BIOLOGY LABORATORY MANUAL

SB015 Lab Manual

Hairline shape

Widow’s peak hairline Straight hairline
https://www.healthline.com/health/wid https://www.ishrs-htforum.org

ows-peak

Chin shape

Cleft chin Smooth chin
https://www.healthline.com/health/cleft https://www.mentalfloss.com/

-chin

Thumb bending

Normal thumb bending Hitchhiker’s thumb
https://askabiologist.asu.edu https://askabiologist.asu.edu

Figure 5.1: The six inherited characteristics in human

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Results

Table 5.1 Observed and expected frequencies of each genotype for six
characteristics in the class

Characteristic Phenotype Genotype Tick (√) Observed
your own frequency of
genotype each phenotype
in the class

Earlobe Free earlobe P_
Tongue Rolling Attached pp
Dimple earlobe C_
Hairline shape Ability of
tongue rolling cc
into “U” shape D_
Inability of dd
tongue rolling H_
into “U” shape hh

Have dimple

Without
dimple
Widow’s peak
hairline

Straight
hairline

Chin shape Cleft chin S_
Thumb ss
Smooth chin T_
tt
Normal
thumb-
bending
Ability to bend
thumb at 60°
angle or more

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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.

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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

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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.

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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.

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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)

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Materials:

Ice-chilled 95% alcohol salt-
Ice cubes detergent
Kiwi/ banana/onion/tomato/watermelon solution
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).

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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?

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BIOLOGY 2
SB025

SB025 Lab Manual

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)

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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)

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Figure 7.3 Different shapes of bacteria
(Adapted from

commons.wikimedia.org/wiki/File:OSC_Microbio_03_03_ProkTable.jpg)

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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.

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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 Gram-
negative, 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.

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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 Gram-
negative bacteria?

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SB025 Lab Manual

EXPERIMENT 8: PLANT DIVERSITY - BRYOPHYTES AND
PTERIDOPHYTES

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 non-
vascular 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

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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.

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Figure 8.1 Capsule of Marchantia sp. (l.s.)
(Adapted from: https://www.morton-pub.com/customize/images/immature-

and-mature-sporophytes-callouts)

Figure 8.2 Archegonia of Marchantia sp. (l.s.)

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(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)

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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.

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SB015 Lab Manual

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).

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Figure 8.7 Selaginella sp.

Figure 8.8 Selaginella sp.
Adapted from https://images.app.goo.gl/q3LCtEuBpNiDd4Tp8

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Figure 8.9 Dryopteris sp.

Figure 8.10 Strobilus of Lycopodium sp. (l.s.)
(Adapted from www.stolaf.edu)

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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.

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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).

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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.

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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.

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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.

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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.

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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?

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SB015 Lab Manual

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 reduction decolourised methylene
(blue/greenish blue) ⇌ (white/light blue)

oxidation

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)

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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.)

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Table 10.1 Colour changes observed for demonstrating redox
reactions in yeast using methylene blue

Treatments Tube A Colour Tube C
Tube B
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.

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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

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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.

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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.

1 cm

Solvent front

Pigment origin

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.

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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.

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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

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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.

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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 66
Cotton wool

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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

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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).

Updated: 15/09/2021 Figure 12.3(b)
Female mice

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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.

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Figure 12.5 Making an incision on the abdominal wall

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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.

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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.

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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).

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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.

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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

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