SB015 Lab Manual Updated: 26/04/2022 25 Hairline shape Widow’s peak hairline https://www.healthline.com/health/wid ows-peak Straight hairline https://www.ishrs-htforum.org Chin shape Cleft chin https://www.healthline.com/health/cleft -chin Smooth chin https://www.mentalfloss.com/ Thumb bending Normal thumb bending https://askabiologist.asu.edu Hitchhiker’s thumb https://askabiologist.asu.edu Figure 5.1: The six inherited characteristics in human
SB015 Lab Manual Updated: 15/09/2021 26 Results Table 5.1 Observed and expected frequencies of each genotype for six characteristics in the class Characteristic Phenotype Genotype Tick (√) your own genotype Observed frequency of each phenotype in the class Earlobe Free earlobe P_ Attached earlobe pp Tongue Rolling Ability of tongue rolling into “U” shape C_ Inability of tongue rolling into “U” shape cc Dimple Have dimple D_ Without dimple dd Hairline shape Widow’s peak hairline H_ Straight hairline hh Chin shape Cleft chin S_ Smooth chin ss Thumb Normal thumbbending T_ Ability to bend thumb at 60° angle or more tt
SB015 Lab Manual Updated: 26/04/2022 27 Experiment 5.2: ABO blood group inheritance Apparatus: Depression slide/Pallet Lancing device Materials: Anti-A and Anti-B serum/ Blood test kit Alcohol swab Cotton ball/ Sterile gauze Sterilized lancet Toothpicks Safety and laboratory precaution 1. DO NOT use same lancet twice or exposed lancet. 2. Dispose all contaminated supply in a given bin. 3. Use different toothpick to mix blood and serum to prevent any inaccuracy. Procedures and Observation: 1. Label two clean and dry slides/ pallet (no. 1 and 2). 2. Wash your hands with soap and let them dry. 3. Swing your hand facing downward for 10 – 15 seconds. 4. Apply alcohol to your middle finger. Firmly place a new sterile lancet off-center on the fingertip. 5. Press the lancet to puncture the fingertip.
SB015 Lab Manual Updated: 15/09/2021 28 Figure 5.2 Guidance for fingerprick (Source: https://www.who.int/diagnostics_laboratory/documents /guidance/fingerprick.pdf?ua=1) 6. Wipe off the first blood drop with sterile gauze or cotton ball. 7. Place the next drop at the center of slide 1 and 2. (Note: Apply gauze or cotton ball to the punctured site until bleeding stops.) 8. Drop an Anti-A serum near the blood on slide 1 and Anti-B serum on slide 2. 9. Mix the blood and serum on slide 1 with a toothpick. Use another toothpick for slide 2. (Note: You belong to A blood group if agglutination occurs on slide 1 only; B blood group if agglutination is observed on slide 2 only; AB blood group if agglutination occurs on both slides 1 and 2; O blood group if no agglutination is seen on both slides). Figure 5.3 Identification of blood type
SB015 Lab Manual Updated: 26/04/2022 29 10. Identify your blood group. Your blood group: 11. Calculate the phenotype frequency of each blood group in the class. Record your observation in Table 5.3. Table 5.3 Phenotype Frequency of blood group in the class Blood group Possible genotypes Phenotype frequency of each blood group A B AB O Questions: Part A: Inheritance of genetic traits in human 1. Individuals with certain heterozygote characteristics are usually called a carrier. What does a carrier mean? 2. A student inherited straight hairline from parents who are both has widow’s peak hairline. Explain the pattern of inheritance. Part B: ABO blood group inheritance 1. Why do you swing your hand for 10 to15 seconds before pricking the tip of your middle finger? 2. Why can’t you use the same lancet twice? 3. Why do you need to wipe off the first blood drop? 4. Why do you need different toothpicks to mix the blood and serum on slides 1 and 2? 5. Can an individual with O blood group donates his blood to an A blood group person? Give reason to your answer.
SB015 Lab Manual Updated: 15/09/2021 30 6. A mother with O blood group gave birth to a baby girl having the same blood group. However, she is not convinced that the baby belongs to her because her husband has AB blood group. She claimed there might be swapping of baby in the nursery. With the aid of genetic diagram, explain your answer.
SB015 Lab Manual Updated: 26/04/2022 31 EXPERIMENT 6: ISOLATING DNA Objectives: At the end of this lesson, students should be able to isolate DNA from plant tissue. Introduction: Each chromosome is a single thread-like structure made up of long molecules of DNA combined with histone protein. The DNA molecule is made up of many small sections called genes. Shortly before cell division occurs, each DNA molecule replicates itself. So, one thread of the chromosome becomes two identical chromatids. As the two chromatids are identical, they will have identical genes. These identical genes are known as allele. In this experiment, you will rupture fruit cells, thus releasing their contents such as protein, DNA, RNA, lipids, ribosomes and various small molecules. DNA is then suspended by alcohol as supernatant layer. The purity of DNA will require further steps. After the isolation of nucleic acids, the solution is still contaminated with proteins which can be removed. To check the success of the removal, a purity determination is performed, which is based on the different absorption characteristics of the proteins and the nucleic acids using UV spectrophotometer. Apparatus: 500 ml beaker Boiling tube Boiling tube rack Mortar and pestle Muslin cloth Water bath (60°C)
SB015 Lab Manual Updated: 15/09/2021 32 Materials: Ice-chilled 95% alcohol Ice cubes Kiwi/ banana/onion/tomato/watermelon 0.292 g ethylenediaminetetraacetic acid (EDTA) 50.000 g sodium dodecyl sulphate or sodium lauryl sulphate (SDS or SLS) 8.770 g sodium chloride 4.410 g sodium citrate 1 litre water Procedures and Observation: 1. Prepare the salt-detergent solution. Stir gently to completely dissolve the salt without producing foam. 2. Pour 10 ml of ice-chilled alcohol into a boiling tube and place it into a beaker containing ice cubes. (Remarks: place the ethanol in the freezer overnight) 3. Peel, slice and mash kiwi/ onion/ tomato/ banana/ watermelon with mortar. 4. Transfer mashed fruit into a beaker and add 100 ml of the salt-detergent solution. Incubate the mixture in the water bath at 60oC for 15 minutes. 5. After 15 minutes, sieve the mixture with muslin cloth and collect the liquid in a beaker. 6. Fill in half of the boiling tube with sieved liquid. 7. Very carefully pour 10ml of ice-chilled alcohol into the side of the boiling tube at a flat angle (Figure 6.1). (Precaution: make sure both liquids do not mix and alcohol form a separate layer on top of the sieved liquid). saltdetergent solution
SB015 Lab Manual Updated: 26/04/2022 33 Figure 6.1 Pour ice chilled alcohol in a flat angle 8. Put the boiling tube into a rack and observe it. Observe and draw the extracted DNA between alcohol and the sieved liquid. Crude DNA should be found in between the alcohol and sieved liquid. Questions: 1. What is the purpose of using the following? a. salt-detergent solution b. ice chilled alcohol c. water bath at 60oC 2. Why do we need to mash the fruits?
BIOLOGY 2 SB025
SB025 Lab Manual Updated: 15/09/2021 36 EXPERIMENT 7: DIVERSITY OF BACTERIA Objectives: At the end of this lesson, students should be able to: i. demonstrate Gram staining technique in classifying bacteria; ii. identify Gram-positive and Gram-negative bacteria; and iii. identify different shapes of bacteria. Introduction: Gram stain is a widely used method of staining bacteria as an aid to their identification. It was originally devised by Hans Christian Joachim Gram, a Danish doctor. Gram stain differentiates two major cell wall types. Bacterial species with walls containing small amount of peptidoglycan and characteristically, lipopolysaccharide, are Gram-negative whereas bacteria with walls containing relatively large amount of peptidoglycan and no lipopolysaccharide are Gram-positive. Apart from Gram staining technique, the identification of bacteria can also be based on shapes. The three most common shapes are spheres, rods and spirals. Apparatus: Inoculation loops Bunsen burner Compound microscopes Forceps Petri dish Staining racks Slides Materials: Crystal violet Cultures of Escherichia coli Cultures of Staphylococcus aureus 95% ethanol Filter paper Immersion oil Iodine Labelling stickers Prepared slides of different types of bacteria Safranin Yoghurt (diluted in water 1:10)
SB015 Lab Manual Updated: 26/04/2022 37 Figure 7.1 Comparative staining and cell wall structures of Gram-positive and Gram-negative bacteria. (Adapted from www.quia.com) Figure 7.2 Gram staining of bacteria (Adapted from http://enfo.agt.bme.hu/drupal/node/9460)
SB025 Lab Manual Updated: 15/09/2021 38 Figure 7.3 Different shapes of bacteria (Adapted from commons.wikimedia.org/wiki/File:OSC_Microbio_03_03_ProkTable.jpg)
SB015 Lab Manual Updated: 26/04/2022 39 Safety Precaution: 1. Wash hands and wear gloves and mask throughout the experiment. 2. Sterilize your workstation with alcohol or any disinfectant liquid. 3. Place a spirit lamp/ Bunsen burner at your workstation. This experiment must be conducted in sterilized environment. 4. DO NOT wash used slide. Place the slide in the basin given. 5. Dispose all contaminated material into a proper bin. Procedures and Observation: 1. Put a slide into the petri dish. Pour 95% alcohol and soak for about 30 seconds. Then use forceps to take out the slide. Let the slide dry and heat it by placing above the flame. 2. Heat the inoculation loop until it glows red to sterilize the loop. 3. Place a loop of sterile distilled water on the slide. 4. Heat the loop again and cool down the loop on the agar of the bacterial culture. Then, transfer a minimal amount of bacterial colony on to slide. 5. Gently heat the slide to fix the bacteria onto the slide. 6. Place the slide on the staining rack. Cover the smear with a few drops of crystal violet and wait for 30 seconds to 1 minute. 7. Gently, rinse the slide with slow running tap water. 8. Cover smear with 2 drops of iodine. Rotate and tilt the slides to allow the iodine to drain. Then, cover again with iodine for 30 seconds to 1 minute. Since the iodine does not mix well with water, this procedure ensures that the iodine will be in contact with the cell walls of the bacteria on the slide. 9. Rinse the slide with water as in step 7.
SB025 Lab Manual Updated: 15/09/2021 40 10. Place several drops of 95% alcohol (decolouriser) evenly over the smears, rotate and tilt the slide. Continue to add alcohol until most of the excess stain is removed and the alcohol running from the slide appears clear. Although there is no recommended time for this step, it usually takes 10-20 seconds to decolourise if exposed to a sufficient amount of decolouriser. (This is the most critical step of the procedures! If the smears are too thick, or if the alcohol is kept on the slide for too long or too short a time, the results will not be accurate. If the alcohol remains on the smear too long, it may also decolourise Gram-positive bacteria.) 11. Add few drops of safranin on the bacterial smear and leave it for approximately 30-45 seconds. (Note: If the bacteria are Gram-positive, it will retain the primary stain (crystal violet) and not take the secondary stain (safranin), causing it to look violet/ purple under a microscope. If the bacteria are Gramnegative, it will lose the primary stain and take the secondary stain, causing it to appear red when viewed under a microscope). 12. Rinse off with water and blot dry with filter paper. 13. Observe the slide under a microscope using oil immersion objective lens. Record your observation in Table 7.1. 14. Repeat steps 1 to 13 for bacteria found in yoghurt.
SB015 Lab Manual Updated: 26/04/2022 41 Table 7.1 Observation results on the type of bacteria, shape, colour and Gram-positive/ Gram-negative Bacteria Shape Colour Gram-positive/ Gram-negative E. coli S. aureus Bacteria from yoghurt Questions: 1. Why Gram-positive bacteria purple in colour while Gram-negative are red? 2. List two examples each of beneficial and harmful Gram-positive bacteria and Gram-negative bacteria. 3. If the iodine step were omitted in the Gram-staining procedure, what colour of stain would you expect from Gram-positive and Gramnegative bacteria?
SB025 Lab Manual Updated: 15/09/2021 42 Objectives: At the end of this lesson, students should be able to: i. observe the diversity of species in bryophytes and pteridophytes; and ii. construct scientific drawing of bryophytes and pteridophytes. Introduction: Bryophytes and pteridophytes are two large groups of spores producing terrestrial plants. Compared to the flowering plants, they have a longer history of evolution. Bryophytes There are three main divisions of bryophytes, namely Bryophyta (mosses), Hepatophyta (liverworts), and Anthocerophyta (hornworts). Bryophytes are the most primitive among the terrestrial plants. They are nonvascular and are confined to moist areas because they lack well developed tissues for transporting water and nutrients. Bryophytes have a root-like structure, which is called rhizoid and have no true stem and leaves. Bryophytes are characterized by clear alternation of generation in its life cycle where the gametophyte generation is dominant. The male reproductive organ is called antheridium and produces flagellated sperms (antherozoids). The sperm fertilizes the egg (oosphere), which is produced by the archegonium that is the female reproductive organ. After fertilization, the zygote develops in the archegonium to produce sporophyte, which grows out from the gametophyte. The sporophyte produces haploid spores, which will eventually give rise to mature gametophytes. Pteridophytes Pteridophytes are the only non-flowering seedless plants possessing true stems with simple vascular tissues, and also true roots and leaves. This enables pteridophytes to achieve larger sizes than the bryophytes. In the tropics, ferns may grow up to 18 m (60 ft). A major difference between pteridophytes and bryophytes is that the diploid sporophyte generation is dominant in pteridophytes. The sporophyte generation of pteridophytes produce spores by meiosis. The pteridophytes may EXPERIMENT 8: PLANT DIVERSITY - BRYOPHYTES AND PTERIDOPHYTES
SB015 Lab Manual Updated: 26/04/2022 43 be homosporous which produce only one type of spore or heterosporous which produces two types of spores. The gametophyte generation of pteridophytes retains two traits that are reminiscent of the bryophyte. Firstly, the small gametophytes lack conducting vessels. Secondly, as in bryophytes, the flagellated sperms require water medium to reach the egg cell, so pteridophytes still depend on the presence of water for sexual reproduction. Experiment 8.1: Bryophytes Apparatus: Compound light microscope Materials: Prepared slides Marchantia sp. - capsule l.s. Marchantia sp. - female gametophyte (archegonium) l.s. Marchantia sp. - male gametophyte (antheridium) l.s. Polytrichum sp. - capsule l.s. *l.s. – longitudinal section Procedures and Observation: 1. Examine the prepared slide which shows the longitudinal sections of Marchantia sp. capsule. Draw and label the seta, foot, sporangium, spores and calyptra for capsule. 2. Examine the prepared slides which show the longitudinal sections of Marchantia sp. antheridium. Draw and label the antheridium and antheridiophore. 3. Examine the prepared slides which show the longitudinal sections of Marchantia sp. archegonium. Draw and label the egg cell, archegonium and archegoniophore. 4. Examine the prepared slides which show the longitudinal sections of Polytrichum sp. capsule. Draw and label the operculum, spore, and seta.
SB025 Lab Manual Updated: 15/09/2021 44 Figure 8.1 Capsule of Marchantia sp. (l.s.) (Adapted from: https://www.morton-pub.com/customize/images/immatureand-mature-sporophytes-callouts) Figure 8.2 Archegonia of Marchantia sp. (l.s.)
SB015 Lab Manual Updated: 26/04/2022 45 (Adapted from http://www.bio.miami.edu/dana/dox/altgen.html) Figure 8.3 Archegonia of Marchantia sp. (l.s.) 400x (Adapted from majorsbiology202) Figure 8.4 Antheridia of Marchantia sp. (l.s.) (Adapted from www.vcbio.science.ru.nl)
SB025 Lab Manual Updated: 15/09/2021 46 Figure 8.5 Antheridia of Marchantia sp. (l.s.) (Adapted from www.vcbio.science) Figure 8.6 Capsule of Polytrichum sp. (l.s.) (Adapted from www.k-state.edu) Questions: Bryophytes 1. State the unique characteristics of bryophytes. 2. How is the transport of substances carried out in bryophytes tissue? How is this feature related to the general size of these plants? 3. What is the process involved in spore formation of bryophytes? 4. Explain the adaptations of bryophytes to the terrestrial environment.
SB015 Lab Manual Updated: 26/04/2022 47 Experiment 8.2 Pteridophytes Apparatus: Compound light microscope Dissecting microscope Magnifying glass Razor blade Tiles Materials: Fresh specimens: Selaginella sp. (Division Lycopodiophyta) Dryopteris sp. (Division Pteridophyta) Prepared slides: Lycopodium sp. – strobilus l.s. Selaginella sp. – strobilus l.s. Procedures and Observation: 1. Examine the fresh specimens of Selaginella sp. to observe the dichotomous branching, types and arrangement of sporophyll and strobilus. 2. Examine the fresh specimens of Dryopteris sp. then draw and label the rhizome, rhizoid, rachis, frond, pinna and sorus. 3. Examine the prepared slides showing longitudinal sections of the strobilus of Lycopodium sp. and Selaginella sp.. Then draw and label sporophyll, sporangium and spore (homosporous or heterosporous).
SB025 Lab Manual Updated: 15/09/2021 48 Figure 8.7 Selaginella sp. Figure 8.8 Selaginella sp. Adapted from https://images.app.goo.gl/q3LCtEuBpNiDd4Tp8
SB015 Lab Manual Updated: 26/04/2022 49 Figure 8.9 Dryopteris sp. Figure 8.10 Strobilus of Lycopodium sp. (l.s.) (Adapted from www.stolaf.edu)
SB025 Lab Manual Updated: 15/09/2021 50 Figure 8.11 Strobilus of Selaginella sp. (l.s.) (Adapted from www.sfsu.edu) Questions: Pteridophytes 1. State the unique characteristics of pteridophytes. 2. Differentiate the spores of Lycopodium sp. and Selaginella sp. 3. Division Pteridophyta is considered to be more advanced than Division Lycopodiophyta. Explain the advanced characteristic of Division Pteridophyta.
SB015 Lab Manual Updated: 26/04/2022 51 EXPERIMENT 9: BIOCATALYSIS Objectives: At the end of this lesson, students should be able to: i. extract catalase from liver tissue; ii. observe the qualitative activity of catalase; iii. measure the quantitative activity of catalase; and iv. determine the factors affecting the catalase activity. Introduction: Enzymes are biological catalysts, normally proteins, synthesized by living organisms. Enzymes speed up reactions by lowering the activation energy. Enzymes are normally very specific. An enzyme catalyses a single reaction that involves one or two specific molecules called substrates. Each enzyme has evolved to function optimally at a particular pH, temperature and salt concentration. Some require the presence of other molecules called coenzymes, derived from water-soluble vitamins, for its function. The rate of reaction also depends on the amount of enzymes present. In this experiment, the enzyme to be extracted and tested is catalase, which present in almost all cells especially in the liver and red blood cells. The substrate for this enzyme is hydrogen peroxide (H2O2). The accumulation of hydrogen peroxide in the body is toxic. Catalase renders the hydrogen peroxide harmless by breaking it down to water and oxygen. catalase 2H2O2 2H2O + O2 The chemical properties of catalase resembles most those of the enzymes. (Note: The success of this experiment depends on the amount of catalase present in the prepared extract. The results of the catalase reaction can be observed clearly if the amount of enzyme in the extract is large. Use a boiling tube to avoid spillage during the reaction).
SB025 Lab Manual Updated: 15/09/2021 52 Apparatus: Beakers (250 ml & 1000 ml) Blade Boiling tube Boiling tube rack Dropper Electronic balance Filter funnel Glass rod Labelling stickers Measuring cylinder (10 ml) Mortar and pestle Muslin cloth Retort stand Rubber stopper Syringe (1 ml) Thermometer Tile Tissue paper Waterbath Materials: Distilled water Fresh liver of a cow/chicken 3% H2O2 solution 1 M H2SO4 Ice cubes 1% KMnO4 solution Phosphate buffer solutions (pH 5, pH 7, pH 9 and pH 11) Safety and laboratory Precaution 1. Ensure all Apparatus: are clean before using it, in order to obtain accurate results. 2. Measure precisely the volume of the solutions used. 3. Handle heated Apparatus: with care.
SB015 Lab Manual Updated: 26/04/2022 53 Experiment 9.1: Estimation of catalase activity Procedures and Observation: 9.1.1 Preparation of catalase extract 1. Cut 10-15 g of fresh liver tissue into small pieces and macerate the tissue in a mortar and pestle. 2. Gradually add 20 ml of water. 3. Filter the mixture into a beaker using the muslin cloth. 4. The filtrate will be the enzyme stock solution to be used in the experiment. 9.1.2 Qualitative test for catalase activity 1. Label two boiling tube as A and B 2. Pour 1 ml H2O2 solution into a boiling tube A. (Caution: H2O2 is a toxic substance). 3. Pour 1 ml enzyme stock solution in boiling tube B. 4. Pour the stock solution in boiling tube B into the boiling tube A. Label the boiling tube as C (enzyme-H2O2 mixture). 5. Record your observation in boiling tube C. Use this boiling tube C for the following experiment. 9.1.3 Estimation of catalase activity 1. Pour 1 ml H2SO4 into boiling tube D. 2. Measure 1 ml of enzyme-H2O2 mixture from boiling tube C and transfer into boiling tube D. Shake well the boiling tube. (Note: In the acidic medium, catalase will be denatured. The remaining H2O2 can be measured using KMnO4 solution. KMnO4 solution reacts with H2O2 in acidic medium.) 5H2O2 + 2KMnO4 + 4H2SO4 2KHSO4 + 2MnSO4 + 8H2O + 5O2 3. Fill the syringe with KMnO4. Then add drops of KMnO4 into test tube D until the red colour remains unchanged for 10 seconds.
SB025 Lab Manual Updated: 15/09/2021 54 4. Determine the amount(ml) of KMnO4 used. (Note: The higher amount of KMnO4 used indicates that more H2O2 is present in the boiling tube D. It means that the H2O2 is not fully broken down by catalase to water and oxygen molecules). 5. To measure catalase activity, we use this formula: Catalase activity = 1 Amount KMnO4 used (ml) Experiment 9.2: Factors affecting the activity of catalase Procedures and Observation: 9.2.1 Temperature 1. Put the following boiling tubes in a beaker containing chilled water (20°C). a) boiling tube A containing 2 ml of enzyme stock solution b) boiling tube B containing 3 ml of H2O2 c) empty boiling tube labelled C 2. Prepare 1 ml H2SO4 in boiling tube 1. 3. Using a thermometer, check the temperature of H2O2 in boiling tube B drops to 20°C, then pour the chilled H2O2 into boiling tube C. 4. Pour the cooled enzyme stock from boiling tube A into boiling tube C. (Note: Make sure that the reaction takes place in the iced-chilled beaker.) Record the time for 4 minutes. 5. After 4 minutes, measure 1 ml of enzyme-H2O2 solution from boiling tube C and transfer it into boiling tube 1. Plug boiling tube 1 with rubber stopper. Shake well and estimate the activity of catalase as conducted in Experiment 9.1.3. 6. Repeat the steps 1 to 5. (Set at different temperatures: 30oC, 40oC and 50oC). Use different boiling tubes. 7. Record the values obtained in Table 9.1 and plot the graph of the approximate catalase activity against temperature.
SB015 Lab Manual Updated: 26/04/2022 55 Table 9.1 The effects of temperature on catalase activity Temperature 20oC 30oC 40oC 50oC Amount (ml) of KMnO4 used Approximate catalase activity (1/amount of KMnO4 used) 9.2.2 pH 1. Label four boiling tubes as 2, 3, 4 and 5. Pour 1 ml of H2SO4 into each boiling tube. 2. Label four new boiling tubes as D, E, F and G. Pour 1 ml of H2O2 into each boiling tube. 3. Measure 2 ml phosphate buffer solution with pH 5, pH 7, pH 9 and pH 11 and add into boiling tube D, E, F and G respectively. Shake them well. (Caution: Always rinse the measuring cylinder before measuring different buffer solution to avoid inaccuracy) 4. Measure 1 ml enzyme stock solution and pour into boiling tube D. Record the time for 4 minutes. 5. After 4 minutes, measure 1 ml of enzyme-H2O2 mixture from boiling tube D and transfer into boiling tube 2. Shake well and estimate the activity of catalase as conducted in Experiment 9.1.3. 6. Repeat the above steps for boiling tubes E, F and G using boiling tubes 3, 4 and 5 respectively. 7. Record the values obtained in Table 9.2 and plot the graph of the approximate catalase activity against pH.
SB025 Lab Manual Updated: 15/09/2021 56 Table 9.2 The effects of pH on catalase activity pH 5 7 9 11 Amount (ml) of KMnO4 used Approximate catalase activity (1/amount of KMnO4 used) Questions: 1. What is the role of H2SO4 in the reaction? 2. Explain the effects of the following factors on the enzymatic reaction: a. temperature b. pH 3. What is the gas that will be released, when hydrogen peroxide is broken down catalyze by catalase? How can we test the gas it releases?
SB015 Lab Manual Updated: 26/04/2022 57 EXPERIMENT 10: CELLULAR RESPIRATION Objectives: At the end of this lesson, students should be able to: i. organize the experiment setting for redox reaction procedures; ii. conduct an experiment on redox reaction in cellular respiration; and iii. explain the biochemical processes in yeast suspension. Introduction: Aerobic cellular respiration produces ATP from glucose. As a cell breaks down the glucose, most of the energy comes as the hydrogens of glucose are released by enzymes in glycolysis and the Krebs cycle. The high energy electrons of the hydrogen are carried to the electron transport chain (ETC) in the forms of NADH and FADH2. We can demonstrate these redox reactions by substituting NAD+ with methylene blue (dye). In the oxidized state, this dye has a blue colour. When it is reduced, it becomes white or light blue as indicated below, hence the reduction has taken place. Methylene blue (blue/greenish blue) reduction ⇌ oxidation decolourised methylene (white/light blue) Apparatus: Beaker (250 ml) Boiling tubes Bunsen burner Cork or rubber stopper Labelling paper Measuring cylinder (10 ml) Pasteur pipette/ Glass dropper Stopwatch Thermometer Tripod stand Waterbath (40oC)
SB025 Lab Manual Updated: 15/09/2021 58 Materials: Methylene blue 0.1% Yeast suspension (5%) added to 1% glucose (freshly prepared) Procedures and Observation: 1. Label 3 boiling tubes as A, B and C. 2. Fill in tube with 10 ml of yeast suspension. 3. Heat tube C in boiling water for 5 minutes. 4. Add 5 drops of methylene blue into each of the boiling tubes using Pasteur pipette. Shake gently to ensure the colour is evenly distributed. 5. Observe the colour of the yeast suspension in all boiling tubes. Record your observation in Table 10.1. 6. Incubate all boiling tubes in the water bath at 40oC for 15 minutes. 7. Observe the colour changes of the yeast suspension in all boiling tubes. Record your observation in Table 10.1. 8. Place tube B in boiling water for 5 minutes. (Note: Make sure the water boils first before placing tube) 9. Plug boiling tube A, B and C with cork or rubber stopper. Press it with your thumb and shake the tube vigorously for 30 seconds. Observe the colour changes of the yeast suspension. Record your observation in Table 10.1. 10. Remove the stopper and incubate all boiling tubes in water bath at 40oC for 15 minutes. 11. Observe the colour of the yeast suspension in each boiling tube. Record your observations in Table 10.1. (Note: Observations are based on the colour changes.)
SB015 Lab Manual Updated: 26/04/2022 59 Table 10.1 Colour changes observed for demonstrating redox reactions in yeast using methylene blue Treatments Colour Tube A Tube B Tube C Before incubation (40oC) After first incubation (40oC) After vigorous shaking After second incubation (40oC) Questions: 1. Explain the redox reaction. 2. What is the substance in a living cell that has the same function as methylene blue? 3. Name the important process which involves substances in question no. 2 above. 4. Are enzymes responsible for the colour changes? State your reason.
SB025 Lab Manual Updated: 15/09/2021 60 EXPERIMENT 11: CHROMATOGRAPHY Objectives: At the end of this lesson, students should be able to: i. demonstrate chromatography technique to separate the photosynthetic pigments; and ii. calculate Rf value. Introduction: The chloroplasts in green plants contain many pigments such as chlorophyll a, chlorophyll b, carotene, phaeophytin and xanthophylls. These pigments have different solubility in certain solvent and they can be separated by chromatography. Paper chromatography is a useful technique for separating and identifying pigments and other molecules from cell extracts that contain a complex mixture of molecules. Typically, a drop of the sample is applied as a spot to a sheet of chromatography paper. The solvent moves up the paper by capillary action, which occurs as a result of the attraction of solvent molecules to the paper and the attraction of solvent molecules to one another. As the solvent moves up the paper, it carries along any substances dissolved in it. The pigments are carried along at different rates because they are attracted to different degrees, to the fibres in the paper through the formation of intermolecular bonds, such as hydrogen bonds. Another factor that is taken into account is molecular weight of the pigment. Apparatus: Beaker (100 ml) Blade Boiling tube rack Boiling tube with cork stopper Chromatography paper strip (Whatman No. 3) with pointed end Dissecting pin/ Capillary tube Filter funnel Forceps Hair dryer Labelling paper Measuring cylinder Mortar and pestle Muslin cloth Spatula
SB015 Lab Manual Updated: 26/04/2022 61 Materials: Fresh leaves: i. Sauropus sp. (Cekur manis) ii. Pandanus sp. (Pandan) iii. Erythrina sp. (Dedap) iv. Coleus sp. (Ati-ati) Solvent (mixture of ether petroleum-acetone at 9:1, freshly prepared) Acetone 80% (Precaution: Should be handled in fume cupboard, do not inhale the fume) Experiment 11.1: Leaf extract preparation Procedures and Observation: 1. Cut approximately 20g of fresh leaves using a blade. 2. Grind the leaves and add 5 ml acetone gradually. 3. Leave them for 10 minutes. 4. Grind again and add another 5 ml acetone. 5. Filter the extraction using muslin cloth. Remarks: Extraction of the pigments also can be done by carefully pressing and moving a coin back and forth more than 10 times on top of the leaf onto the chromatography paper until enough pigments are placed on the chromatography paper.
SB025 Lab Manual Updated: 15/09/2021 62 Experiment 11.2: Paper Chromatography 1. Prepare 3-5ml of solvent in the boiling tube. (Note: Close the boiling tube with cork before use to prevent the solvent evaporate.) (Precaution: Make sure the solvent stock is always close to prevent evaporation) 2. By using a pencil, draw a horizontal line 1 cm from the end of the chromatography paper. Then mark 2cm from the pointed end of the chromatography paper. Refer to Figure 11.1. Figure 11.1: Suggestion measurement for chromatography paper 3. Using the blunt end/ head of dissecting pin or a capillary tube, place a drop of the leaf extract on the chromatography strip at the origin. Let the drop dry completely. Repeat the process more than 15 times to build up a small area of concentrated pigment. (Precaution: Use forceps to handle the chromatography strip throughout the experiment). 4. Attach the blunt end of the chromatography strip on a cork with a dissecting pin. 5. Suspend the strip straight into the boiling tube containing solvent. (Note: The pointed end of the chromatography strip should be dipped into the solvent, but make sure that the pigment spot (pigment origin) is not immersed in the solvent.) 6. Place boiling tube vertically in the boiling tube rack. 7. Let the solvent rise until it reaches the solvent front. 1 cm Solvent front Pigment origin
SB015 Lab Manual Updated: 26/04/2022 63 8. Remove the chromatography paper. Mark the pigmented area. 9. Dispose the solvent by diluting it with running tap water and then pour the solvent into the sink. 10. Identify the pigment colour and measure the distance for each pigment. 11. Calculate the Rf value for each pigment using the following formula: Rf = Distance moved by the pigment from the origin Distance moved by the solvent from the origin Figure 11.1 Paper chromatography set up using a boiling tube 12. Record your results in the Table 11.1.
SB025 Lab Manual Updated: 15/09/2021 64 Table 11.1 Photosynthetic pigments and the observed Rf values Pigment Colour Observed Rf value Chlorophyll b Chlorophyll a Xanthophyll Phaeophytin Carotene (Remarks – it is recommended that different groups of plant be used) Figure 11.2 Chromatography paper shows the distance for each pigment from origin
SB015 Lab Manual Updated: 26/04/2022 65 Questions: 1. Do the leaf extracts from different plants contain the same pigments? Explain why. 2. Name the most common pigment which is usually found in many plants. Explain your answer. 3. Name the five common photosynthetic pigments in plant and state its role. Extension Try this experiment using C4 or CAM leave. Observe the difference between the pigment intensity for C3, C4 and CAM plant.
SB025 Lab Manual Updated: 15/09/2021 66 EXPERIMENT 12: MAMMAL ORGAN SYSTEM Objectives: At the end of this lesson, students should be able to: i. demonstrate dissecting skill; and ii. examine the organ systems in mammal: Digestive, Circulatory, Respiratory, Urogenital and Nervous System. Introduction: An organ system is a group of anatomical structures that work together to perform a specific function or task. Although we learn about each organ system as a distinct entity, the functions of the body's organ systems overlap considerably, and your body could not function without the cooperation of all its organ systems. In fact, the failure of even one organ system could lead to severe disability or even death. A mammalian body is composed of different organ systems which include the following: ● Integumentary ● Muscular ● Skeletal ● Nervous ● Circulatory ● Lymphatic ● Respiratory ● Endocrine ● Urinary/excretory ● Reproductive ● Digestive Apparatus: Dissecting set Dissecting pins Dissecting tray Petri dish (Note: Demonstration by the lecturer on how to use the dissecting set.) Material: Chloroform Cotton wool
SB015 Lab Manual Updated: 26/04/2022 67 Disposable gloves Freshly killed mice (with chloroform) Surgical mask Safety Precaution: 1. Wear mask and glove during the experiment 2. Be careful while handling sharp Apparatus: Procedures and Observation: 12.1 Digestive, Circulatory, Respiratory, Urogenital 1. Lay down the mice on a dissecting tray, with its ventral surface facing upward. Spread the legs and pin at 45° angle as shown in Figure 12.1. Figure 12.1 Pin the legs of the mice at 45° angle 2. Use forceps to lift the skin on the mid-ventral line (Figure 12.2). Figure 12.2 Lifting the skin on the mid ventral line
SB025 Lab Manual Updated: 15/09/2021 68 3. Slit the skin along the mid-ventral line with scissor. Figure 12.3(a) Male mice (Note: Keep the scissors as low as possible to avoid from cutting the body wall underneath the skin.) Male: Cut straight up until you reach the lower jaw. Cut straight down, until around the penis and end at the scrotal sacs (Figure 12.3a). Female: Cut the skin as described for the male, but continue to cut straight down, passing on either side of the urinary and genital apertures to the anus (Figure 12.3b). Figure 12.3(b) Female mice
SB015 Lab Manual Updated: 26/04/2022 69 4. Cut through the skin towards the end of each limb. With a scalpel, separate and pull the skin aside to expose the abdominal wall (Figure 12.4). (Note: Be careful not to tear off the nerves and muscles at the axillary region.) Figure 12.4 Exposing the abdominal wall 5. Stretch the skin and pin it back as shown in Figure 12.5. Lift the abdominal wall with forceps and make an incision as shown. Using a blunt end scissors, cut through the body wall to expose the components of the abdomen.
SB025 Lab Manual Updated: 15/09/2021 70 Figure 12.5 Making an incision on the abdominal wall
SB015 Lab Manual Updated: 26/04/2022 71 Figure 12.6 Exposing the internal anatomy of the abdomen 6. Pin aside the abdominal wall (Figure 12.6). 7. Observe the digestive and reproductive systems of the mice. 8. Remove the fat bodies as shown in Figure 12.7 when necessary.
SB025 Lab Manual Updated: 15/09/2021 72 Figure 12.7 Exposing the lower abdominal region (Note: Do not use sharp instruments while observing internal organs.) Male: i. Cut the ureters. Pin the bladder, seminal vesicle and rectum. ii. Remove the fat body on the right of the mice. iii. The blood vessels can be traced through the right groin by easing away the muscle and connective tissue with forceps. Trim with a pair of scissors if necessary. iv. Remove the remains of the mesentery and fat to display the aorta and posterior vena cava.
SB015 Lab Manual Updated: 26/04/2022 73 Female: i. Cut the ureters. ii. Pin the rectum. iii. Lay aside the vagina and bladder as shown and pin it if necessary. iv. The blood vessels can be traced through the right groin by easing away the muscle and connective tissue with forceps. Trim with a pair of scissors if necessary. v. Remove the remains of the mesentery and fat to display the aorta and posterior vena cava. 9. Cut with blunt end scissor through the side wall of the thorax along the line indicated as shown in Figure 12.8. Figure: 12.8 Exposing the thoracic cavity 10. Continue the cut to the apex by turning the ventral part of the thoracic wall aside and pull it slightly to avoid cutting the heart. Repeat on the other side to remove the ventral part of the thoracic wall entirely. Remove the loose parts of the pleura (refer to Figure 12.8).
SB025 Lab Manual Updated: 15/09/2021 74 11. Observe the components of the thorax as they appear at this stage. Refer to Figure 12.9. Figure 12.9 Components of the thorax 12. Remove the thymus gland as shown in Figure 12.10. Clear away the fat tissues around the great vessels.
SB015 Lab Manual Updated: 26/04/2022 75 Figure 12.10 Removing the thymus gland 13. Pin the heart to the right of the mice. Observe the structures in Figure 12.11. Figure 12.11 Circulatory system of the mice