IAS Biology
Unit 3
CORE PRACTICAL 1
Use a semi-quantitative method with Benedict’s reagent to estimate the concentrations of
reducing sugars and with iodine solution to estimate the concentrations of starch, using color
standards
Estimating the concertation of reducing sugar in an unknown solution:
Steps
1. Decide on a range of glucose concentrations before beginning to work.[e.g. 0.25%,
0.5%, 1.0%, 1.5%, 2.0%]
2. Label 6 test tubes and the 5 beakers with the concentrations you’ve decided on, and
one test tube will be labeled with your control experiment
3. Using a clean syringe, distilled water, and 0.25% glucose, fill the labeled beaker with
5 cm3 of glucose solution. Repeat this for the four other concentrations.
4. Using a clean syringe, fill the 6 labeled test tubes with 2 cm3 of Benedict’s reagent
5. Add 1 cm3 of each glucose solution to its corresponding labeled test tube, and add
1 cm3 of the control solution to the 6th test tube [e.g. fruit juice]
6. Give each test tube a gentle shake to ensure the contents are well-mixed, and place
the test tubes in a water bath of a known temperature, and start a timer [e.g. 5
minutes]
7. After the [5 minutes] take out the test tubes carefully, using tongs, as to not injure
yourself.
8. Observe the test tubes and record the color change for each concentration
Section 1 – Independent and dependent variables
Dependent variable – color of Benedict’s reagent
Independent variable – glucose concentration
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
Volume of glucose solutions – using a syringe
Volume of Benedict’s reagent – using a syringe
Concentration of Benedict’s reagent – taking all samples from the same bottle
Surrounding temperature in water – using a water bath
Time given for color change in water bath – using a timer
Section 3 – Color change
Blue → Green → Yellow → Orange → Brick Red
How to know the concentration?
The concentration won’t be exactly known, but the color of the unknown solution will guide
us towards the approximate value. For example, if the color of 0.5% glucose was yellow, and
the color of the unknown solution was also yellow, then we know that the unknown solution
is near 0.5% concentration.
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Estimating the concertation of starch in an unknown solution:
Steps
1. Decide on a range of starch concentrations before beginning to work.[e.g. 0.25%,
0.5%, 1.0%, 1.5%, 2.0%]
2. Label 6 test tubes and the 5 beakers with the concentrations you’ve decided on, and
one test tube will be labeled with your control experiment
3. Using a clean syringe, distilled water, and 0.25% starch, fill the labeled beaker with 5
cm3 of starch solution. Repeat this for the four other concentrations.
4. Using a clean syringe, fill the 6 labeled test tubes with 0.5 cm3 of Iodine solution and
10 cm3 of distilled water
5. Add 5 cm3 of each starch solution to its corresponding labeled test tube, and add
5 cm3 of the control solution to the 6th test tube [e.g. potato]
6. Give each test tube a gentle shake to ensure the contents are well-mixed
7. Observe the test tubes and record the color change for each concentration
Section 1 – Independent and dependent variables
Dependent variable – color of Iodine solution
Independent variable – starch concentration
Section 2 – Control variables (variable – how it is controlled)
1. Volume of starch solutions – using a syringe
2. Volume of Iodine solutions – using a syringe
3. Concentration of Benedict’s solution – taking all samples from the same bottle
Section 3 – Color change
Red-Brown → Blue-Black
How to know the concentration?
This is sort of different from that of the glucose concentration. Here, the use of a colorimeter
may be required. This is due to the fact that color changes only once, and it is specifically
blue-black, so a colorimeter is used to figure out the intensity of blue-black for each
concentration, then the intensity of the blue-black in the control is compared to the
concentrations, and whichever is closest in intensity, will be closest in concentration to the
control experiment
Safety measures for both experiments (risk – how to prevent it)
1. Skin contact with benedict’s reagent – wear eye goggles, gloves, and a lab coat
2. Burns due to hot test tubes – use tongs to carry the test tubes when they’re hot
3. Toxicity of fruit juice/control solution – keep the fruit juice/control solution in a closed
container and do not taste it
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 2
Investigate the Vitamin C content of food and drink
Estimating Vitamin C content in an unknown solution
Steps
1. Decide on a range of Vitamin C concentrations, in order to compare them to the food
or drink [e.g. 0.25%, 0.5%, 1.0%, 1.5%, and 2.0%]
2. Start by using a syringe to draw up 5 cm3 of 1% DCPIP and shake it well, then add
1 cm3 of that DCPIP to a test tube, labeled with the desired concentration
3. Use a clean syringe to draw up 5 cm3 of 0.25% Vitamin C solution
4. Dropwise, add the Vitamin C solution to the test tube containing the 1 cm3 DCPIP,
and after each tube, shake the tube gently. Once you’ve noticed the blue color of the
DCPIP disappear, then stop adding drops, and record the volume of the Vitamin C
solution needed to do this
5. Repeat the steps 2-4 with the same concentration of Vitamin C solution, and record
the average volume of Vitamin C solution needed to decolorize the DCPIP
6. Repeat the steps 2-5 with other Vitamin C concentrations, recording the data in a
suitable table
7. Now, to start with your control experiment, add 1 cm3 of DCPIP to a test tube, and
use a syringe to draw up 5 cm3 of fruit juice, as your control experiment
8. Dropwise, add the fruit juice to the test tube containing the 1 cm3 DCPIP, and after
each drop, shake gently to ensure that the contents were mixed. Once you’ve noticed
the blue color of the DCPIP disappearing, then stop adding drops, and record the
volume of fruit juice needed to do so
9. Repeat steps 6-7 with the same fruit juice and record the average volume of fruit juice
needed to decolorize the DCPIP
Section 1 – Independent and dependent variables
Dependent variable – color of DCPIP
Independent variable – Vitamin C concentration
Section 2 – Control variables (variable – how it is controlled)
1. Volume of DCPIP – using a syringe
2. Concentration of DCPIP – obtaining the DCPIP from the same container
3. Volume of Vitamin C solution and fruit juice – using a syringe
Section 3 – Color change
Blue → Colorless
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IAS Biology
Unit 3
How to know the concentration?
This is different for this experiment specifically. You may be asked to use the following
formula in order to obtain the concentration of the control solution [in our example; fruit
juice], so long you have the needed data, and only the concentration is missing:
Concentration of Vitamin C in fruit juice =
Volume of standard solution X concentration of standard
Volume of fruit juice
solution
Key example: If the volume of the standard solution was 2 cm3 and the volume of the fruit
juice was 1.5 cm3, while the concentration of the standard solution was 1%, what would be
the concentration of Vitamin C in the fruit juice?
Concentration of Vitamin C in fruit juice =
2
1.5
X 10 = 13.3 mg
Safety measures (risk – how to prevent it)
1. Splash damage of hot water or DCPIP – wear eye protection
2. Skin contact with DCPIP and test tube solutions – wear gloves and a lab coat
3. Toxicity of fruit juice/control solution – keep the fruit juice/control solution in a closed
container and do not taste it
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 3
Investigate membrane properties including the effect of alcohol and temperature on
membrane permeability
Effect of temperature on membrane permeability
Steps
1. First of all, prepare eight water baths of a range of temperatures
[0,5°c,10°c,15°c,20°c,25°c,30°c,35°c]
2. Using a clean syringe, add 10 cm3 to eight test tubes (10 cm3 to each) and label them
with the temperature of the water bath in which they’ll be placed
3. Place each test tubes into its corresponding water bath, to acclimatize them, and start
a timer for 5 minutes. Check the temperature of each water bath using a
thermometer, as the temperature will not be exact, it’ll be approximate
4. Use a cork borer to cut 8 beetroot cylinders, and then use a knife and a white tile to
trim them all to the same length (e.g. 1 cm3)
5. Wash the beetroot cylinders with distilled water until water runs clean, and pat them
with a paper towel so that no pigment due to cutting stays on the cylinder.
6. Add one beetroot cylinder to each test tube in the water bath, and start a timer for 15
minutes
7. Shake the test tubes once, then discard the beetroot cylinder from each test tube, but
keep the liquid
8. Set the colorimeter to a blue/green filter and percentage transmission, making sure
you’ve zeroed the colorimeter using a blank cuvette filled with distilled water.
9. Transfer the liquid from each test tube into a cuvette, then place the cuvette inside the
colorimeter and take the percentage transmission reading. Record your results in a
suitable table
Section 1 – Independent and dependent variables
Dependent variable – percentage transmission/color intensity (membrane permeability)
Independent variable – temperature
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
6.
Volume of distilled water in each test tube – using a syringe
Time in water bath – using a stop clock
Diameter of beetroot cylinders – using cork borer
Length and width of beetroot cylinders – using ruler and a white tile
Age of beetroot cylinder – obtain from the same plant
Type of beetroot cylinder – obtain from the same tissue
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Effect of alcohol on membrane permeability
Steps
1. Add 10 cm3 of ethanol to 5 test tubes, of different concentrations [e.g. 0.2%, 0.4%,
0.6%, 0.8%, 1.0%]
2. Use a cork borer to extract 5 beetroot cylinders, then trim them to the same size using
a knife and a white tile (e.g. 1 cm). Afterwards, wash the cylinders with distilled
water and pat them gently with a paper towel, to make sure that none of the pigment
due to cutting stays
3. Add a beetroot cylinder to each of the test tubes, then leave them for 15 minutes to
soak in the ethanol
4. Shake the test tubes once, then discard the cylinders from the test tubes but keep the
liquid
5. Set the colorimeter to blue/green filter and percentage transmission, making sure
you’ve zeroed it using a blank cuvette filled with distilled water
6. Transfer the liquid from each test tube to a cuvette, and place the cuvette in the
colorimeter, taking a percentage transmission reading. Record your results in a
suitable table.
Section 1 – Independent and dependent variables
Dependent variable – percentage transmission/color intensity (membrane permeability)
Independent variable – alcohol concentration
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
6.
Volume of distilled water in each test tube – using a syringe
Volume of ethanol – using a syringe
Diameter of beetroot cylinders – using cork borer
Length and width of beetroot cylinders – using ruler and a white tile
Age of beetroot cylinder – obtain from the same plant
Type of beetroot cylinder – obtain from the same tissue
Safety measures (risk – how to prevent it)
1. Water baths above 50°c may scald – keep away from the water bath when opening it
as to not get splashed with hot water or burnt by steam, and wear eye goggles
2. Sharp items such as cork borer and knives may injure your hand – always cut or push
downwards onto the tile, and wear gloves if necessary
3. Ethanol is highly flammable – keep ethanol away from naked flames, and always keep
stoppers on ethanol bottles
4. Electric plugs – do not handle electric plugs or switches with wet hands
5. Contamination – always wash hands with soap when you’re done working
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IAS Biology
Unit 3
How to see the effect?
In order to see the effect of temperature on membrane permeability, we’ll be using tables and
graphs as shown below. The higher the transmission or color intensity, the greater the effect
and the more permeable the membrane. As a rule of thumb for most cases, as the
temperature or ethanol concentration increases, so will the permeability of the membrane,
because high temperatures denature the shape of proteins and enzymes in the membrane,
making it more permeable, and high alcohol concentrations cause the phospholipids in the
cell membrane to become emulsified.
0 / 0.2
Temperature/°c or Ethanol concentration/%
10 / 0.4
20 / 0.6
Percentage
transmission
of membrane
Mean
transmission
of membrane
Temperature's effect on membrane permeability
Percentage transmission
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0
Temperature/°c or Ethanol concentration
In the exam, you may be asked to fill in data, draw graphs, or even title graphs. This page
gives you an idea about how your work should look like, given that the independent variable
is the same (however, it usually is not)
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 4
Investigate the effect of temperature, pH, enzyme concentration and substrate
concentration on the initial rate of enzyme-catalyzed reaction
Effect of temperature on the initial rate of enzyme-catalyzed reaction
Steps
1. Prepare five water baths of a range of your desired temperatures [e.g. 10°c, 20°c,
30°c, 40°c, 50°c], and use a thermometer to ensure that the water baths are of the
correct temperatures
2. Add 2 cm3 of 1% trypsin to 5 test tubes, and label each test tube with a temperature
from your range [i.e. first test tube labeled with 10°c], and place each test tube to its
corresponding water bath
3. Add 2 cm3 of milk to 5 different test tubes, and label each test tube with a
temperature from your range [i.e. first test tube labeled with 10°c], and place each
test tube to its corresponding water bath
4. Leave the test tubes for 5 minutes so that they reach the required temperature
5. Before taking out the test tubes, zero the colorimeter using a solution of 2 cm3 of 1%
trypsin and 2 cm3 of distilled water in a cuvette
6. Take out the test tubes and pour 2 cm3 of the test tube containing milk of the first
temperature into the 2nd cuvette
7. Take out the test tube and pour 2 cm3 of the 1% trypsin into the same cuvette
containing the milk, from the same water bath as that of the test tube containing milk.
Then mix the two solutions quickly and place your solution into the colorimeter
8. Start the data logger and measure the color absorbance immediately, then keep
measuring the absorbance once every 15 seconds for 5 minutes or until there is no
significant change in readings
9. Rinse the cuvette, and repeat the steps 6-8 for the other four temperatures making
sure you’ve zeroed the colorimeter before taking the readings, then record your data
in a suitable table
Section 1 – Independent and dependent variables
Dependent variable – initial rate of enzyme-catalyzed reaction/enzyme activity
Independent variable – temperature
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
6.
Volume of trypsin in each test tube – using a syringe
Concentration of trypsin in each test tube – way of preparation
Concentration of milk suspension - way of preparation
Volume of milk – using a syringe
Time in water bath – using a stop clock
pH of the solution – using a buffer
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Effect of pH on the initial rate of enzyme-catalyzed reaction
Steps
1. Select a range of buffer solutions (the pH range) which you want to investigate in
relation to the enzyme activity [e.g. 4,5,6,7,8]
2. Start off by placing 1 cm3 of buffer, 2 cm3 of distilled water, and 1 cm3 of trypsin
solution into a cuvette and use it in order to zero the colorimeter.
3. Pour 1 cm3 of trypsin, 1cm3 of your first buffer solution, and 2 cm3 of milk into a
cuvette and working quickly, mix the solution [buffer and trypsin] and the milk b
shaking gently, and place the cuvette into the colorimeter and start the data logger
4. Measure the absorbance immediately, then once every 15 seconds for 5 minutes or
until there is no significant change in readings
5. Rinse the cuvette and repeat the steps 2-4 for the other pH values, but remember to
zero the colorimeter using the solution in step 2 before taking the readings, and
record your data in a suitable table
Section 1 – Independent and dependent variables
Dependent variable – initial rate of enzyme-catalyzed reaction/enzyme activity
Independent variable – pH
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
Volume of trypsin in each test tube – using a syringe
Concentration of trypsin in each test tube – way of preparation
Volume of milk – using a syringe
Temperature – using a water bath, an AC unit, or a thermostat
Concentration of milk suspension - taking the milk suspension samples from the same
container and through the way of preparation
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IAS Biology
Unit 3
Effect of enzyme concentration on the initial rate of enzyme-catalyzed reaction
Steps
1. First of all, check your dilution calculations and decide on a range of enzyme
concentrations [e.g. 0.2%, 0.4%, 0.6%, 0.8%, 1.0%], then use a solution of 2 cm3 of
trypsin solution and 2 cm3 of distilled water in a cuvette to zero the colorimeter
2. Working quickly, mix 2 cm3 of milk suspension and 2 cm3 of the 0.2% trypsin in the
second cuvette, and place the cuvette in a colorimeter, and start the data logger
3. Measure absorbance immediately, then once every 15 seconds for 5 minutes or until
there is no significant change in results
4. Rinse the cuvette that you’ve used for the first solution with distilled water, and repeat
the steps 2-4 for the 4 other concentrations, making sure you’ve zeroed the
colorimeter before taking the recordings, and record your data in a suitable table
Section 1 – Independent and dependent variables
Dependent variable – initial rate of enzyme-catalyzed reaction/enzyme activity
Independent variable – enzyme concentration
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
Volume of trypsin in each test tube – using a syringe
Volume of milk – using a syringe
Temperature – using a water bath, an AC unit, or a thermostat
pH – by the use of a buffer
Concentration of milk suspension – way of preparation
Effect of substrate concentration on the initial rate of enzyme-catalyzed reaction
Steps
1. First of all, check your dilution calculations and decide on a range substrate
concentrations, in this case, the milk [e.g. 0.2%, 0.4%, 0.6%, 0.8%, 1.0%], then use
a solution of 2 cm3 of distilled water and 2 cm3 of 1% trypsin in a cuvette to zero the
colorimeter reading
2. Working quickly, mix 2 cm3 of 1% trypsin and 2 cm3 of 0.2% milk suspension in a
cuvette, and place the cuvette in a colorimeter and start the data logger
3. Measure absorbance immediately, then once every 15 seconds for 5 minutes or until
there is no significant change in readings
4. Rinse the cuvette that you’ve used for the first solution with distilled water and repeat
the steps 2-4 for the 4 other concentrations, making sure you’ve zeroed the
colorimeter before taking the readings, and record your data in a suitable table.
Section 1 – Independent and dependent variables
Dependent variable – initial rate of enzyme-catalyzed reaction/enzyme activity
Independent variable – substrate concentration
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IAS Biology
Unit 3
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
5.
Volume of trypsin in each test tube – using a syringe
Volume of milk – using a syringe
Temperature – using a water bath, an AC unit, or a thermostat
pH – by the use of a buffer
Concentration of trypsin – way of preparation
Safety measures (risk – how to prevent it)
1. Any trypsin splashing onto your eyes or body – wear goggles, a lab coat, and gloves
2. Inflammation on skin due to trypsin – wash your skin with cold running water
3. Water baths above 50°c may scald – keep away from the water bath when opening it
as to not get splashed with hot water or burnt by steam, and wear eye goggles
How to display the data?
As we’ve said before, for each experiment, you’ll record your results in a table, and
afterwards, you’ll plot them on a graph and see the trend or effect of the independent
variable on the initial rate of enzyme activity. Below, we’ll take temperature as an example,
Note: This table should work for all independent variables (the numbers are imaginary)
Initial rate of enzyme
activity
Temperature/°c
0
10
20
30
40
50
Initial rate of enzyme activity
0
0.1
0.2
0.25
0.3
0.3
Effect of temperature on initial
rate of enzyme activity
0.4
0.3
0.2
0.1
0
0
20
40
60
Temperature
To measure the
rate of reaction,
draw a line at the steepest point and use the following formula to figure out the gradient,
which gives you the initial rate of reaction
Change in Y-axis
Change in X-axis
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IAS Biology
Unit 3
Gradient =
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 5 & 7
(I) Use a light microscope to make observations and labelled drawings of suitable animal cells
(II) Use a graticule with a microscope to make measurements and understand the concept of scale
(III) Use a light microscope to make observations of transverse sections of roots, stems, leaves, plant
tissues
(IV) Identify Sclerenchyma fibers, Phloem, Sieve tubes, Xylem vessels, and their location
Before we begin the steps for the core practical, we’ll need to familiarize ourselves with the
inner workings of the microscope. First of all, there are mainly two types of microscope, the
light microscope and the electron microscope. Below is a comparison between the 2
microscopes:
Category
Magnification
Resolution
Cost
Specimen
Observation to view the
image
Light Microscope
1500x
0.2 μm
Cheap
Usually dead; can sometimes be
alive
Using a light beam from
underneath or light from the sun
Electron Microscope
500 000x
0.1-1.0 nm
Expensive
Always dead due to being
placed in a vacuum
Uses a beam of electrons
scattered by the specimen
The light microscope
A light microscope is usually used to look at specimens in institutions such as schools, as it is
cheap, reliable, and can give a good image of the specimen. When using the light microscope,
you’ll need to stain the specimen so that it is visible. There are a number of stains used:
1.
2.
3.
4.
Hematoxylin – stains the nuclei of animal and plant cells
Methylene blue – stains the nuclei of animal cells
Acetocarmine – stains the chromosomes in dividing nuclei in animals and plants
Iodine – stains starch containing material
There are specific advantages and disadvantages to the light microscope
Advantages of light microscope
Can see parts of living animals or plants, which is
useful when comparing prepared slides with living
tissue
Relatively cheap
Relatively light and portable; can be used anywhere
Disadvantages of light microscope
Preserving and staining tissue can produce
artifacts; which are not part of the living tissue,
but occur due to the process of preserving and
can be mistaken for living tissue
Limited powers of resolution and magnification
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IAS Biology
Unit 3
In examinations, you may be asked to calculate the magnification of a certain image, if you’ve
been given the real measurement, in order to do this, use the following formula:Magnification =
Image size
Actual size
The electron microscope
The electron microscope is usually used in large labs and scientific institutions to look at the
detailed ultrastructure of specimen, and when using it, specimen must be in vacuum in order
for the microscope to work, as air would scatter the electrons and produce a blurred image of
the tissue, so they’ll always be dead. When staining the specimen, heavy metal ions such as
uranium and lead are used. Afterwards, the microscope will produce two types of electron
micrographs, which are:
1. Transmission electron micrographs (TEMs) – two dimensional images, similar to those
from the electron microscope, but clearer
2. Scanning electron micrographs (SEMs) – have a lower magnification than TEMs, but
show images in three dimensions
The electron microscope also has some advantages and disadvantages
Advantages of light microscope
Huge powers of magnification and resolution; allow us
to see the exact ultrastructure of specimen
Disadvantages of light microscope
All specimen must be examined in vacuum, so it
is impossible to observe live tissue
Specimen undergo extreme treatment which may
produce artifacts
Extremely expensive
The instrument is large, must be kept at constant
temperature and pressure, and is expensive to
maintain
(I) Calibration of the graticule
Here, you’ll have to compare the graticule scale with the micrometer scale in order to figure
out the length of one eyepiece unit, using the following steps:
1. Place a micrometer slide on the stage of the microscope, and
focus on the micrometer scale using the low-power objective lens
2. Move the slide and rotate the eyepiece to align the scales of the
eyepiece graticule and the micrometer scale in the field of view
3. Count the number of divisions on the eyepiece graticule and
compare them to a known length on the micrometer scale to
figure out the length of one eyepiece unit
4. Repeat steps 1-3 with the medium-power and high-power
objective lens
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IAS Biology
Unit 3
(II) Making observations
Once you’ve calibrated the graticule, you’ll be able to observe the cells, through the following
steps:
1. Wash your hands with soap and water, then take a cotton bud and rub it gently onto
the inside of your cheek to obtain somatic cells, then rub it in a circle on a microscopic
slide, and place the cotton bud in a beaker of disinfectant solution
2. Add a few drops of methylene blue to the sample on the microscopic slide, then cover
it with a cover slip
3. Turn the objective lens to low power, and examine the stained slide under the
microscope. In order to do this, bring the lens as close to the slide as possible, while
watching from the slide instrument, and look through the eyepiece using the coarse
focusing knob to focus, while moving the lens away from the stage as to not damage
the slide and the lens. Lastly, use the fine focus until a clear view of the cells is
established
4. Carefully sketch a few cells
5. Use an eyepiece graticule to measure the cell’s diameter. Add the scale bar, a title,
and the total magnification to your sketched diagram. The total magnification is not
the same as the magnification of the drawing
Total magnification = magnification of objective lens x magnification of eyepiece lens
6. Afterwards, turn the objective lens onto medium-power, and focus until the cells are
clear and distinct, identifying as many details as you can. Finally, turn the objective
lens onto the high-power, and focus until the details are as clear as possible, using the
fine-focusing knob, then draw the details of the cell as accurately as possible.
7. Measure the length and breadth of two cells and include the measurements in your
sketched diagram, then turn off the microscope and place the microscopic slide in a
beaker of disinfectant solution
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IAS Biology
Unit 3
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
(III & IV) Using a light microscope to observe transverse sections of plant parts, and identify
plant tissue such as sclerenchyma fibers, phloem, sieve tubes, xylem vessels, and their location
Here, we won’t be examining animal cells, instead we’ll be observing plant cells and
identifying their tissues instead, through the following steps:
1. First of all, prepare thin sections of plant stem by adding a few drops of water to a
white tile, wetting your razor to reduce friction, and holding the plant cell firmly while
keeping away from the edge of the razor, and cutting several thin transverse sections.
Transfer the sections to a water-containing watch glass
2. Choose the thinnest transverse section, place it on a microscopic slide, add one drop
of water and remove any excess water by touching the edge of the stem with
absorbent paper
3. Wearing gloves and eye protection, add two drops of toluidine blue O stain to the
stem and leave it for 2-4 minutes. Then, add a cover slip and gently remove excess
stain using a paper towel
4. Turn the objective lens to low power, and examine the stained slide under the
microscope. In order to do this, bring the lens as close to the slide as possible, while
watching from the slide instrument, and look through the eyepiece using the coarse
focusing knob to focus, while moving the lens away from the stage as to not damage
the slide and the lens. Lastly, use the fine focus until a clear view of the cells is
established
5. Staying on low-power or medium-power objective lenses, annotate a diagram of the
section while showing the arrangement of the tissues, without drawing any cell details
6. Repeat steps 1-5 using a root piece
7. To make a leaf cross section, place the leaf between two thin polystyrene blocks, and
gently stroke the polystyrene and the leaf with your razor to stroke off 4 or 5 thin
sections
8. Select the thinnest section, and place it on a microscopic slide, then add 1 or 2 drops
of water and cover it using a cover slip
9. Repeat steps 4 and 5 with the leaf
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IAS Biology
Unit 3
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Safety measures (risk – how to prevent it)
1. Methylene blue may be harmful when swallowed – handle the stain with care
2. Skin contact with methylene blue, iodine, or toluidine blue O – wear gloves, goggles,
and a lab coat, and if any spills happen, clean them immediately
3. Biohazardous material and cotton buds – these could be harmful, so they should be
placed in disinfectant solution immediately after use, and cotton buds should be used
once by one person only
4. Daylight illumination with light microscope – do not use a light microscope with
daylight illumination under the sunlight, as it might strike your eye and cause
blindness
5. Razor blades and mounted needles – take care when using them and always cut away
from oneself
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 6
Prepare and stain a root tip squash to observe the stages of mitosis
Observing mitosis under the microscope
Steps
1. Fill a small bottle with 1mol dm-3 of hydrochloric acid and place it in a water bath of
temperature 55°c and leave it for 15 minutes to allow the acid to warm up
2. Place a garlic clove at the top of the bottle making sure the roots are submerged, and
leave it at 55°c for 5 minutes in order to soften the roots
3. Take out the garlic clove and rinse it with water to remove any excess HCL, then use
a pair of sharp scissors to cut the root tips to 5-10 mm pieces making sure they fall
onto a watch glass. Afterwards, add a drop of acetic orcein and cover the watch glass
with a lid that is slightly loose (to prevent ejection of liquid during heating), then heat
the watch glass in a 55°c water bath for 5 minutes to intensify the color
4. Remove the root tips from the watch glass, place them onto the microscopic slide and
tease them apart using a mounted needle, then add one drop of acetic orcein [you
may heat the slide to intensify the color], then add a drop of water to wash off any
excess acetic orcein. Cover the slide with a cover slip
5. Wrap the slide with several layers of a paper towel and gently squash the root tips
while not twisting as to not break the cover slip
6. Place the root tips onto the stage of the microscope, and start with the low power,
while adjusting the focus and centering the cells in the field of view
7. Move up to the high power, and identify as many stages of mitosis as you can. Count
the number of cells undergoing each stage of mitosis, as well as interphase, while
recording your results in a suitable table, you can also calculate the mitotic index
Mitotic index =
Number of cells in mitosis
Total number of cells
8. Draw and annotate one cell from each of the stages you have identified, making simple
outlines of the cells and a group of chromosomes in them. Try to show the relative size
and positions of chromosomes in the cells, and include a small description of what is
happening in the drawing
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Safety measures (risk – how to prevent it)
1. Acetic orcein – acetic orcein is corrosive and causes burns, so always wear goggles, a
lab coat, and if exposed to orcein, wash your skin for 10 minutes
2. HCL – avoid skin contact with HCL and wear gloves
3. Water baths above 50°c may scald – keep away from the water bath when opening it
as to not get splashed with hot water or burnt by steam, and wear eye goggles
4. Daylight illumination with light microscope – do not use a light microscope with
daylight illumination under the sunlight, as it might strike your eye and cause
blindness
5. Glassware and Scissors – take care when using them and always cut away from
oneself
6. Electric plugs – keep away from them when your hands are wet, and shut them off
when you aren’t using them
Risk assessment
With this experiment, you may need to do a risk assessment, as follows
The risk will be placed in the appropriate square for it based on your evaluation and previous
experience
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 8
Determining the tensile strength of plant fibers
Measuring tensile strength
Steps
Tensile strength is the measure of how much stress or weight a plant fiber can handle before
breaking. We can measure it for plant fibers using the following steps (for the purpose of being
clear, we’ll use celery stalks as an example). It is important to know the graph of tensile strength
1. Remove nine fibrous strings from celery stalks, using a knife and a white cutting tile
surface
2. Inspect the strings to make sure there are no cuts or breaks, and that all fibers have a
constant diameter along them. Divide the 9 fibers into 3 groups [3 fibers 10 cm long, 3
fibers 15 cm long, and 3 fibers 20 cm long].
3. Carefully clamp the 10 cm string between the two stands shown below, ensuring it is
held securely, place cushioning under the fiber, and start adding weights 10g at a time
until the string breaks. Record the mass required to break the fiber
4. Repeat the steps for the 2 other 10 cm fibers and record the mean mass needed to break
the fiber, then repeat the experiment for the other fibers
Section 1 – Independent and dependent variables
Dependent variable – breaking strength/tensile strength
Independent variable – weight of masses
Section 2 – Control variables (variable – how it is controlled)
1.
2.
3.
4.
Plant from which the fiber was obtained – same tissue and plant
Length of fiber – using a ruler
Temperature – using a water bath, an AC unit, or a thermostat
Diameter of fiber – using a Vernier caliper to cut them all to the same diameter
Safety measures (risk – how to prevent it)
1. Weights slamming the bench or feet – place cushioning under the middle of the fiber
2. Knife and spring scale – take care when using them and always cut away from oneself
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
CORE PRACTICAL 9
Investigate the antimicrobial properties of plants, including aseptic techniques for
the handling of bacteria
Investigating antimicrobial properties
Steps
Before we start, at all times you should follow aseptic techniques, meaning you’ll keep a
yellow Bunsen burner beside you, and disinfect anything you use
1. Wash your hands with soap and water, disinfect your bench and leave it to dry for 10
minutes then wipe it
2. Place a piece of garlic and use a mortar and pestle to grind it into paste, then add 10
cm3 of alcohol to it to make an extract
3. Place a paper disc in the mortar to soak up the garlic and alcohol solution, then place
the paper disc on an agar plate to dry
4. Using a marker pen, mark the 4 sides of the agar plate seeded with bacteria, one side
with garlic, one side with mint, and 2 with control.
5. Open the lid of the Petri dish containing the bacteria-seeded agar away from yourself
and only slightly, then use your forceps to place the paper disc containing the garlic
extract in the correct quarter, and close the dish.
6. Clean the mortar and pestle, then repeat steps 2-6 with mint
7. Add a small amount of alcohol to a small beaker, and sock to paper discs in the beaker,
then remove them and place them on a sterile agar plate to dry
8. Open the lid of the bacteria seeded Petri dish away from yourself and only slightly, and
place the two paper discs in the control labelled quarters
9. Place one small piece of tape on each quarter, but don’t cover the dish completely
10. Invert the plate and incubate it at 27°c for 24 hours.
11. Afterwards, use a ruler to measure the inhibition zone caused by the extracts, and record
your data in a table
12. Clear all of your equipment and sterilize the Petri dishes which you used.
Section 1 – Independent and dependent variables
Dependent variable – plant extract
Independent variable – zone of inhibition
Section 2 – Control variables (variable – how it is controlled)
1. Time of soaking – stop clock
2. Size of paper disc – taking them from the same container
3. Temperature – incubator
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi
IAS Biology
Unit 3
Safety measures (risk – how to prevent it)
1.
2.
3.
4.
5.
Microorganisms are hazardous – only open the Petri dish slightly when performing the
experiment, wash your hands before and after, and keep a yellow Bunsen burner flame
beside you to prevent contamination
Inoculated plates – do not open them, use the alternative plate provided for you
Ethanol – do not decant it near a naked flame and use stoppers when not using ethanol
Contamination – use aseptic techniques when transferring the bacteria to the Petri
dishes, disinfect your bench when your done working, and any contaminated Petri
dishes should not be opened, and should be destroyed by a lab technician
Skin contact with disinfectant – carefully handle the disinfectant and wear gloves
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© 2018 – Arranged and Edited by: Abdulrahman Tabaza and Omar Minawi