e-BOOK MANUAL AMALI BIOLOGY

SB015 Lab Manual

Questions

1. Individuals with certain heterozygous characteristics are usually
called a carrier. What does a carrier mean?

2. A student inherited left handedness from parents who are both
right handed. Explain the pattern of inheritance.

3. What is the expected frequency for a person having tongue
rolling ability and attached earlobe?

4. What is the expected frequency for a person to have all six
recessive characteristics?

Exercise 5.2: ABO blood group inheritance

ABO blood groups in human are examples of multiple alleles of a
single gene and also codominant alleles. Each individual inherited any
one of four blood types, i.e. A, B, AB or O. Type A groups are
determined by the presence of antigen A found on the surface of red
blood cells (erythrocytes), while the blood plasma contains B antibody
which agglutinates type B blood. Individuals with type B blood have
antigen B and antibody A which agglutinates type A blood. Individuals
with type AB blood have both antigen A and antigen B but without
antibodies A or B. Finally, individuals with type O blood have
antibody A and antibody B but without any antigen. Table 5.2 shows
individual characteristics for all ABO blood groups.

Table 5.2Individual characteristics for all ABO blood groups

Blood group Antigen Antibodies present Agglutinated
(phenotypes) present in blood plasma blood group
(serum)
A on
B erythrocytes Anti-B B
AB A
A Anti-A none
O A and B
B none
Anti-A and
A and B
Anti-B
none

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Apparatus

Depression slide/Pallet
Lancing device

Materials

Anti-A and Anti-B serum/ Blood test kit
Alcohol swab
Sterilized lancet
Toothpicks

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 for 10 – 15 seconds.

(Caution: Do not use the same lancet twice or exposed lancet)
4. Apply alcohol to your middle finger. Prick the tip of the middle

finger using sterilized lancet.
5. Wipe off the first blood drop.
6. Place the next drop at the center of slide 1 and 2.
7. Drop an Anti-A serum near the blood on slide 1 and Anti-B

serum on slide 2.
8. Mix the blood and serum on slide 1 with a toothpick. Use

another toothpick for slide 2.
9. 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.
10. Calculate the frequency of each blood group in the class. Record
your observation in Table 5.3.

Your blood group:

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Table 5.3Frequency of blood group in the class

Blood group Possible genotypes Frequency of each
blood group
A
B
AB
O

Questions

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.
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 babies in the nursery.
Explain.

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SB015 Lab Manual

EXPERIMENT 6: BASIC TECHNIQUES IN ISOLATING DNA

Course Learning Objective: Conduct biology laboratory work on
microscopy, biological molecules, histology and genetics information by
applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To isolate DNA from plant tissue.

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

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.

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SB015 Lab Manual

Apparatus

Mortar and pestle
500 ml beaker
Muslin cloth
Boiling tube
Boiling tube rack
Water bath (60 °C)

Materials

Kiwi /banana/onion/tomato/watermelon salt-
Ice-chilled 95% alcohol detergent
Ice cubes solution
50.000g sodium dodecyl sulfate or sodium lauryl sulfate (SDS or SLS)
8.770g sodium chloride
4.410g sodium citrate
0.292g ethylenediaminetetraacetic acid (EDTA)
1 liter water

Procedures and Observation

Exercise: Isolation of crude DNA.

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 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.

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7. Very carefully pour 10ml of ice-chilled alcohol into the side of
the boiling tube (at flat angle). (Remark: make sure both liquid
do not mix and alcohol form a separate layer on top of the sieved
liquid)

8. Put the boiling tube into a rack and observe it. Observe 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

2. Why do we need to mash the fruits?

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BIOLOGY 2
SB025

SB025 Lab Manual

EXPERIMENT 7: DIVERSITY OF BACTERIA

Course Learning Objective: Conduct biology laboratory work on
diversity of bacteria and plant, biocatalysis, cellular respiration,
chromatography and dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To demonstrate Gram staining technique in classifying bacteria
ii. To identify Gram-positive and Gram-negative bacteria
iii. To identify different shapes of bacteria

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

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.

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Apparatus

Compound microscopes
Slides
Wash bottle
Bunsen burner
Bacterial loops
Petri dish
Forceps
Staining racks

Materials

Prepared slides of different types of bacteria
Cultures of Escherichia coli
Cultures of Staphylococcus aureus
Yoghurt (diluted in water 1:10)
Immersion oil
Safranin
Crystal violet
95% ethanol
Iodine
Filter paper
Labelling stickers

Figure 7.1 Comparative staining and cell wall structures of Gram-positive
and Gram-negative bacteria. (Adapted from www.quia.com)

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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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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. Place a loop of sterile distilled water on the slide and put a little
bit of bacterial colony.

3. Gently heat the slide to fix the bacteria onto the slide.
4. Place the slide on the staining rack. Cover the smear with single

drop of crystal violet and wait for 30 seconds to one minute.
5. Gently, rinse the slide with slow running water.
6. 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 one 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.
7. Rinse the slide with water as in step 6.
8. 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.
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. 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.
9. Add few drops of safranin on the bacterial smear and leave it for
approximately 30-45 seconds.
Colourless Gram-negative cell will readily accept the light
red safranin stain, while the already dark coloured Gram-
positive cell will undergo no change at all.
10. Rinse off with water and blot dry with filter paper.
11. Observe the slide under oil immersion magnification and
describe your observation in terms of types of bacteria, shape,
colour and determine whether it is Gram-positive or Gram-
negative.
12. Repeat steps 2-11 for microorganisms 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 +ve/-ve
E. coli

S. aureus
Bacteria from

yoghurt

Questions

1. Why Gram-positive bacteria purple in colour while Gram-negative
are red?

2. List some examples 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

Course Learning Objective: Conduct biology laboratory work on
diversity of bacteria and plant, biocatalysis, cellular respiration,
chromatography and dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To observe the diversity of species in bryophytes and

pteridophytes.
ii. To construct scientific drawing of bryophytes and

pteridophytes.

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

Introduction

Bryophytes and pteridophytes are two large groups of spore 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.

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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
vascular tissues – xylem and phloem. 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 gametophyte generation retains two traits that are
reminiscent of the bryophyte. Firstly, the small gametophytes lack
conducting vessels. Secondly, as in bryophytes, the flagellated sperms
(antherozoids) require water medium to reach the egg (oosphere), so
pteridophytes still depend on the presence of water for sexual
reproduction. Pteridophytes have true stems with vascular tissues, and
also true roots and leaves.

Exercise 8.1 Bryophytes

Apparatus

Compound microscope

Materials

Prepared slides
Marchantia sp. - capsule l.s
Marchantia sp. - male gametophyte (antheridium) l.s
Marchantia sp. - female gametophyte (archegonium) l.s
Polytrichum sp. - capsule l.s

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SB025 Lab Manual

Procedures and Observation

1. Examine the prepared slides which show the longitudinal
sections of Marchantia sp. capsule, antheridium and
archegonium. Draw and label the seta, foot, sporangium, spores
and calyptra.

2. Examine the prepared slides which show the longitudinal
sections of Polytrichum sp. capsule. Draw and label the
operculum, spore, peristome, annulus, calyptra, seta and capsule.

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

mature-sporophytes-callouts

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Figure 8.2 Archegonia of Marchantia sp. (l.s)
(Adapted from http://www.bio.miami.edu/dana/dox/altgen.html)

Figure 8.3 Archegonia of Marchantia sp. (l.s) 400x 41
(Adapted from majorsbiology202)

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SB025 Lab Manual

Figure 8.4 Antheridia of Marchantia sp. (l.s)
(Adapted from www.vcbio.science.ru.nl)

Figure 8.5 Antheridia of Marchantia sp. (l.s) 42
(Adapted from www.vcbio.science.

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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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SB025 Lab Manual

Exercise 8.2 Pteridophytes

Apparatus

Compound 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 specimens of Selaginella sp. Observe the
dichotomous branching, types and arrangement of sporophyll and
strobilus.

2. Examine the specimens of Dryopteris sp. 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. Draw and label
sporophyll, sporangium and spore (homosporous or
heterosporous).

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Figure 8.7 Selaginella sp.

Figure 8.8 Dryopteris sp.

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Figure 8.9 Strobilus of Lycopodium sp. (l.s)
(Adapted from www.stolaf.edu)

Figure 8.10 Strobilus of Selaginella sp. (l.s) 46
(Adapted from www.sfsu.edu)

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Questions

Pteridophytes

1. State the unique characteristics of pteridophytes.
2. Fern sporophytes have an underground stem called rhizomes.

How do you distinguish that rhizomes are stems and not roots?
3. Compare the spores of Lycopodium sp. and Selaginella sp.
4. Division Pteridophyta is considered to be more advanced than

Division Lycopodiophyta. Explain the advanced characteristic of
Division Pteridophyta.

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SB025 Lab Manual

EXPERIMENT 9: BIOCATALYSIS

Course Learning Objective: Conduct biology laboratory work on
diversity of bacteria and plant, biocatalysis, cellular respiration,
chromatography and dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To extract catalase from liver tissue
ii. To observe the qualitative activity of catalase.
iii. To measure the quantitative activity of catalase.
iv. To determine the factors affecting the catalase activity.

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

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 2H2O + O2
2H2O2

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
test tube to avoid spillage during the reaction).

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SB025 Lab Manual

Apparatus

Beakers (250 ml & 1000 ml)
Glass rod
Boiling tube
Boiling tube rack
Blade
Dropper
Filter funnel
Measuring cylinder (10 ml)
Mortar and pestle
Muslin cloth
Labelling stickers
Retort stand
Rubber stopper
Syringe (1 ml)
Thermometer
Tile
Tissue paper
Waterbath

Materials

Fresh liver of a cow/chicken
Distilled water
3 % H2O2 solution
1 M H2SO4
1 % KMnO4 solution
Ice cubes
Phosphate buffer solutions (pH 5, pH 7, pH 9 and pH 11)

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Exercise 9.1: Estimation of catalase activity

Procedures and Observation

A) 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.

B) 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. Using a dropper, add the stock solution in boiling tube B into the

boiling tube A. Label the tube as boiling tube C.
5. Observe and explain the activity of the enzyme. Use this boiling tube C

for the following.

C) Estimation of catalase activity

1. Pour 1 ml H2SO4 into boiling tube D.
2. Transfer 1 ml of enzyme-H2O2 mixture from C into D. Shake well the

boiling tube.

3. The left over H2O2 solution that does not react can be measured using
KMnO4 solution.

4. KMnO4 solution reacts with H2O2 in acid medium.

5H2O2 + 2KMnO4 + 4H2SO4 2KHSO4 + 2MnSO4 +
8H2O + 5O2

5. Fill the syringe with KMnO4.
6. Using the syringe, add drops of KMnO4 into test tube D until the red

colour remains unchanged for 10 seconds.
7. Determine the amount of KMnO4 used. The value shows the activity of

catalase.
8. The more KMnO4 is used indicates that more H2O2 is present in the

mixture. It means that the H2O2 is not fully broken down by catalyst to
oxygen molecules.

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Exercise 9.2: Factors affecting the activity of catalase

Procedures and observation

A)Temperature

1. Put the following boiling tubes in a beaker containing chilled water
(20°C).
a) boiling tube A containing 2ml 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 5.
3. When the temperature in the boiling tube B containing H2O2 drop to

20°C, pour the chilled H2O2 into boiling tube C.
4. Pour the cooled enzyme stock from boiling tube A into boiling tube C.

Make sure that the reaction takes place in the iced-chilled beaker.
Record the time.
5. After 4 minutes, transfer 1 ml of solution from boiling tube C into
boiling tube 5 and then plug with rubber stopper. Shake well and
estimate the activity of catalase as conducted in Exercise 15.1C.
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.

Temperature 20oC 30oC 40oC 50oC

Amount (ml) of
KMnO4 used

Approximate
catalase activity

(1/amount of
KMnO4 used)

Table 9.1 The effects of temperature on catalase activity

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B) pH

1. Label four boiling tubes as 6, 7, 8 and 9.
2. Pour 1 ml of H2SO4 into each boiling tube.
3. Label four boiling tubes as F, G, H and I.
4. Pour 1 ml of H2O2 into each boiling tube.
1. Add 2 ml phosphate buffer solution with pH 5, pH7, pH 9 and pH 11

to boiling tube F, G, H and I, respectively. Shake them well.
2. Pour 1 ml enzyme stock solution into boiling tube F. Record the time

for 4 minutes.
3. After 4 minutes, transfer 1 ml of solution from boiling tube F into

boiling tube 6. Shake well and estimate the activity of catalase as
conducted in Exercise 15.1C.
4. Repeat the above steps for boiling tubes G, H and I using boiling tubes
7, 8 and 9 respectively.
5. Record the values obtained in Table 9.2 and plot the graph of the
approximate catalase activity against pH.

pH 5 7 9 11

Amount (ml) of
KMnO4 used

Approximate
catalase activity

(1/amount of
KMnO4 used)

Table 9.2 The effects of pH on catalase activity

Notes:
1. Ensure all apparatus are clean, in order to obtain accurate results.
2. Measure precisely the volume of the solutions 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

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SB025 Lab Manual

EXPERIMENT 10: CELLULAR RESPIRATION

Course Learning Objective: Conduct biology laboratory work on diversity of
bacteria and plant, biocatalysis, cellular respiration, chromatography and
dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:

i. To organize the experiment setting for redox reaction procedures
ii.To conduct an experiment on redox reaction in cellular respiration
iii.To explain the biochemical processes in yeast suspension

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

Introduction

Aerobic cellular respiration produces ATP from glucose. As an organism
breaks down the glucose, most of the energy comes as the hydrogens of
glucose are removed by enzymes in glycolysis and the citric acid cycle. The
electrons of the hydrogens 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. 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) oxidation (white/light blue)

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SB025 Lab Manual

Apparatus

Beaker (250 mL)
Boiling tubes
Bunsen burner
Cork or rubber stopper
Dropper
Labelling paper
Measuring cylinder (10 mL)
Pasteur pipette
Stopwatch
Thermometer
Tripod stand
Water bath (380C – 420C)

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 tubes using Pasteur

pipette. Shake gently to ensure the colour is evenly distributed.
5. Incubate all tubes in the water bath (40oC) for 15 minutes.
6. Observe the colour changes in all tubes.
7. Heat tube B in boiling water for 5 minutes.
8. Plug 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. Remove the stopper and incubate all tubes in water
bath (40oC) for 15 minutes.
9. Observe the colour of the yeast suspension precipitate in each tube.
Note: Observations are based on the colour changes.
10. Record your observations in Table 10.1.

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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 √
Boiling 5 minutes √ √ √
5 drops of √ √
methylene blue √
First incubation √ √
(40oC)
After first √
incubation

Boiling 5 minutes

Vigorous shaking

Second incubation
(40oC)
After second
incubation

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. Explain the biochemical processes based on the observations in boiling

tubes A, B and C.
5. Are enzymes responsible for the colour changes? State your reason.

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SB025 Lab Manual

EXPERIMENT 11: PHOTOSYNTHESIS

Course Learning Objective: Conduct biology laboratory work on
diversity of bacteria and plant, biocatalysis, cellular respiration,
chromatography and dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To demonstrate chromatography technique to separate the

photosynthetic pigments
ii. To calculate Rf value

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

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 size.

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Apparatus

Beaker (100 mL)
Blade
Boiling tube rack
Boiling tube with cork stopper
Chromatography paper strip (Whatman No. 3) with pointed end
Dissecting pin
Filter funnel
Forceps
Hair dryer
Labelling paper
Measuring cylinder
Mortar and pestle
Muslin cloth
Spatula

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% (Should be handled in fume cupboard, do not inhale the
fume)

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Procedures and Observation

Exercise 11.1 Chlorophyll extract preparation

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.

Exercise 11.2 Paper Chromatography

1. Using the tip of dissecting pin, place a drop of the chlorophyll
extract on the chromatography strip. Let the drop dry completely.
Repeat the process more than 15 times to build up a small area of
concentrated pigment.
(Caution: Use forceps to handle the chromatography strip
throughout the experiment).

2. Attach the paper strip to the stopper with a pin. Suspend the strip
straight into the boiling tube that contains 3-5 mL solvent. The
bottom of the paper should be dipped into the solvent, but make
sure that the pigment spot (point of origin) is not immersed in
the solvent. Place the chromatography paper strip vertically in
the tube rack.

3. Let the solvent rise until its front reaches 1cm from the top of the
strip.

4. Remove the chromatography paper strip and mark the solvent
front with pencil. Mark the pigmented area.

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5. 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

stop cork

pin

Whatman No. 3
filter paper

pigment extract
solvent

Figure 11.1 Paper chromatography set up using a boiling tube

6. Record your results in the Table 11.1.

Table 11.1 Photosynthetic pigments and the observed Rf values

Pigment Colour Standard Rf Observed Rf
Chlorophyll b Yellow-green value value

0.45

Chlorophyll a Blue-green 0.65

Xanthophyll Yellow 0.71

Phaeophytin Grey 0.83

Carotene Orange 0.95

(Remarks – it is recommended that different groups of plant be used)

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11.2 cm SB025 Lab Manual

6.4 cm Solvent front
5.2 cm 12 cm
4.1 cm

Pigment origin

Solvent origin

Figure 11.2 Paper chromatography shows the value for each pigment.

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. Why do plants have different types of pigment?

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EXPERIMENT 12: MAMMAL ORGAN SYSTEM

Course Learning Objective: Conduct biology laboratory work on
diversity of bacteria and plant, biocatalysis, cellular respiration,
chromatography and dissecting technics by applying manipulative skills.
(P3, CLO 2, PLO 2, MQF LOD 2)

Learning Outcomes:
At the end of this lesson, students should be able to:
i. To demonstrate dissecting skill.
ii. To examine the organ systems in mammal: Digestive,

Circulatory, Respiratory, Urogenital and Nervous System.

Student Learning Time (SLT):

Face-to-face Non face-to-face

2 hour 0

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 of its organ systems. In fact, the failure
of even one organ system could lead to severe disability or even death.

A mammallian body is composed of different organ systems which
include the following:

 Integumentary
 Muscular
 Skeletal
 Nervous
 Circulatory
 Lymphatic
 Respiratory
 Endocrine
 Urinary/excretory
 Reproductive
 Digestive

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Apparatus
Dissecting set
Dissecting pins
Dissecting tray
Petri dish
Note: Demonstration by the lecturer on how to use the dissecting kit.

Material
Chloroform
Cotton wool
Disposable gloves
Mice
Surgical mask

Procedures and Observation
1. Put the mice to sleep.
2. 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.

Dissecting tray
Figure 12.1 Pin the legs of the mice at 45° angle

3. Use forceps to lift the skin on the mid-ventral line (Figure 12.2).

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Figure 12.2 Lifting the skin on the mid ventral line
4. Slit the skin along the mid-ventral line.

Penis
Scrotal sac

Figure 12.3(a) Male mice

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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,
till 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

4. Cut through the skin towards the end of each limb. 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.

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Figure 12.4 Exposing the abdominal wall

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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 pair of scissors, cut through the body wall to

Figure 12.5 Making an incision on the abdominal wall
expose the components of the abdomen.

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6. Pin aside the abdominal wall (Figure 12.6).

Figure 12.6 Exposing the internal anatomy of the abdomen

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7. Observe the digestive and reproductive systems of the mice.
8. Remove the fat bodies as shown in Figure 12.7 when
necessary.

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.

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iii. The blood vessels can be traced through the right groin by
easing away the muscle and connective tissue with forceps.
iv. Trim with a pair of scissors if necessary.
Female: Remove the remains of the mesentery and fat to display the
aorta and posterior vena cava.

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.
vi. Cut through the side wall of the thorax along the line
indicated as shown in Figure 12.8.

9. 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

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wall entirely. Remove the loose parts of the pleura (refer to Figure
12.8).

Figure: 12.8 Exposing the thoracic cavity

10. Observe and draw the components of the thorax as they appear
at this stage. Refer to Figure 12.9.

FFigiugruer1e21.92.C9 oCmopmopnoennetns tosfotfhtehtehtohroarxax

10. Remove the thymus gland as shown in Figure 12.10. Clear away
the fat tissues around the great vessels.

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FFiigguurree1122.1.100 RReemmoovviinngg tthhe thymus gland
11. Pin the heart to the right of the mice. Observe the structures in

Figure 12.11 Circulatory system of the mice 71
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SB025 Lab Manual

Figure 12.11.

12. Based on your observations, draw a labelled diagram of the
organ systems in mammal; Digestive System, Circulatory
System, Respiratory System, Urogenital System and Nervous
System.

Figure 12.12 Digestive System

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Figure 12.13 Circulatory System.

Figure 12.14 Respiratory System 73
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Figure 12.15 Urogenital System (Male)

Figure 12.16 Urogenital System (Female) 74
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