. Calculate oxygen consumption rates per organism or per unit time
  16. Compare respiratory rates between different organism types or environmental conditions

Practical 17.1: Investigating Pigments in a Leaf Using Paper Chromatography

  1. Tear fresh green leaves into small pieces
  2. Place the leaf pieces in a clean mortar
  3. Add a small amount of sand to the mortar (to help break down cell walls)
  4. Add calcium carbonate (chalk) to the sand and leaves
  5. Add a few mL of organic solvent (petroleum ether or ethanol) to create a paste
  6. Use a pestle to grind the mixture thoroughly until the leaves become pulpy
  7. Continue grinding until pigment colors are clearly visible in the solution
  8. Pour the pigment extract carefully into a small beaker or test tube
  9. Cut a strip of chromatography paper to the appropriate length
  10. Draw a pencil baseline 2 cm from the bottom of the paper strip
  11. Use a pencil capillary to place a small drop of pigment extract on the baseline
  12. Allow the spot to dry completely
  13. Repeat applications (2-3 times) to concentrate the pigment, allowing drying between applications
  14. Pour solvent into a chromatography tank to a depth of about 1 cm
  15. Suspend the paper strip vertically in the tank with the baseline above the solvent level
  16. Cover the tank to prevent solvent evaporation
  17. Allow the solvent to rise through the paper via capillary action
  18. Stop the experiment when the solvent front reaches near the top (about 1-2 cm from the top)
  19. Remove the paper and immediately mark the solvent front with a pencil
  20. Allow the paper to dry completely
  21. Measure the distance traveled by each pigment color from the baseline
  22. Measure the total distance traveled by the solvent front
  23. Calculate the Rf value for each pigment: Rf = distance of pigment ÷ distance of solvent
  24. Identify each pigment based on known Rf values (chlorophyll a, chlorophyll b, carotene, xanthophyll)

Practical 17.2: Investigating the Light-Dependent Stage of Photosynthesis

  1. Prepare fresh large leaves from plants such as variegated species or Swiss chard
  2. Cut leaves into small pieces using a scalpel
  3. Place the leaf pieces in a clean mortar with sand
  4. Add isolation medium solution to the mortar
  5. Grind the leaves thoroughly with a pestle until a chlorophyll suspension forms
  6. Pour the green suspension into a beaker
  7. Prepare several cuvettes (small clear containers for spectrophotometer use)
  8. Using a dropping pipette, add drops of chlorophyll isolation medium to each cuvette
  9. Add drops of the leaf extract to each cuvette until color is visible
  10. Seal each cuvette with a cap to prevent evaporation
  11. Place one cuvette in a spectrophotometer as a blank (control with no leaf extract)
  12. Measure baseline light absorption using the spectrophotometer at 400 nm (blue light)
  13. Expose experimental cuvettes to different light wavelengths (red, green, blue, yellow, UV)
  14. For each wavelength, shine light of that color onto the cuvette for a set time period
  15. Measure light absorption immediately after exposure using the spectrophotometer
  16. Record the intensity of color changes for each wavelength
  17. Repeat measurements for each wavelength to ensure accuracy
  18. Create a graph showing absorption (y-axis) against wavelength (x-axis)
  19. Identify which wavelengths produce the greatest changes in absorption
  20. Determine which colors are most effective for photosynthetic light-dependent reactions

Practical 17.3: Investigating the Effect of Carbon Dioxide Concentration on the Rate of Photosynthesis

  1. Collect fresh pieces of Canadian pondweed (Elodea canadensis)
  2. Measure the initial length of the plant using a ruler
  3. Place pondweed in experimental tubes filled with water
  4. Prepare three different isolation medium solutions: 11 mL acid, 11 mL base, 11 mL sodium hydrogencarbonate
  5. Set up apparatus with transparent plastic tubing containing pondweed submerged in water
  6. Connect the apparatus with the labeled isolation medium solutions in sequence
  7. Place the apparatus vertically using a clamp and stand
  8. Connect a capillary tube to the end of the apparatus to measure gas movement
  9. Fill the capillary tube completely with water initially
  10. Gently push down the plunger to force water into the capillary tubing
  11. Record the initial position of the water meniscus in the capillary tube
  12. Measure the distance the water meniscus moves at regular time intervals (every 5 minutes)
  13. Record movement over a 15-30 minute observation period
  14. The meniscus movement indicates oxygen production rate and photosynthesis rate
  15. Repeat the entire procedure using each of the three isolation medium solutions
  16. Calculate the rate of photosynthesis for each carbon dioxide concentration
  17. Create a data table showing CO₂ concentration, time, and distance moved
  18. Plot results on a graph with CO₂ concentration on x-axis and photosynthesis rate on y-axis
  19. Identify the point where carbon dioxide becomes saturating
  20. Analyze the relationship between CO₂ concentration and photosynthetic rate

Practical 20.1: Measuring Population Growth in a Culture of Chlorella

  1. Obtain a hemocytometer slide and place it under a light microscope
  2. Examine the gridded squares on the hemocytometer
  3. Select the two-pointer objective lens of the microscope
  4. Place a small sample of Chlorella culture on the hemocytometer
  5. Place the coverslip carefully on top to create a uniform depth
  6. Focus on one of the gridded squares using low power first
  7. Switch to high power and count cells within defined grid squares
  8. Count cells in at least 5 different squares to get accurate average counts
  9. Follow the standard counting rule: count cells in top-left to center, avoid over-counting
  10. Calculate the average number of cells per square
  11. Multiply by the appropriate dilution factor (usually 10⁴) to get cells per mL
  12. Record this as the initial cell count at day 0
  13. Prepare culture flasks containing nutrient growth medium
  14. Inoculate each flask with Chlorella cells at the calculated initial density
  15. Place flasks under controlled light conditions with lamps providing illumination
  16. Maintain constant temperature (approximately 20-25°C)
  17. Ensure gentle aeration of the cultures
  18. At the same time each day for 12-14 days, remove a sample from the culture
  19. Use the hemocytometer to count cells in the daily sample
  20. Record cell densities in a data table with date and cell count
  21. Plot cell number (y-axis) against time in days (x-axis) on a graph
  22. Analyze the curve shape to identify growth phases: lag phase, exponential phase, stationary phase
  23. Calculate generation time using the exponential growth data
  24. Calculate specific growth rates
  25. Write conclusions about population dynamics and growth patterns observed