Chapter 1 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Eye protection should be worn at all times.
Practical 1.1
A: Calibrating an eyepiece graticule
B: Preparing a slide of onion epidermal cells
It is recommended that a ready-made solution of 2% iodine in potassium iodide is purchased.
The ready-made solution is low hazard.
Practical 1.2
Preparing a slide of human cheek cells
It is recommended that a ready-made solution of 0.1% methylene blue is purchased. The ready-made
solution is low hazard.
Cotton buds, slides and cover slips should all be transferred to a 5% bleach (sodium hypochlorite)
solution immediately after use, and left there for at least 15 minutes. Alternatively, domestic chlorinebased bleach can be used. This will usually have a concentration of less than 5% and therefore, a lower
hazard rating. It should be labelled ‘irritant’.
Slides and cover slips can then be washed for reuse, following normal procedure. Used cotton buds,
once disinfected, can be disposed of in a sealed bag with the normal refuse.
Practical 1.3
Preparing a slide of Elodea leaf cells
It is recommended that a ready-made solution of 2% iodine in potassium iodide is purchased.
The ready-made solution is low hazard.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
If Elodea is not available, teachers can find a suitable locally available species of aquatic plant. An
alternative is to use leaves of moss plants.
Practical 1.4 Preparing slides of potato tubers and banana fruit to investigate
starch grains
It is recommended that a ready-made solution of 2% iodine in potassium iodide is purchased.
The ready-made solution is low hazard.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 1.1
A: Calibrating an eyepiece graticule
B: Preparing a slide of onion epidermal cells
Safety
Wear eye protection.
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
microscope
eyepiece graticule
stage micrometer
sharp knife or scalpel
small onion (Allium cepa)
slides and cover slips
•
•
•
•
•
•
forceps
mounted needle
2% iodine in potassium iodide solution
dropping pipette
filter paper
eye protection
Introduction
In this practical, you will:
• calibrate an eyepiece graticule
• make a temporary preparation of some onion epidermal cells
• stain the cells so that you can see structures within them
• set up a light microscope and use it to make observations and measurements
• make a drawing of the onion cells
• use the eyepiece graticule you calibrated to measure the size of the cells.
Procedure
A
Calibrating an eyepiece graticule
1
If it does not already have one, insert a graticule into the eyepiece of the microscope by
unscrewing the top lens, resting the graticule on the rim halfway down and replacing the top lens.
2
Place a stage micrometer slide on the stage of the microscope. The smallest division on the stage
micrometer equals 100 m.
3
Using the low-power objective, focus the microscope on the stage micrometer. Rotate the eyepiece
and move the slide to superimpose the scales of the eyepiece graticule and the stage micrometer
(shown in the diagram on the next page).
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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4
Count the number of divisions on the eyepiece graticule equivalent to 100 m on the stage
micrometer and then calculate the length that one eyepiece division is equivalent to. For example,
if three divisions are equal to 100 m, then each division is equal to 33.3 m at low power. Record
your answer.
5
Repeat step 4 for the medium-power and high-power objectives. You have now calibrated the
eyepiece graticule and you can use it to measure cells in the preparation below, or in the
preparations in the next practicals.
B
Preparing a slide of onion epidermal cells
1
The fleshy layers inside an onion are known as ‘leaves’ and store nutrients. Cut open an onion and
separate some of the leaves. Peel off one of the thin layers of epidermal tissue from the inner
concave surface of a leaf and transfer it to a drop of water on a microscope slide. Use forceps and a
mounted needle to make sure that the tissue is not folded.
2
Place two drops of iodine solution onto the tissue. Gently lower a coverslip onto the slide using a
mounted needle. Use a piece of filter paper to absorb excess stain. Place some filter paper over the
coverslip and gently press to flatten the specimen.
3
Place the slide on the stage of the microscope and use the low-power objective to locate the cells.
Now use the high-power objective to select three adjacent cells that are clearly visible in your field
of view.
4
Make a large, labelled drawing of these three epidermal cells. Use a sharp pencil (HB) and a ruler
to draw the label lines and labels.
5
Use the eyepiece graticule to measure the length of one of the epidermal cells that you have drawn.
Now measure the same cell in your drawing.
6
Calculate the magnification of your drawing, using the formula:
magnification =
length of drawing of cell
actual length of cell
Remember that both lengths must be measured in the same units, e.g. micrometres (m). Write the
magnification underneath your drawing.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 1.2
Preparing a slide of human cheek cells
Safety
Wear eye protection.
Take care when using sharps.
The disinfectant is an irritant. If it contacts the skin, wash off immediately with plenty of water.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
microscope
calibrated eyepiece graticule
slides and cover slips
forceps
mounted needle
dropping pipette
•
•
•
•
•
filter paper
cotton bud
small beaker of disinfectant
0.1% methylene blue solution
eye protection
Introduction
In this practical, you will:
• make a temporary preparation of human cheek cells
• stain the cells so that you can see structures within them
• set up a light microscope and use it to make observations and measurements
• make a drawing of the cheek cells
• use an eyepiece graticule to measure the size of the cells.
Procedure
1
Use a clean cotton bud to gently scrape some material from the lining inside one of your cheeks.
Smear the material onto a clean dry slide and immediately put the cotton bud into a beaker of
disinfectant.
2
Place two drops of methylene blue solution onto the material on the slide. Place a cover slip over
the stain and lower the cover slip gently onto the specimen using a mounted needle. Use a piece of
filter paper to absorb excess stain. Place some filter paper over the cover slip and gently press to
flatten the specimen.
3
Place the slide on the stage of the microscope and use the high-power objective to select three cells
that are clearly visible in your field of view. Make a large, labelled drawing of these cells.
4
Use the eyepiece graticule that you calibrated in Practical 1.1 to measure the diameter of one of the
cells that you have drawn. Now measure the same cell in your drawing. Calculate the
magnification of your drawing, using the formula:
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
1
magnification =
length of drawing of cell
actual length of cell
Remember that both lengths must be measured in the same units e.g. micrometres, (m). Write the
magnification underneath your drawing.
5
When you have finished with your slide, place it in the beaker of disinfectant.
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
2
Practical 1.3
Preparing a slide of Elodea leaf cells
Safety
Wear eye protection.
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
microscope
calibrated eyepiece graticule
scalpel
piece of Elodea plant (Canadian pondweed)
slides and cover slips
forceps
•
•
•
•
•
•
mounted needle
dropping pipette
distilled water
2% iodine in potassium iodide solution
filter paper
eye protection
Introduction
In this practical, you will:
• make a temporary preparation of Elodea leaf cells
• set up a light microscope and use it to make observations and measurements
• make a drawing of the leaf cells
• use an eyepiece graticule to measure the size of the cells.
Procedure
1
Use forceps to remove a single leaf from a piece of Elodea canadensis plant. Place the leaf on a
clean microscope slide and cut a small square of the leaf using a sharp scalpel. Elodea leaves are a
curled shape and it is important to prepare a flat piece of leaf to use in the preparation: it should be
about 3 mm wide. Try to use the edge of a leaf, avoiding the thicker midrib.
2
Transfer the piece of leaf to another microscope slide. Add two drops of distilled water to the
specimen. Place a cover slip over the water and lower it gently onto the specimen using a mounted
needle. Use a piece of filter paper to absorb excess water. The cover slip must lie flat.
3
Observe the specimen under the microscope, using low-power, medium-power and then highpower objectives. Select an area where cells are clearly visible. You should be able to see the cell
walls and many chloroplasts. More chloroplasts will be visible around the edges of the cells, just
inside the cell walls. This is because the middle of each cell is occupied by the sap vacuole. If you
watch a cell for a few minutes, you may see the chloroplasts moving in the cell as a result of
cytoplasmic streaming. (Under the microscope this appears faster than normal, due to the extra
kinetic energy provided by the heat from the lamp.)
4
It is difficult to see nuclei in unstained leaf cells. You could try staining the cells with iodine to
show up the nuclei. Repeat steps 1–3 using iodine solution instead of water.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
5
Use the high-power objective to select a few leaf cells. Make a large, labelled drawing of these
cells. Use the eyepiece graticule that you calibrated in Practical 1.1 to measure the length of one of
the cells that you have drawn. Now measure the length of this cell in your drawing.
6
Calculate the magnification of your drawing, using the formula:
length of drawing of cell
actual length of cell
Remember that both lengths must be measured in the same units, e.g. micrometres (m). Write the
magnification underneath your drawing.
magnification =
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 1.4
Preparing slides of potato tubers and banana fruit to investigate
starch grains
Safety
Wear eye protection.
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
microscope
calibrated eyepiece graticule
scalpel
potato tuber (Solanum tuberosum)
banana fruit (Musa sp.)
slides and cover slips
forceps
•
•
•
•
•
•
mounted needle
dropping pipette
distilled water
2% iodine in potassium iodide solution
filter paper
eye protection
Introduction
In this practical, you will:
• make a temporary preparation of cells from a potato tuber and a banana to show starch grains
• set up a light microscope and use it to make observations and measurements
• stain the starch grains using the irrigation technique
• make a drawing of the cells
• use an eyepiece graticule to measure the size of the starch grains.
Procedure
1
Use a mounted needle or a scalpel to scrape some tissue from the cut surface of a potato. Place the
tissue on a clean microscope slide and add two drops of distilled water. Place a cover slip over the
water and lower it gently onto the specimen using a mounted needle. Use a piece of filter paper to
absorb excess water.
2
Observe the potato cells under the high-power objective of the microscope. You should see the cell
walls of the potato cells. Each cell contains a number of rounded starch grains, called amyloplasts.
3
Stain the starch grains with iodine solution using a technique called irrigation (see diagram on the
next page). Place a drop of iodine solution on the slide so that it touches the edge of the cover slip.
Use a piece of filter paper to draw the iodine across the slide under the cover slip, by touching the
edge of the cover slip on the side opposite the iodine. The starch grains will stain blue with the
iodine.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
4
Use the high-power objective to select a few potato tuber cells. Make a large, labelled drawing of
these cells and their starch grains. Use the eyepiece graticule calibrated in Practical 1.1 to measure
the length of one of the cells that you have drawn. Now measure the length of this cell in your
drawing.
5
Calculate the magnification of your drawing, using the formula:
magnification =
length of drawing of cell
actual length of cell
Remember that both lengths must be measured in the same units, e.g. micrometres (m). Write the
magnification underneath your drawing.
6
Repeat steps 1–5 using tissue from the middle of a banana fruit, which also contains starch.
Compare the shape of the grains in the two tissues. Starch grains in banana are more elongated in
shape than those in potato.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 2 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Eye protection should be worn at all times.
Practical 2.1 Tests for biological molecules
It is recommended that ready-made solutions of 2% iodine in potassium iodide, Benedict’s solution
and biuret reagent are purchased. The ready-made solutions are low hazard.
Sodium hydroxide solution is an irritant, and hazardous to eyes even at low concentrations.
Ethanol and its vapour are highly flammable – keep them well away from sources of ignition.
In the Benedict’s test, tubes should be heated in a thermostatic water bath or in a beaker of water
above a Bunsen burner. They should not be heated directly in a Bunsen flame, which is likely to cause
the contents to jump out of the tube. Using a thermostatic water bath avoids the need to use a Bunsen
burner altogether. The test will work at temperatures just below boiling point, so for added safety the
electric water bath could be maintained at 90 °C.
The sucrose should be uncontaminated with glucose. Use analytical reagent (AR) sucrose rather than
granulated sugar.
The biuret test only works on soluble proteins. If separate biuret solutions are used, students should
note the darker blue colour produced when the two solutions are mixed, which is not to be confused
with the lilac colour of a positive biuret test.
The iodine test for starch only works with amylose, where the helices of amylose trap the iodine
molecules forming a blue–black compound. It does not work with amylopectin, which is composed of
much shorter helices that are unable to trap the iodine. Iodine will also stain glycogen, but the colour
produced is a dark red–brown.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 2.2
Identifying three biological molecules
It is recommended that ready-made solutions of 2% iodine in potassium iodide, Benedict’s solution
and biuret reagent are purchased. The ready-made solutions are low hazard.
Risk assessment should consider the problem of allergic responses to enzymes.
The tubes should be set up as follows:
• 30 cm3 of 0.1% starch solution in tube A
• 30 cm3 of 1% sucrose solution in tube B
• 30 cm3 of 1% sucrase solution in tube C.
Both sucrase and sucrose solutions should be uncontaminated with glucose. Use AR sucrose rather
than granulated sugar, and pure sucrase rather than invertase.
Students should use the reagents to identify A as starch, B as non-reducing sugar and C as protein.
C is therefore the enzyme.
If students mix C (sucrase) with B and test the mixture for reducing sugars, the test should be positive
due to the hydrolysis of the sucrose into glucose and fructose.
If students then mix A and C, and test the mixture for presence of reducing sugars, the test will be
negative, confirming that C is not amylase.
Practical 2.3 Semi-quantitative and quantitative tests for reducing sugars
It is recommended that a ready-made Benedict’s solution is purchased. The ready-made solution is
low hazard.
The unknown glucose solutions should be selected so that they are within the range of the measured
standards.
Students will probably need a second lesson to complete part C. It is only possible to carry out this
assay on a limited range of glucose concentrations. This is an extension activity to test more able
students, particularly in step 5.
The absorbance scale on the colorimeter must be used. The colorimeter should be zeroed with the
blank (that is, the filtrate from the 2.5% glucose solution) each time a reading is taken.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 2.1
Tests for biological molecules
Safety
Wear eye protection.
Take care when using heating apparatus.
Dilute sodium hydroxide solution is an irritant. If it contacts the skin, wash off immediately with
plenty of water.
Ethanol is highly flammable. Keep it well away from sources of ignition.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
six test tubes
test tube rack
labels or marker pen
thermostatically controlled water
bath at 100 °C
thermometer
test tube holder
dropping pipettes
spotting tile
10 cm3 of 0.1% starch solution
10 cm3 of 1% glucose solution
•
•
•
•
•
•
•
•
•
•
20 cm3 of 1% sucrose solution
5 cm3 of 1% protein solution (e.g. albumin)
a few drops of vegetable oil, e.g. olive oil or cooking oil
2% iodine in potassium iodide solution
Benedict’s solution
0.1 mol dm–3 sodium hydroxide solution
0.1 mol dm–3 hydrochloric acid
biuret reagent (or separate biuret solutions)
ethanol (or methylated spirits)
eye protection
Introduction
In this practical, you will:
• carry out tests for starch, reducing sugar, non-reducing sugar, protein and lipid.
Procedure
1
Carry out the tests for biological molecules shown in the table overleaf.
2
With the sucrose solution, try the reducing sugar test on the solution first, followed by the test for
non-reducing sugar.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Tests for biological molecules
Molecule
Test
Result if test is positive (and explanation)
Starch
Using a pipette, place a drop of starch
solution in a depression in a spotting
tile. Add a drop of iodine solution.
A blue–black colour is formed.
Place about 10 cm3 of glucose solution
in a test tube. Add a few drops of
Benedict’s solution. Stand the tube in
the water bath at 100 °C.
A brick-red precipitate is formed.
Reducing
sugar
(glucose)
A coloured polyiodide complex is formed
with starch.
The reducing sugar reduces the copper(II)
ions in the Benedict’s to copper(I) oxide.
(If a lower concentration of reducing sugar
is used, the colour may be green, yellow or
orange.)
Nonreducing
sugar
(sucrose)
Place about 10 cm3 of sucrose solution
in a test tube. Add three drops of dilute
hydrochloric acid.
Shake the tube and place it in the water
bath at 100 °C for 5 minutes. Remove
the tube and allow it to cool. Add three
drops of dilute sodium hydroxide
solution and mix, to neutralise the acid.
A brick-red precipitate is formed.
The acid hydrolyses the sucrose into
glucose and fructose, which both give a
positive Benedict’s test.
Repeat the reducing sugar test as above.
Protein
Lipid
Place about 5 cm3 of protein solution in
a test tube. Add an equal volume of
biuret reagent.
A lilac (mauve) solution is formed.
Place one drop of vegetable oil in a
clean, dry test tube. Add about 5 cm3 of
ethanol and shake thoroughly to
dissolve the oil. Pour the mixture into a
test tube three-quarters filled with cold
water.
A white emulsion is formed on the surface
of the water.
Nitrogen atoms in the peptide bonds of the
protein form a lilac complex with copper(II)
ions in the biuret reagent.
The alcohol mixes with the water, leaving
the lipid to form an emulsion of
microscopic droplets suspended at the
surface.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 2.2
Identifying three biological molecules
Safety
Wear eye protection.
Take care when using heating apparatus.
Dilute sodium hydroxide solution is an irritant. If it contacts the skin, wash off immediately with
plenty of water.
Apparatus and materials
•
•
•
•
•
•
•
•
eight test tubes
test-tube rack
labels or marker pen
thermostatically controlled water
bath maintained at 100 °C
thermometer
test-tube holder
three dropping pipettes
spotting tile
•
•
•
•
•
•
•
•
three tubes, labelled A, B and C, each containing 30 cm3
of a different unknown solution
2% iodine in potassium iodide solution
Benedict’s solution
0.1 mol dm–3 sodium hydroxide
0.1 mol dm–3 hydrochloric acid
biuret solution (or separate biuret solutions)
eye protection
pH paper
Introduction
In this practical, you will:
• identify which of three solutions A, B and C is an enzyme
• identify the other two solutions of different carbohydrates.
You are provided with three solutions labelled A, B and C. One of the solutions is an enzyme.
The other two are solutions of different carbohydrates, one of which is the substrate for the enzyme.
Procedure
1
Identify each solution using the reagents provided.
2
Present the results that you obtain in a suitable format.
3
When you have found out which solution is the enzyme, investigate the effect of the enzyme on
each of the two carbohydrate solutions.
4
State your conclusions and how you arrived at them.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 2.3
Semi-quantitative and quantitative tests for reducing sugars
Safety
Wear eye protection.
Take care when using heating apparatus.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
15 boiling tubes
test-tube rack
10 cm3 syringes
5 cm3 syringes
1 cm3 syringes
distilled water
thermostatically controlled water
bath maintained at 75 °C
six small beakers
filter funnel
filter paper
•
•
•
•
•
•
•
•
•
•
•
coloured pencils
stopwatch
colorimeter and cuvettes
50 cm3 of 10% glucose solution
Benedict’s solution
20 cm3 of lemon juice
20 cm3 of unknown glucose solution, labelled A
20 cm3 of unknown glucose solution, labelled B
eye protection
pipettes
labels or marker pen
Introduction
In this practical, you will:
• make a serial dilution of glucose
• test the different concentrations of glucose with Benedict’s solution
• make a colour chart
• use your colour chart to estimate the concentration of reducing sugar in some unknown solutions
• use a colorimeter to increase the sensitivity of the reducing sugar test.
Procedure
It is important to avoid contamination of solutions. Use a clean syringe for measuring out volumes of
different solutions.
A Making a serial dilution of glucose
1
Label five boiling tubes 1 to 5. Using a 10 cm3 syringe, place 10.0 cm3 of 10% glucose solution in
tube 1.
2
Using a 1 cm3 syringe, take 1.0 cm3 of the solution from tube 1 and transfer it to tube 2. Using a
10 cm3 syringe, add 9.0 cm3 of distilled water to tube 2 and mix the contents. The 1.0 cm3 of 10%
glucose solution has now been diluted ten times to make a 1% solution.
3
Repeat step 2, diluting the 1% solution in tube 2, to produce a 0.1% solution in tube 3. Repeat the
process with tubes 4 and 5. Tubes 1 to 5 now contain a serial dilution of the original glucose
solution, with the following concentrations: 10%, 1%, 0.1%, 0.01% and 0.001%.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
4
Tubes 1 to 4 have only 9.0 cm3 of solution left in them, but tube 5 has 10.0 cm3. Remove 1.0 cm3
of solution from tube 5 so that, for the Benedict’s test, all tubes start with the same volume of
solution.
5
Using a syringe, add 5.0 cm3 of Benedict’s solution to each tube, and place the tubes in a water
bath at 75 °C for 9 minutes.
6
Remove the tubes from the water bath and return them to the test-tube rack. Use coloured pencils
to make a chart of the colours.
B
Estimating the concentration of reducing sugar in some unknown solutions
1
Into three separate boiling tubes place 9.0 cm3 of either unknown solution A, unknown solution B
or the lemon juice. Label the tubes.
2
Add 5.0 cm3 of Benedict’s solution to each of the three tubes and heat in the water bath at 75 °C
for 9 minutes as in part A.
3
Compare the colours of the three tubes with those obtained from part A and estimate the
concentrations of reducing sugar present.
C
Extension: using a colorimeter to increase the sensitivity of the Benedict’s test
1
Make up a series of dilutions of the 10% glucose solution, of concentrations 0%, 0.5%, 1.0%,
1.5%, 2.0% and 2.5%, using distilled water. It is best to construct a table first to show how you
will make these dilutions. Have the table checked before you carry on.
2
Transfer 0.5 cm3 of each of your solutions to a labelled boiling tube, and add 5.0 cm3 of Benedict’s
solution to each tube. Place all the tubes in the water bath at 75 °C for 5 minutes.
3
Remove the tubes from the water bath and filter the contents of each tube into a clean, labelled
test tube. Using a pipette, transfer some of each filtrate to labelled colorimeter cuvettes.
4
Using an orange filter in the colorimeter, place the cuvette containing the filtrate from the
2.5% solution into the colorimeter. Set the colorimeter to zero absorbance using this solution.
Now read the absorbance of the other filtrates.
5
Process your results so that you could use the information to determine the concentration of
glucose in an unknown solution.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 3 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Enzymes in powdered form (i.e. if the teacher or technician makes up the solution) are harmful. They
are irritating to the eyes, there is a risk of serious damage to the eyes, and they may cause sensitisation
by inhalation.
Enzyme solutions equal to or stronger than 1% (w/v) are irritants. Enzyme solutions less than 1%
(w/v) are low hazards.
The risk assessment should consider the problem of allergic responses to enzymes.
Eye protection should be worn at all times.
Practical 3.1
amylase
Following the course of an enzyme-catalysed reaction using
It is recommended that a ready-made solution of 2% iodine in potassium iodide is purchased. The
ready-made solution is low hazard.
If time is short, you can carry out part A before the lesson and give students the results to enable them
to plot a standard curve.
Practical 3.2
Investigating the effect of temperature on the activity of trypsin
The denaturation of trypsin at high temperatures can be investigated further by exposing trypsin
solution to high temperatures (60 °C or 70 °C work well) for different periods of time. A suitable range
of times is 1, 2, 3, 5, 10 and 15 minutes. If the trypsin is cooled to room temperature after this heat
treatment, its remaining activity can be found using the method described.
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1
This investigation can also be modified to find the effect of dilute copper(II) sulfate solution on the
activity of trypsin. Copper(II) ions act as a non-competitive inhibitor of many enzymes, including
trypsin. The students can be provided with a serial dilution of copper sulfate, in the range 0.1 mol dm−3
to 0.1 mmol dm−3. Addition of 1.0 cm3 of 0.1 mol dm–3 copper sulfate will completely denature the
trypsin, resulting in zero activity. Lower concentrations will reduce the activity compared with a
control treated with 1.0 cm3 of distilled water. All solutions should be equilibrated to 40 °C, before
mixing and maintaining at this temperature. The turbidity of the solutions can be compared with the
end-point colour standard, which should be made up with 5.0 cm3 of milk suspension plus 5.0 cm3 of
dilute hydrochloric acid and 1.0 cm3 of the corresponding dilution of copper sulfate.
If the teacher makes up the copper(II) sulfate solution, note that the solid is harmful if swallowed,
irritating to eyes and skin, and very toxic to aquatic organisms (dilute with large volumes of water for
disposal).
Practical 3.3
of catalase
Investigating the effect of substrate concentration on the activity
The 2.5 mol dm–3 hydrogen peroxide solution should be labelled ‘harmful’. It is harmful if swallowed,
and can cause serious damage to eyes.
Practical 3.4A Investigating the effect of enzyme concentration on the activity
of rennin
Practical 3.4B Developing a procedure to investigate the effect of enzyme
concentration on the activity of rennin
Chapter P1 in the Coursebook features this investigation. Teachers might like to use this practical to
introduce their students to many of the key practical skills for the course.
Practical 3.4A includes a full set of instructions for the investigation.
Practical 3.4B is a second version of the same investigation, which gives students an opportunity to
make their own decisions about some aspects of the procedure and presentation of results. If students
read the relevant parts of Chapter P1 before doing this version, they should be able to make these
decisions. It would also be a good idea if they have the Coursebook available as they carry out the
practical.
Rennin can be purchased cheaply from supermarkets, pharmacies or health food shops in the form of
rennet – a liquid derived from the stomachs of young calves. Rennet contains several proteolytic
enzymes, including rennin. An alternative is to use powdered rennin, which should be supplied as a
1% solution.
If not available locally, liquid rennet and powdered rennin are both available from scientific supply
companies; rennet is also available from companies that supply domestic cheese-makers.
In the past, the only source of rennin was the stomachs of calves; now non-animal (vegetarian) rennet
is produced from genetically modified microorganisms such as yeast. Liquid and solid forms are
available. If a solid form is used, start with a 1% stock solution. Concentrations of rennin in rennet
vary considerably, so it is advisable to trial the method using different dilutions before using this
practical with students. Fresh, pasteurised full fat (3%) milk is the best substrate to use but the
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practical will also work with powdered milk and other forms of milk. Again, trialling is important to
make sure the students can achieve a suitable end point.
Practical 3.5
The hydrolysis of a protein by pepsin
Dilute hydrochloric acid should be labelled ‘irritant’.
Practical 3.6
Investigating the effect of pH on the activity of amylase
Iodine solution (iodine in potassium iodide) at a concentration of less than 0.1 mol dm–3 is low hazard,
so it is recommended that a ready-made solution stock solution of this concentration is purchased.
Make a 10-fold dilution of the stock solution.
Make a fresh 1% starch solution by mixing 5 g of soluble starch with cold water and dissolving the
paste in 500 cm3 of boiling water. Continue boiling until the solution is clear, and allow to cool.
Make a fresh solution of 1% amylase. The enzyme loses activity with storage.
The optimum pH for amylase activity is about pH6. Using the suggested volumes and concentrations
of reactants, complete digestion of the starch at pH6 should take about a minute. Check that this is the
case before the practical, and if the reaction is too slow or too fast, change the concentration of
enzyme to achieve a reasonable time for completion of the reaction.
Practical 3.7
catalase
Following the course of an enzyme-catalysed reaction, using
The 2.5 mol dm–3 hydrogen peroxide solution should be labelled ‘harmful’. It is harmful if swallowed,
and can cause serious damage to eyes.
The concentrations of yeast and hydrogen peroxide may need to be adjusted to achieve suitable results.
The technician or teacher can carry out some preliminary experiments before the lesson for this
purpose.
Practical 3.8
sucrase
Following the course of an enzyme-catalysed reaction, using
If the teacher is making up the solution of potassium permanganate, the solid permanganate is
oxidising and can cause fires if in contact with combustible material. It is harmful if swallowed, and
very toxic to aquatic organisms (dilute with large volumes of water for disposal).
Dilute sulfuric acid should be labelled ‘irritant’.
If time is short, you could provide students with the dilutions of glucose, or they could prepare them in
one lesson and monitor the sucrase reaction in a subsequent lesson. The practical could be modified to
find the effect of a different concentration of sucrase on the course of the reaction.
Pure sucrase must be used. Invertase is an impure form of sucrase contaminated with reducing sugar.
Similarly, pure (Analar®) sucrose must be used, rather than granulated sugar.
In part B, enzyme activity ceases after it has been added to the assay mixture, where it is denatured by
the acid.
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Practical 3.9 Investigating the effect of the concentration of an inhibitor on
the activity of catalase
Copper sulfate solution is harmful and irritant. Hydrogen peroxide is harmful and irritant, and may
bleach clothing or skin and cause burns. Spillages should be washed off immediately using plenty of
water. Yeast suspension is low hazard.
Make up the solutions as follows, scaling up as necessary:
At least 1 hour before the class practical, make a fresh 1% yeast suspension by adding 2 g of dried
(baker’s) yeast to 100 cm3 of warm distilled water in a beaker and making up to 200 cm3 with warm
distilled water. Sir well.
Prepare 3% hydrogen peroxide solution by placing 250 cm3 of stock 6% (20 vol) hydrogen peroxide in
a beaker and making up to 500 cm3 with distilled water. The solution should be covered to prevent
evaporation.
Prepare 3% copper sulfate solution by dissolving 6.0 g of hydrated copper sulfate (CuSO4.5H2O) in
100 cm3 of distilled water in a beaker and making up to 200 cm3 with distilled water.
The volumes and concentrations of solutions suggested in the procedure should produce a suitable
volume of gas from the control tube, but the teacher or technician should carry out the practical to
check that this is the case, and adjust the volumes or concentrations if necessary.
If gas syringes are not available, the oxygen can be collected over water in an inverted measuring
cylinder.
An alternative source of catalase is potato tissue. Take a large potato and peel it. Chop the peeled
potato into small pieces and macerate it in a blender with an approximately equal volume of distilled
water. Pour the macerate into a beaker and allow it to settle for several hours. A clear solution
containing catalase will form above the potato tissue and can be decanted. (Alternatively the extract
can be filtered into an Erlenmeyer flask, using a filter pump).
Practical 3.10
in solution
Comparing the activities of immobilised invertase and invertase
The buffer solution, sucrose and invertase solutions are slightly acidic and corrosive. Invertase powder
is harmful by inhalation and skin contact. Plastic gloves and a dust mask should be worn when
handling the powdered enzyme.
Including the controls, there are six reaction mixtures to be tested in this practical:
1 sucrose solution + immobilised invertase
2 sucrose solution + invertase in solution
3 sucrose solution (control)
4 buffer + invertase in solution (control)
5 buffer + immobilised invertase (control)
6 sucrose solution + effluent from immobilised enzyme.
This is too much for all students to complete in a limited time, so the work will need to be divided up
between groups. All should do 1 and 2, and the controls and effluent (3–6) should be shared out
between them.
It is important to use a water bath. The optimum temperature of invertase is about 55 °C. the reactions
should proceed at a suitable rate at 35 °C, but will be too slow at room temperature.
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Enzyme samples from different sources can vary, so it is suggested to try out a sample of invertase
before the full practical, in case a higher or lower concentration than that suggested in these
instructions is required.
Commercially available bottles of buffer tablets are available, for various pHs, which are simply
dissolved in water to form the desired buffer solution. These ready-made buffer solutions may be
preferred to making up buffer using sodium ethanoate and ethanoic acid.
Make up the solutions as follows, scaling up as necessary:
• Prepare 1 dm3 of 3% sodium alginate suspension by stirring 30 g of sodium alginate into 1 dm3 of
distilled water. Wear a disposable dust mask and gloves when weighing out the alginate. Prepare
the solution immediately before use.
• Prepare 3 dm3 of sodium ethanoate buffer (pH 4.7) by mixing 1300 cm3 of 0.2 mol dm–3 ethanoic
(acetic) acid with 1700 cm3 of 0.2 mol dm–3 sodium ethanoate (acetate).
• Prepare 1% invertase solution by dissolving 5.0 g of powdered invertase in 400 cm3 of the above
buffer. Make up to 1 dm3 with buffer.
• Weighed portions (0.125 g) of solid (powdered) invertase should be supplied in weighing boats in
a draught-free area such as a fume cupboard.
• Prepare 15% sucrose solution in buffer immediately before the practical (the sucrose slowly
hydrolyses at the pH of the buffer). Dissolve 150 g in 800 cm3 of buffer and make up to 1 dm3 with
buffer.
• Prepare 3% calcium chloride solution by dissolving 30 g of calcium chloride in 800 cm3 of
distilled water and making up to 1 dm3 with distilled water.
All solutions must be made up using distilled water. Calcium ions in tap water will cause the sodium
alginate to set.
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Practical 3.1
Following the course of an enzyme-catalysed reaction using
amylase
Safety
Wear eye protection.
1% amylase solution is an irritant and some people may be allergic to the enzyme. If it contacts the
skin, wash off immediately with plenty of water.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
colorimeter and cuvettes
water bath set at 25 °C
250 cm3 beaker
25 cm3 beaker
two plastic beakers
100 cm3 measuring cylinder
20 cm3 measuring cylinder
1 cm3 syringe
two 5 cm3 syringes
10 cm3 syringe
•
•
•
•
•
•
•
•
•
boiling tube
test-tube rack
stopwatch
distilled water
15 cm3 of 2% iodine in potassium iodide solution
60 cm3 of stock starch suspension (10.0 g dm–3) at 25 °C
10 cm3 of stock amylase solution (10.0 g dm–3) at 25 °C
eye protection
stirring rod
Introduction
In this practical, you will:
• construct a standard curve of absorbance against concentration of starch
• measure the course of the hydrolysis of starch by the enzyme amylase.
Iodine gives a blue–black colour with starch. The intensity of the colour is proportional to the
concentration of the starch. This intensity can be measured in a colorimeter, and used to follow the
course of the reaction. As the amylase hydrolyses the starch, the blue colour that starch forms with the
iodine decreases in intensity.
Procedure
A
Constructing a standard curve
1
Using the measuring cylinders, dilute some standard iodine solution 1 : 20 with distilled water, in
the large beaker. Make up 200 cm3 of this diluted iodine. Stir to mix thoroughly.
2
Set up the colorimeter with a red filter.
3
Place some of the diluted iodine solution in a cuvette, and place the cuvette in the colorimeter.
Set the absorbance reading of the colorimeter to zero. This is the blank solution (i.e. one
containing no starch). This blank must be kept for comparison throughout the rest of the
experiment.
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4
The stock starch suspension has a concentration of 10.0 g dm–3. Prepare ten dilutions of this stock
solution as shown in the table below.
Diluting stock starch solution
volume of stock
starch suspension /
cm3
volume of distilled
water / cm3
concentration of
diluted starch
suspension / g dm–3
Stock
1
2
3
4
5
6
7
8
9
10
10.0
7.0
6.0
5.0
4.0
3.0
1.0
0.6
0.4
0.1
0.05
0.0
3.0
4.0
5.0
6.0
7.0
9.0
9.4
9.6
9.9
9.95
10.0
7.0
6.0
5.0
4.0
3.0
1.0
0.6
0.4
0.1
0.05
5
Place 5.0 cm3 of the diluted iodine solution into a 25 cm3 beaker. Use a 1 cm3 syringe to add
0.5 cm3 of the 10.0 g dm–3 starch suspension to the iodine. Stir thoroughly. Pour this mixture into a
second cuvette and obtain an absorbance reading in the colorimeter.
6
Repeat step 5 for each of your prepared starch suspensions. Remember to wash out the syringe and
beaker after you have tested each mixture.
7
Draw a graph of absorbance against concentration of starch. This is a conversion graph or standard
curve.
B
Following the course of an enzyme-catalysed reaction
1
Using a 5 cm3 syringe, place 5.0 cm3 of the diluted iodine solution into a 25 cm3 beaker.
2
Place 15.0 cm3 of starch suspension and 5.0 cm3 of amylase solution from the stock in the water
bath into separate plastic beakers, using the 10 cm3 syringe and a clean 5 cm3 syringe respectively.
Mix these together and stir thoroughly. Now add 0.5 cm3 of the mixture to the beaker of iodine
solution, using the 1 cm3 syringe. Mix, and start the stopwatch. Pour some of the mixture into a
cuvette, and take a colorimeter reading. This must be done quickly. Pour the remaining starch–
amylase mixture into a boiling tube and place the mixture in the water bath.
3
Wash the syringes and beaker with distilled water and place another 5.0 cm3 of iodine solution into
the beaker. Take another 0.5 cm3 sample from the boiling tube 2 minutes after the enzyme and
substrate were first mixed, and repeat the procedure, sampling from the mixture every 2 minutes
for 30 minutes (or as long as it takes to get a reasonable change in the colorimeter reading).
4
Record the results in a suitable table. Use the standard curve to convert meter readings into
concentrations of starch, and record these values in the table.
5
Plot a graph of starch concentration against time.
6
Describe the shape of the curve, and suggest an explanation to account for its shape.
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Practical 3.2
Investigating the effect of temperature on the activity of trypsin
Safety
Wear eye protection.
Take care when using heating apparatus.
Apparatus and materials
•
•
•
•
•
•
•
•
•
12 boiling tubes
labels or marker pen
test-tube rack
four 5 cm3 syringes
large beaker (water bath)
thermometer
tripod
gauze
Bunsen burner
•
•
•
•
•
•
•
•
•
heat-proof mat
test-tube holder
stopwatch
stirring rod
50 cm3 of 4% suspension of powdered milk
30 cm3 of 0.5% trypsin solution
10 cm3 of 0.1 mol dm–3 hydrochloric acid
10 cm3 of distilled water
eye protection
Introduction
In this practical, you will:
• investigate the activity of trypsin.
Powdered milk contains the protein casein. A suspension of this milk in water is opaque and white but
it becomes translucent after hydrolysis by the enzyme trypsin. Casein is also hydrolysed by dilute
hydrochloric acid, which is a convenient way of preparing an end-point colour standard.
Procedure
1
Label four syringes ‘milk’, ‘water’, ‘acid’ and ‘trypsin’. Use these syringes to dispense the correct
solutions throughout the investigation, in order to avoid contamination.
2
Label two boiling tubes A and B. Stir the powdered milk suspension thoroughly, and use the
syringe labelled ‘milk’ to place 5.0 cm3 of the suspension in each tube. Using the correct syringes,
add 5.0 cm3 of distilled water to tube A and 5.0 cm3 of 0.1 mol dm–3 hydrochloric acid to tube B.
These tubes will be kept and used to compare the start and end points of the reaction during the
subsequent investigation.
3
Place 5.0 cm3 of the milk suspension in another boiling tube labelled C, and 5.0 cm3 of the trypsin
solution in a fourth tube labelled D.
4
Prepare a water bath at room temperature.
5
Place tubes C and D in the water bath. Allow 5 minutes for the tubes to equilibrate to the
temperature of the water bath. Now add the enzyme to the milk suspension, mix, and return this
tube to the water bath. Start the stopwatch.
6
Observe the tube closely and note the time when the tube has reached the colour of the end-point
tube B.
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7
Repeat steps 3–6 at four more temperatures between 20 and 70 °C, each time using the water bath
and heating apparatus to maintain the temperature of the reaction as constant as possible.
8
Convert your recorded times into rates. Rate is proportional to
of
9
1
. Multiplying the values
time
1
by 1000 gives rates in more manageable figures (the units are arbitrary).
time
Present your results in a suitable format.
10 Address the following points and questions.
a
From your results, what general conclusion can you make about the effects of temperature on
enzyme activity?
b What further work might you do to confirm your conclusions?
c
Discuss how changes in temperature affect enzyme activity.
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2
Practical 3.3
Investigating the effect of substrate concentration on the activity
of catalase
Safety
Wear eye protection.
The normal safety precautions associated with the use of chemicals apply.
Hydrogen peroxide solution can bleach clothing or skin and cause burns. Spillages should be washed
off immediately using plenty of water.
Apparatus and materials
•
•
•
•
•
•
four boiling tubes
bung to fit tubes, with delivery tube attached
distilled water
65 cm3 of 2.5 mol dm–3 hydrogen peroxide solution
cylinders of potato tuber tissue of uniform diameter,
about 25 cm long in total
two 10 cm3 syringes
•
•
•
•
•
•
•
100 cm3 gas syringe
retort stand
boss and clamp
scalpel
ruler
stopwatch
eye protection
Introduction
In this practical, you will:
• investigate the effect of the concentration of the substrate (hydrogen peroxide) on the rate of the
reaction, using catalase present in potato tuber tissue.
The enzyme catalase breaks down hydrogen peroxide into oxygen and water.
Procedure
1
Label four boiling tubes A to D. Using the distilled water and 2.5 mol dm–3 hydrogen peroxide
solution, make up four different mixtures as shown in the table below.
Four solutions of hydrogen peroxide
Tube
A
B
C
D
2
Volume of distilled
water / cm3
8.0
6.0
4.0
2.0
Volume of hydrogen
peroxide / cm3
2.0
4.0
6.0
8.0
Final hydrogen peroxide
concentration / mol dm–3
0.5
1.0
1.5
2.0
Attach the bung and delivery tube to boiling tube A and attach the end of the delivery tube to the
gas syringe, clamped horizontally.
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3
With a sharp scalpel and a ruler, cut ten slices 1–2 mm thick from the potato tuber tissue cylinder
supplied. Immerse all ten slices in the hydrogen peroxide solution in tube A. After 45 seconds
replace the bung and record the volume of gas produced in 1 minute.
4
Discard the contents of tube A, rinse the tube and repeat the procedure twice more, recording the
volume of gas produced on each occasion.
5
Now repeat steps 3–4 using tubes B, C and D. Record your results in a suitable table and plot as a
graph.
6
Address the following points and questions.
a
Write a balanced equation for the reaction that is taking place in tubes A to D.
b What was the effect of increasing the concentration of hydrogen peroxide on the rate of
reaction?
c
What were the chief sources of error in this investigation?
d If you were to increase the concentration of hydrogen peroxide further still, how would you
expect the rate of reaction to change?
e
Explain the relationship between substrate concentration and the rate of reaction.
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2
Practical 3.4A
Investigating the effect of enzyme concentration on the activity
of rennin
Safety
Wear eye protection.
Take care when using heating apparatus.
Avoid skin contact with the enzyme solution. Wash off any splashes.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
10 test tubes
test-tube rack
two 1 cm3 syringes
10 cm3 syringe
large beaker (water bath)
Bunsen burner
heat-proof mat
tripod
gauze
about 10 cm3 distilled water
•
•
•
•
•
•
•
one bung to fit the test tubes
stop watch
marker pen
thermometer
piece of black card about 10 cm × 10 cm
about 10 cm3 liquid rennet (or the same
volume of 1% rennin prepared from the
powdered form)
50 cm3 pasteurised milk
Introduction
In this practical, you will:
• investigate how changing the concentration of rennin affects its activity.
All young mammals feed on milk. Rennin is an enzyme that is found in the stomachs of young
mammals. Its substrate is a protein in milk called casein. The rennin breaks the casein down, causing
the milk to clot (Chapter P1, Figure P1.1, page 247 in the Coursebook).
For this experiment you will use rennet, which is an extract obtained from the stomach lining of young
calves. Rennet contains small amounts of rennin. The activity of the enzyme will be established by
measuring how quickly it causes milk to clot.
Procedure
Before starting, read through the whole procedure, particularly how to determine the end-point in
step 11 (you should try observing for an end-point using milk and undiluted enzyme before you begin
the experiment).
1
Before you start the practical work, prepare a table like the one on the next page for recording your
results.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Record of results for the effect of enzyme concentration on the activity of rennin
Tube
Concentration
of rennet / %
1
2
3
4
5
10
8
6
4
2
A
Time at
start / s
0
B
Time at endpoint / s
A–B
Time to reach
end-point / s
Rate of reaction
(1000/time in
seconds)
2
Place five test tubes in a test-tube rack.
3
Using a 10 cm3 syringe add 9 cm3 of pasteurised milk to each of the tubes.
4
Set up a water bath at 37 °C. Maintain this temperature to within ± 2 °C throughout the experiment.
5
Place the tubes of milk in the water bath.
6
Label five more test tubes 1 to 5 and place them in a test tube rack.
7
Using separate 1 cm3 syringes for the rennet and distilled water, prepare a series of dilutions of the
rennet solution provided as shown in the table below.
Dilutions of rennet
Tube
Volume of rennet solution / cm3
Volume of distilled water / cm3
1
1.0
0.0
2
0.8
0.2
3
0.6
0.4
4
0.4
0.6
5
0.2
0.8
8
Remove a tube of milk from the water bath and pour its contents into tube 1. Place a bung in the
tube and invert the tube to mix the contents. Place tube 1 back into the water bath at 37 °C.
9
Start the stop watch and leave it running throughout the experiment.
10 Repeat step 8 for tubes 2, 3, 4 and 5. For each tube, using your results table, note the time when
the milk is added. These are the start times and must be subtracted from the finish times at the end
of the experiment.
11 The milk will gradually ‘set’ (coagulate or clot). About every 30 to 60 seconds, check all five
tubes for the end-point.* Rock each tube gently from side to side when checking. Record the time
at each end-point in your results table. If no reaction is visible after 15 minutes, record the time
taken as infinity (∞) and the rate as zero.
*The end-point can be taken as the time taken for a ‘partial set’ (when the developing clot resists
movement or when small white granules begin to appear – these can be seen more clearly by
holding a piece of black card behind the tube and tilting the tube). It is easier to judge a ‘complete
set’. This is when the contents of the tube become solid, but it takes longer.
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2
12 Calculate the time taken to reach the end-point for each test tube and enter the results in your table.
13 Convert each time taken to a rate of reaction:
rate of reaction =
1000
t
where t = the time taken in seconds.
Enter your results in the table.
14 Decide which is the independent variable and which is the dependent variable in this experiment.
Plot a graph to show the effect of rennet concentration on the rate of reaction.
15 Address the following points and questions
a
State two controlled variables (control variables) in this experiment.
b Explain why the concentration of rennet in tube 1 is 10%.
c
Describe the results shown in the graph.
d Explain the effect of changing the concentration of enzyme (rennet) on the rate of reaction.
e
A sixth tube could be set up containing 1 cm3 water and 9 cm3 milk. What would be the
purpose of this tube?
f
State two limitations of the investigation.
g
Suggest one improvement to the method that will address each of the limitations that you have
given.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
3
Practical 3.4B
Developing a procedure to investigate the effect of enzyme
concentration on the activity of rennin
Safety
Wear eye protection.
Take care when using heating apparatus.
Avoid skin contact with the enzyme solution. Wash off any splashes.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
10 test tubes
test-tube rack
two 1 cm3 syringes
10 cm3 syringe
large beaker (water bath)
Bunsen burner
water tripod
Bunsen burner
heat-proof mat
one bung to fit test tubes
•
•
•
•
•
•
•
stop watch
marker pen
thermometer
piece of black card about 10 cm × 10 cm
about 10 cm3 liquid rennet (or the same
volume of 1% rennin prepared from the
powdered form)
50 cm3 pasteurised milk
about 10 cm3 distilled water
Introduction
In this practical, you will:
• develop a method to investigate how changing the concentration of rennin affects its activity.
All young mammals feed on milk. Rennin is an enzyme that is found in the stomachs of young
mammals. Its substrate is a protein in milk called casein. The rennin breaks the casein down, causing
the milk to clot (Chapter P1, Figure P1.1 page 247 in the Coursebook).
For this experiment, you will use rennet, which is an extract obtained from the stomach lining of
young calves. Rennet contains small amounts of rennin. The activity of the enzyme will be established
by measuring how quickly it causes milk to clot.
You will need to decide the following:
• the temperature at which to carry out the experiment
• the number of dilutions of the rennet
• the intervals between dilutions (in other words, what dilutions will you use?)
• the procedure for diluting the rennet
• the end-point used
• how to present your results.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Procedure
Before starting, read through the whole procedure, particularly how to determine the end-point in
step 11 (you should try observing for an end-point using milk and undiluted enzyme before you begin
the experiment). If you are unsure about any of your decisions, ask your teacher for advice.
1
Decide how many dilutions of the rennet you will use. You will need one tube of milk for each
dilution. Place the chosen number of tubes in a test tube rack. Using a 10 cm3 syringe, add 9 cm3
of pasteurised milk to each tube.
2
Set up a water bath at the chosen temperature. Maintain this temperature to within ± 2 °C
throughout the experiment.
3
Place the tubes of milk in the water bath.
4
Place one empty test tube in a test tube rack for every tube of milk in the water bath (for example,
if there are three tubes of milk, place three empty test tubes in a rack). Label the tubes 1, 2, 3, etc.
These will be the tubes in which the rennet will be placed in step 6 below.
5
You will now need to prepare your dilution series for the rennet. Decide what dilutions you will
prepare. You will need 1 cm3 of rennet solution in each numbered tube. Make a table to show the
volume of rennet and the volume of distilled water needed to prepare 1 cm3 of each dilution. Ask
your teacher to check your table before you proceed.
6
Using separate 1 cm3 syringes for the rennet and distilled water, prepare your chosen series of
dilutions. Use one numbered test tube for each dilution.
7
Make a table to record your results. You will need to record the tube number, the concentration of
rennet and the relevant times from the stop clock (read steps 9 and 10 below). Also add a column
for the rate of reaction, which you will calculate after the experiment.
8
Remove a tube of milk from the water bath and pour its contents into tube 1. Place a bung or
stopper in the tube and invert the tube to mix the contents. Replace tube 1 in the water bath.
9
Start the stop watch and leave it running throughout the experiment.
10 Repeat step 8 for tubes 2, 3, etc. For each tube, note the time when the milk is added in your table.
These are the start times and must be subtracted from the finish times at the end of the experiment.
11 The milk will gradually ‘set’ (coagulate or clot). Small white granules begin to appear – these can
be seen more clearly by holding a piece of black card behind the tube and tilting the tube. About
every 30 to 60 seconds, check all your tubes for the end-point (decide this for yourself). Rock each
tube gently from side to side when checking. Record the time at each end-point in your table. If no
reaction is visible after 15 minutes, record the time taken as infinity (∞) and the rate as zero.
12 Calculate the time taken to reach the end-point for each test tube and enter the results in your table.
13 Convert each time taken to a rate of reaction:
rate of reaction =
1000
t
where t = the time taken in seconds.
Enter your results in the table.
14 Decide which is the independent variable and which is the dependent variable in this experiment.
Plot a graph to show the effect of rennet concentration on the rate of reaction.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
15 Address the following points and questions.
a
State the temperature at which you carried out the experiment and explain your choice.
b State two controlled variables (control variables) in this experiment.
c
Describe the results as shown in the graph.
d Explain the effect of changing the concentration of enzyme (rennet) on the rate of reaction.
e
State two limitations of the investigation.
f
Suggest one improvement to the method that will address each of the limitations that you have
given.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
3
Chapter 4 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Eye protection should be worn at all times.
Practical 4.1
Investigating the properties of cell membranes
Ethanol and its vapour are highly flammable. Keep them well away from the Bunsen burner and other
sources of ignition. Ethanol should be labelled ‘flammable’ and, if methylated spirits are used,
‘harmful’. Dilute hydrochloric acid should be labelled ‘irritant’.
Discs of beetroot tissue should be washed thoroughly before the investigation to remove traces of
pigment from the outside of the discs. This takes at least 15 minutes – if students stop rinsing the discs
too early, the tissue in the control experiment will continue to ‘bleed’.
A large cork borer can be used to cut discs of the same diameter. Fresh beetroot must be used (not
beetroot preserved in vinegar).
The effect of temperature on the integrity of the membrane could be investigated by placing the
beetroot into tubes maintained at different temperatures for the same length of time, perhaps
10 minutes. Temperatures between 20 C and 100 C, at 10 C intervals, could be tried.
Practical 4.2
Investigating plasmolysis
White onion can be used instead of red onion, although the pigment in the red onion cells makes it
easier to see plasmolysis taking place. Another alternative is to use the epidermis of rhubarb petioles.
If plasmolysis is too rapid in 1 mol dm–3 sucrose solution, try 0.5 mol dm–3 instead.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 4.3
Finding the mean water potential of potato tuber cells
Use a sharp knife to cut flat sheets of potato, 10 mm thick. Then cut the sheets into 10 mm strips,
removing all skin. Thirty minutes is usually long enough to allow a great enough percentage change in
mass of the potato tissue, although it would be better to leave the tissue until no further change in mass
occurs. If short lesson times are a problem, the dilutions of sucrose solution could be made up in the
previous lesson and stored in the fridge until required. To extend the time the tissue sections are in the
solutions, different teaching sets could collaborate over the weighing of the sections.
Cutting the chips into smaller slices increases the surface area for osmotic water movement, but
introduces further errors in consistency of blotting, as well as differences in the total surface area
exposed in each tube. It is better to control these factors by using larger chips.
The method takes too much time for students to carry out repeats at each concentration of sucrose. To
evaluate reliability, they can pool their results for the % change in mass at each concentration, and
calculate mean values.
Practical 4.4 Estimating the solute potential of cell sap by the incipient
plasmolysis method
White onion can be used instead of red onion, although the pigment in the red onion cells makes it
easier to see plasmolysis taking place. Another alternative is to use the epidermis of rhubarb petioles.
If there is time, for each glucose solution, students can find the % of plasmolysed cells from different
areas of the tissue and average their results.
Practical 4.5
Investigating the properties of Visking tubing
All solutions in the first part are low hazard.
The second part of the practical (steps 13–20) is included as extension material, which can be carried
out if time is available. This is Chardakoff’s method (the ‘hanging drop’ method). It shows that there
has been a change in the concentrations of the solutions inside the Visking tubing bags that were
placed in tubes A and C.
Methylene blue will stain the skin. If any stain contacts the skin it must be washed off under the tap.
Practical 4.6
Rates of diffusion in ‘cells’ of different sizes
The teacher or technician will need to experiment with the concentration of potassium permanganate
(potassium manganate(VII)) in order to produce a suitably dyed agar jelly that changes colour in a
reasonable time.
A different concentration of acid can also be tried.
Practical 4.7
membranes
Investigating the effect of ethanol on the permeability of cell
Ethanol is flammable and should be kept away from naked flames.
Methylene blue will stain the skin; students must use forceps for handling the plant tissue. If any stain
contacts the skin it must be washed off under the tap.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Solutions and reagents can be supplied in beakers or other suitable containers that will allow for the
removal of solution using a syringe.
To prepare 32% ethanol, take 32 cm3 of ethanol and make up to 100 cm3 with distilled water (scale up
volumes to make enough for the class). The dilute ethanol should be supplied in a covered container to
prevent evaporation.
Suitable plant tissues are potato tuber, sweet potato or cassava. The tissue should be prepared 24 hours
before the practical. Use fresh material that has not be stored or refrigerated for a long period of time.
Remove the outer skin and cut into lengths with a cross-section of 0.5 cm  0.5 cm.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 4.1
Investigating the properties of cell membranes
Safety
Wear eye protection.
Take care when using heating apparatus.
Dilute hydrochloric acid is an irritant. Spillages should be washed off immediately using plenty of
water.
Ethanol is highly flammable – keep it well away from the Bunsen flame.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
12 boiling tubes
labels or marker pen
test-tube rack
large beaker (water bath)
tripod
gauze
Bunsen burner
heat-proof mat
test-tube holder
colorimeter
•
•
•
•
•
•
•
thermometer
50 cm3 of distilled water
15 cm3 of ethanol
15 cm3 of dilute hydrochloric acid
three 10 cm3 syringes
20 discs of fresh beetroot (Beta vulgaris) about 2 mm thick,
of a suitable size to fit the boiling tubes (discs should be
rinsed under running water until no further colour is lost –
this takes at least 15 minutes)
eye protection
Introduction
In this practical, you will:
• investigate the effects of various solutions and different temperatures on plant cell membranes.
Beetroot cells contain a red pigment in their cell sap. The pigment is normally retained in the vacuole
by the membrane around the vacuole (the tonoplast).
Procedure
1
Label six boiling tubes 1 to 6. Place three discs of the beetroot in each tube.
2
Add 10 cm3 of distilled water to tube 1, 10 cm3 of ethanol to tube 2 and 10 cm3 of dilute
hydrochloric acid to tube 3. Leave these three tubes in the rack for observation.
Add 10 cm3 of distilled water to tubes 4, 5 and 6. Half fill the beaker with tap water and place
tubes 4, 5 and 6 in the beaker. Heat the water and remove tube 4 when the temperature reaches
40 °C. Continue to heat the water and remove tube 5 when the temperature reaches 65 °C. Again,
continue to heat the water and remove tube 6 when the temperature reaches 100 °C.
Pour off the solution from tubes 1–6 into six clean boiling tubes also labelled 1–6.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
5
Examine each tube against a white background and compare their contents. Note the depth of
colour and record your observations in a table.
6
Use a colorimeter fitted with a green filter to measure the absorbance of the different solutions
from tubes 1–6 against a blank of distilled water. Add the absorbance readings to your table.
7
Explain the depth of colour produced in each of the tubes.
8
Explain how you might modify the second half of the experiment to investigate more fully the
effect of temperature on the integrity of the beetroot membranes. Consider how exposure to
different temperatures might be better controlled and suggest suitable temperature intervals.
If there is time, you could carry out this investigation.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 4.2
Investigating plasmolysis
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Take care when using heating apparatus.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
microscope
two microscope slides
two cover slips
dropping pipettes
scalpel
forceps
filter paper
•
•
•
•
•
•
watch glass
red onion (Allium sp.)
knife
chopping board or tile
20 cm3 of distilled water
20 cm3 of sucrose solution (1 mol dm–3)
Introduction
In this practical, you will:
• make a temporary preparation of epidermal cells from a red onion
• observe plasmolysis and deplasmolysis in these epidermal cells.
In plant cells, plasmolysis takes place when the cell surface membrane pulls away from the cell wall,
leaving a space between the membrane and the wall. Plasmolysis is caused by water passing out of the
cell. This may be due to osmosis, or it may happen if the cell loses water by evaporation. Cells can
recover from a plasmolysed state if they take up enough water. This recovery is called deplasmolysis.
You can observe plasmolysis and deplasmolysis in cells taken from the bulb of a red onion.
Procedure
1
Remove some small squares of epidermal tissue from the inner concave surface of one of the outer
storage ‘leaves’ of a red onion bulb (see the diagram below). The squares must be small enough to
fit under a cover slip. It is easiest to make some square cuts in the inner surface of the leaf before
you remove the epidermis. Place the pieces of tissue in some distilled water in a watch glass.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
2
Mount a piece of the epidermal tissue in water on a microscope slide. Keep the tissue flat, with no
air bubbles under the cover slip.
3
Observe these normal cells through the medium-power and high-power objectives of the
microscope.
4
Mount another piece of onion epidermis in 1 mol dm–3 sucrose solution on a clean microscope
slide. Again observe the onion cells under the medium-power and high-power objectives. Can you
see plasmolysis taking place? If you watch carefully over several minutes, you should be able to
see the cell surface membrane pull away from the inside of the cell wall.
5
Draw and label a few plasmolysed cells (Figure 4.14 on page 85 of the Coursebook).
6
Irrigate the slide with distilled water. Place a drop of water on the slide so that it touches the edge
of the cover slip. Use a piece of filter paper to draw the water across the slide under the cover slip,
by touching the edge of the cover slip on the side opposite the water (see the diagram below).
7
Observe the cells for any signs of deplasmolysis. If you do not see any change, remove the cover
slip and soak the tissue in distilled water in the watch glass for a few minutes. Replace the cover
slip and try again.
8
Address the following points and questions, using the term water potential where you can.
a
Have the cells plasmolysed in the sucrose solution? Have all the cells plasmolysed? If not, can
you explain why?
b Why do the cells deplasmolyse in distilled water?
c
What fills the space between the cell wall and the cell surface membrane in a plasmolysed
cell?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 4.3
Finding the mean water potential of potato tuber cells
Safety
Take care when using sharps.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
six boiling tubes
labels or marker pen
test-tube rack
sharp knife
chopping board
potato tuber tissue (Solanum tuberosum)
•
•
•
•
•
paper towels
two 20 cm3 syringes
balance weighing to 0.01 g
100 cm3 of 1 mol dm–3 sucrose solution
100 cm3 of distilled water
Introduction
In this practical, you will:
• establish the mean water potential of potato tuber cells.
If pieces of potato tuber tissue are placed in different concentrations of sucrose solution, the tissue
gains or loses mass depending on the water potential of the solution. If there is no change in the mass
of the tissue, then the sucrose solution has a water potential which is the same as that of the tissue
itself.
Procedure
1
Label six boiling tubes 1 to 6. Using distilled water and 1 mol dm–3 sucrose solution, make up six
different dilutions of sucrose solution as shown in the table below. Shake each mixture thoroughly.
Dilutions of sucrose in tubes 1–6
Tube
2
Volume of distilled
water / cm3
Volume of 1 mol dm–3 sucrose
solution / cm3
Molarity / mol dm–3
1
20
0
0.0
2
16
4
0.2
3
12
8
0.4
4
8
12
0.6
5
4
16
0.8
6
0
20
1.0
Using a sharp knife, cut six potato tuber sections at least 10 mm in width and 50 mm long.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
3
Blot each section gently to remove any surface moisture. Weigh the first section and place it in
tube 1, making a note of the mass. Repeat for tubes 2 to 6.
4
Leave the potato tuber sections in the sucrose solutions for at least 30 minutes.
5
Remove the tuber sections one at a time; gently blot the surface of the potato and re-weigh. Make
sure that you don’t mix up the sections.
6
Calculate the percentage change in mass of each section, using the formula:
percentage change in mass =
change in mass
´ 100%
starting mass
7
Plot a graph of the percentage change in mass against the molarity of the sucrose solution.
(The x-axis should intercept with the y-axis at 0% change in mass.)
8
Find the point where the line of your graph crosses the x-axis (i.e. where there is no change in
mass). This is the molarity of a solution that has a water potential equal to the water potential of
the potato tuber cells.
9
Use the table below to work out the water potential of a solution having this molarity. To do this, it
is best to plot a second graph, of water potential against molarity.
Water potential for a range of sucrose solutions
Molarity of sucrose solution / mol dm–3
Water potential / kPa
0.0
0
0.2
–540
0.4
–1120
0.6
–1800
0.8
–2580
1.0
–3500
10 Explain your results using the term water potential.
11 Describe the limitations of using a change in mass as a measure of osmosis.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 5 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Eye protection should be worn at all times.
Practical 5.1
Investigating mitosis in root tips
A stock solution of acetic orcein contains 2.2 g of orcein dissolved in 100 cm3 of glacial ethanoic
(acetic) acid, and is corrosive. Dilute 10 cm3 of this solution with 12 cm3 of water before use. Wear
eye protection and gloves. Carry out the preparation and dilution in a fume cupboard. The diluted
solution should be discarded after the practical.
Students should wear gloves when handling acetic orcein stain.
Dilute (1 mol dm–3) hydrochloric acid should be labelled ‘irritant’.
Root tips of other species such as onion or garlic (Allium sp.) can be used. However, broad bean (Vicia
faba) has a smaller number of larger chromosomes, which are easier to see through the microscope.
Soak the beans for 24 hours and then plant them in moist potting compost. This is better than using
jars with blotting paper, where the root tips tend to dry out. The beans take about 10 days (depending
on the temperature) to grow suitable roots. Keep them well watered. Two or three beans will usually
provide enough root tips for a class.
Activity 5.1
Investigating a karyotype
There are no specific safety issues.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 5.1
Investigating mitosis in root tips
Safety
Wear eye protection.
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Take care when using heating apparatus.
Do not inhale the acetic orcein fumes. Wear gloves when handling the stain.
Dilute (1 mol dm–3) hydrochloric acid is an irritant. Spillages should be washed off immediately using
plenty of water.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
•
scissors
scalpel
forceps
mounted needle
dropping pipettes
filter paper
microscope
slides
cover slips
Bunsen burner
prepared slide of root tip showing mitosis
•
•
•
•
•
•
•
•
•
•
acetic orcein stain
boiling tube
test-tube rack
aluminium foil
thermostatically controlled water bath set at 70 °C
10 cm3 of 1 mol dm–3 hydrochloric acid
calibrated eyepiece graticule
broad bean (Vicia faba) plants with roots about
5–10 cm in length
eye protection
protective gloves
Introduction
In this practical, you will:
• make a temporary preparation of root tips of broad bean (Vicia faba).
Part of a plant root tip contains meristematic tissue, where stages of mitosis can be seen. You should
make a few slide preparations so that you can use the best one for making observations and drawings.
Before you look at your own preparations through the microscope, you should look at a prepared slide
of a root tip and identify cells that are undergoing mitosis. A good time to do this is while the roots are
staining in the acetic orcein (step 5). Acetic orcein stains the chromosomes red.
Procedure
1
Place about 10 cm3 of acetic orcein stain in a boiling tube. Add three drops of 1 mol dm–3
hydrochloric acid. Cover the tube with a cap of aluminium foil. Acetic orcein has a strong vinegarlike smell which can be an irritant. The foil reduces the escape of fumes from the tube.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
2
Remove a bean plant from the compost in which it has been growing, taking care not to damage
the root tips. Gently wash the roots under the tap, collecting the compost in a sieve to avoid
blocking the sink.
3
Using scissors cut a few undamaged roots about 5 cm long from the plant. Make sure you note
which end is the growing tip of the root.
4
Remove the foil cap from the boiling tube and place the roots, tips down, in the acetic orcein. It
doesn’t matter if the whole root is not fully submerged in the stain as long as the tip is. Replace the
foil cap.
5
Place the boiling tube in a water bath set at 70 °C. After 30–40 minutes, remove the tube and stand
it in a test-tube rack. (Use the heating time to look at prepared slides of root tips.)
6
Wear protective gloves from now on, to avoid getting the stain on your hands. Use forceps to
transfer a stained root tip to a microscope slide. Use a scalpel to cut off and discard all but the
terminal 3–4 mm of the tip.
7
Add two drops of the acetic orcein from the boiling tube. Gently lower a cover slip onto the
specimen, using a mounted needle. Place a filter paper on top of the cover slip and firmly press
down to squash the tissue. The aim is to spread out the cells to a single layer, but without altering
their relative positions along the root tip. You can use the handle of a mounted needle to complete
the squashing. The tissue should spread out to cover an area about 5 mm × 5 mm – don’t be afraid
to squash the preparation well, or you won’t be able to see individual cells. Prepare two or three
root-tip samples in this way.
8
Observe each of your slides using the low-power objective lens. Cells undergoing division are
small and square in shape, with the nucleus taking up most of the cell during interphase (see
Figure 5.9 in the Coursebook). Find an area containing cells in different stages of mitosis. Select
the slide which shows the cells most clearly.
9
Look at the dividing cells using the high-power objective and identify cells in interphase,
prophase, metaphase, anaphase and telophase. Make labelled drawings of each of these stages. Use
a calibrated eyepiece graticule to measure a cell and add a scale to your drawing.
10 Estimate the percentage of cells in the meristematic area that are undergoing mitosis. This is
equivalent to the proportion of the cell cycle taken up by mitosis.
(Note that interphase is not part of mitosis.)
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 6 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Enzymes in powdered form (i.e. if the teacher or technician makes up the solution) are harmful. They
are irritating to the eyes, there is a risk of serious damage to the eyes, and they may cause sensitisation
by inhalation.
Enzyme solutions equal to or stronger than 1% (w/v) are irritants. Enzyme solutions less than 1%
(w/v) are low hazards.
The risk assessment should consider the problem of allergic responses to enzymes.
Eye protection should be worn at all times.
Practical 6.1
Extracting DNA from onion bulb cells
Ethanol and its vapour are highly flammable. Keep them well away from the Bunsen burner and other
sources of ignition. Ethanol should be labelled ‘flammable’.
Novo neutrase™ solution can be used instead of pepsin. It is available from the National Centre for
Biotechnology Education.
The threads of DNA can also be stained with acetic orcein. Other plant materials such as kiwi fruit
(Actinidia sp.) or banana can be used instead of onion bulb tissue.
Activity 6.1
DNA and RNA
There are no specific safety issues.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 6.1
Extracting DNA from onion bulb cells
Safety
Wear eye protection.
Take care when using sharps.
Take care when using heating apparatus.
Ethanol is highly flammable – keep it well away from the Bunsen flame.
Enzyme solutions equal to or stronger than 1% (w/v) are irritants.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
glass rod
blender
water bath set at 60 °C
two 400 cm3 beakers
boiling tube
test-tube rack
funnel and coarse filter paper (or coffee
filter paper)
Pasteur pipette
knife or scalpel
microscope slide
•
•
•
•
•
•
•
•
chopping board
onion bulb (Allium sp.)
universal indicator
600 cm3 beaker containing small amount of ice
(as an ice-water bath)
3 g of sodium chloride and 10 cm3 of washing-up
liquid dissolved in 100 cm3 of distilled water
10 cm3 of 95% ethanol stored in a freezer for at
least 12 hours before use
2 cm3 of 1% pepsin
eye protection
Introduction
In this practical, you will:
• extract some DNA from onion cells.
Procedure
1
Cut an onion bulb into small pieces and transfer them to a beaker. Add approximately 100 cm3 of
the salt and detergent mixture, and stir thoroughly.
2
Place the beaker in a water bath at 60 °C for 15 minutes.
The detergent forms complexes around the membrane phospholipids and proteins, causing them to
precipitate out of solution. The sodium ions from the salt shield the negatively charged phosphate
groups of the DNA molecules, causing them to coalesce. At 60 °C, nuclease enzymes, which
would otherwise start to fragment the DNA, are partially denatured.
3
Cool the mixture in an ice-water bath for 5 minutes, stirring frequently.
This slows down the denaturation of DNA that would occur if a high temperature were
maintained.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
4
Pour the mixture into a blender and blend for 5 seconds.
This permits the release of DNA by further degrading the cell walls and membranes. However, the
DNA will be broken up if blending is carried out for more than 5 seconds.
5
Filter the mixture into a beaker, ensuring that the foam on the surface of the mixture does not
contaminate the filtrate. Note that filtration may take some time to complete.
6
Transfer 6 cm3 of the filtrate to a clean boiling tube.
7
Add four drops of the 1% pepsin solution provided. Mix thoroughly.
Pepsin hydrolyses the proteins in the mixture to amino acids.
8
Slowly pour 9 cm3 of ice-cold 95% ethanol down the side of the tube so that it forms a layer over
the filtrate and enzyme mixture. Leave the tube for a few minutes without disturbing it.
9
Using a Pasteur pipette, try to draw up some threads of the fibrous material that forms in the cold
ethanol and transfer the material to a slide. These threads contain DNA from the onion cells. Add a
drop of universal indicator to confirm they are acidic.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 7 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Eye protection should be worn at all times.
Practical 7.1
Investigating stem, root and leaf structure
Phloroglucinol (benzene-1,3,5-triol) is an irritant.
The fresh pieces of stem should be from a plant with a translucent stem. Sections are very easy to cut
from a Busy Lizzie (Impatiens sp.). Celery is not suitable for this exercise, as celery stalks are petioles
rather than stems.
Practical 7.2
Investigating xylem and phloem
Solid eosin is an irritant, take care not to inhale the powder or get it on skin when making up the
solution.
Celery (Apiaceae graveolens) tissue can be torn lengthways to reveal vascular bundles. These can be
teased out and cut into lengths to be given to students. The tissue can be teased apart with mounted
needles and provided to students in watch glasses so that they can see individual cells.
Practical 7.3
potometer
Investigating the rate of transpiration of a leafy shoot using a
It is important to use a twig with a stem that is a good fit in the rubber tubing. When the apparatus and
twig are held in the clamp, water will be seen to drip out of the end of the capillary tubing if there are
any air leaks. Twigs can take some time to equilibrate.
A hair drier or fan can be used to find the effect of moving air on the rate of transpiration. A suitable
control would be the same shoot placed in still air, or in a clear plastic bag, and allowed to equilibrate.
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Other factors (including temperature, light intensity, humidity, and wind speed) must be maintained
the same in each treatment.
It is possible to compare rates of transpiration in two twigs of the same species, or two different
species such as a mesophyte and a plant showing xerophytic adaptations. In this case, the rate must be
measured as volume of water taken up per unit leaf area. A xerophyte would be expected to have a
lower rate of transpiration per unit leaf area, assuming that all other conditions are the same.
Practical 7.4
Preparing a slide of epidermal cells from a lettuce leaf
It is recommended that a ready-made solution of 2% iodine in potassium iodide is purchased. The
ready-made solution is low hazard.
Practical 7.5
Investigating stomatal density
This exercise can be carried out with other species of plant, such as Pelargonium, although privet is
easier, and best tried first. Other comparisons can be made, such as stomatal densities of plants of the
same species from sunny and shady habitats.
The nail varnish takes at least 20 minutes to dry fully.
Test the nail varnish on leaves before the lesson: some cheaper brands do not work well.
Practical 7.6
Investigating the leaves of xerophytes
A simple model of a marram grass leaf can be made with a piece of paper rolled lengthwise. This
helps students to relate the transverse section to the whole leaf.
Teachers may be able to find suitable local examples of xerophytes for the students to study.
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Practical 7.1
Investigating stem, root and leaf structure
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Phloroglucinol (benzene-1,3,5-triol) is an irritant. Spillages should be washed off immediately using
plenty of water.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
two pieces of a non-woody stem
(e.g. Impatiens sp.);
one of the pieces should have been standing in
water for 24–48 hours, the other should have
been standing in water with a little eosin added to
it for the same time
prepared slide of TS of a young stem
(e.g. sunflower (Helianthus sp.) or buttercup
(Ranunculus sp.))
prepared slide of TS of a young root
(e.g. buttercup)
prepared slide of TS of a leaf
from a mesophyte (e.g. privet (Ligustrum sp.))
•
•
•
•
•
•
•
•
•
•
•
privet leaf
teat pipette
carrot taproot (Daucus sp.)
hand lens
10 cm3 of acidified phloroglucinol
slides
forceps
tile
single-edged razor blade or sharp scalpel
microscope
calibrated eyepiece graticule
Introduction
In this practical, you will:
• examine the distribution of vascular tissue in roots, stems and leaves
• observe the two plant transport tissues, xylem and phloem.
Procedure
1
You are provided with a stem that has been standing in water coloured with a dye. Use a singleedged razor blade or a sharp scalpel to cut a thin cross-section. Put the cross-section on a
microscope slide and study it through a hand lens. If it is thin enough, you may be able to observe
it using the low-power objective of your microscope.
2
Draw a low-power plan to show the distribution of the coloured dye in the stem section.
Annotate your drawing to explain which tissues (xylem, phloem or others) contain the dye.
3
You are also provided with a piece of stem that has been standing in water, not a coloured dye.
Cut a similar section of this stem and place it on a microscope slide. Add some acidified
phloroglucinol to stain lignified cell walls. What colour do you see? Compare the two sections and
explain your observation.
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4
Use the low-power objective of your microscope to examine the prepared slide of a transverse
section (TS) of a stem. Draw a low-power plan to show the distribution of the tissues. Label your
drawing, using Figure 7.4 on page 129 of the Coursebook to help you. Use a ruler or calibrated
eyepiece graticule to measure the width of the stem section. Calculate the magnification of your
drawing and add this to the drawing.
5
Repeat steps 3 and 4 using a piece of carrot root and a prepared slide of a cross-section of a root.
6
Use a razor blade to cut across a leaf to give a cross-section. Use a hand lens to look at the surfaces
of the leaf and the section that you have cut. Notice the shape of the leaf in cross-section and the
position of the upper and lower surfaces.
7
Use a hand lens to look at the prepared slide of a TS of a privet leaf. Compare what you see with
the cut section of your leaf. Put the slide on the microscope and use low power and medium power
to focus on the leaf section. Make sure you are viewing it the right way round – with the upper
surface at the top of the field of view.
8
Repeat step 4 using the prepared slide of privet leaf. Use Figure 7.7 on page 130 of the
Coursebook to help you.
9
Construct a diagram to show the pathway taken by water as it travels through a plant from the soil
surrounding the root hairs to the atmosphere surrounding a leaf. In your diagram, show the
position of the xylem tissue in root, stem and leaf.
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Practical 7.2
Investigating xylem and phloem
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
a length of fresh celery (Apiaceae graveolens.)
petiole that has been left standing in water with a
little added eosin for 24–48 hours
vascular bundles teased out from celery petiole
tissue
prepared slide of TS of a young stem, (e.g.
cucumber (Cucurbita sp.) or sunflower
(Helianthus sp.))
prepared slide of LS of a stem (e.g. cucumber or
sunflower)
•
•
•
•
•
•
•
•
•
hand lens
slides
cover slips
tile
single-edged razor blade or sharp scalpel
forceps
mounted needles
microscope
calibrated eyepiece graticule
Introduction
In this practical, you will:
• study the microscopic structure of xylem and phloem tissues within the vascular bundle.
Procedure
1
You are provided with a petiole of celery that has been standing in water containing a coloured
dye. Use a single-edged razor blade or a sharp scalpel to cut a thin cross-section of the petiole.
Place the cross-section on a microscope slide and study it through a hand lens. If it is thin enough,
you may be able to observe it under the low power of your microscope.
2
Draw the cross-section of the celery petiole to show the distribution of the areas that are stained by
the dye. Label these areas.
3
Cut the celery petiole so you have a piece about 1 cm in length. Make another cut lengthways
through the middle of one of the vascular bundles. Now cut a thin longitudinal section from this
area and transfer your section onto a microscope slide. Observe the section using the low-power
objective to see if any xylem vessels are visible. If you have cut a thin enough section, you can
mount it in water, add a cover slip and observe using the medium-power objective. Use Figure 7.6
on page 130 of the Coursebook to help you identify xylem tissue. If you are not successful, repeat
this step until you cut a section that is thin enough.
4
Write an illustrated description of what you can see under the microscope.
5
You are provided with some vascular tissue that has been dissected from a celery petiole. Use
mounted needles to tease the white xylem tissue from the green phloem. Cut lengths of xylem and
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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phloem and mount them in water on separate microscope slides. Add cover slips. Observe using
the low-power and high-power objectives of your microscope. Add any further information you
have discovered to your description in step 4.
6
Use the low-power objective of your microscope to examine the prepared slide of a transverse
section (TS) of a stem. Find the xylem. Use the high-power objective to examine carefully the
xylem tissue. Compare what you can see with Figure 7.7 on page 130 of the Coursebook.
7
Using the high-power objective of your microscope to observe, draw three adjacent xylem vessels.
Use a calibrated eyepiece graticule to measure the width of one of the xylem vessels and indicate
this on your drawing. No labels are required.
8
Examine a prepared longitudinal section (LS) of stem under high power for xylem vessels. Add
any further observations to the illustrated description you made in step 4.
9
Look again at the prepared slide of a transverse section of a stem. This time, find phloem tissue.
Use the high-power objective to examine the phloem tissue carefully. Look for any sieve plates
with sieve pores – sometimes these are visible in cross-sections.
10 Make a drawing to show the details of three adjacent phloem sieve tube elements and their
companion cells. Use a calibrated eyepiece graticule to measure the width of one of the sieve tube
elements and indicate this on your drawing. Use labels to identify the two types of cell you have
drawn.
11 Examine a longitudinal section of stem using the high-power objective and look for phloem tissue.
Compare what you can see with Figure 7.23 on page 142 of the Coursebook.
12 Write an illustrated description of your observations of phloem tissue.
13 Construct a table to show the differences between xylem and phloem tissue that are visible using a
light microscope.
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Practical 7.3
Investigating the rate of transpiration of a leafy shoot using
a potometer
Safety
Take care when using sharps.
Take care when using mains-operated lamps in close proximity to water.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
length of capillary tubing (about 30 cm) with a
short connecting piece of rubber tubing attached
5 cm3 syringe
leafy twig with leaves that can be removed in
pairs (e.g. laurel (Laurus sp.)) in water
scissors
•
•
•
•
•
paper tissues
retort stand and clamp
lamp
stopwatch
petroleum jelly
Introduction
In this practical, you will:
• investigate the effect of leaf area on the rate of transpiration, using a simple photometer
• plan an experiment to find out how an environmental factor affects the rate of transpiration.
Procedure
1
Attach a syringe to the end of the capillary
tubing and draw water carefully through the
tubing. Submerge the syringe and the capillary
tubing in water and detach the syringe, ensuring
that there are no air bubbles left in the capillary
tubing.
2
Submerge the cut end of the leafy twig and
attach it to the capillary tubing under water,
making sure that the attachment is watertight
and being careful to exclude air bubbles.
3
Remove the twig and the attached capillary
tubing (see diagram opposite) and wipe excess
water from the surface of both using paper
tissues. Clamp the capillary tubing vertically and
place a lamp about 15 cm from the twig.
4
Measure and record the distance moved by the
water meniscus in the capillary tubing over a
five-minute interval. If necessary, repeat until
the rate of uptake has stabilised.
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5
Convert the distance l moved by the water in 1 minute into a rate of uptake, using the formula πr2l,
where r is the internal radius of the capillary tubing (in mm).
For example, if the internal radius is 0.75 mm, and the water moved 115 mm in 1 minute:
rate of uptake = π  0.75  0.75  115
= 3.14  0.75  0.75  115
= 203 mm3 min–1
6
Remove the apical (top) pair of leaves. Place a little petroleum jelly on the stem where each leaf
has been removed, and measure and record the rate of water uptake.
7
Continue removing successive pairs of leaves from the twig, each time measuring and recording
the rate of water uptake after removal of each pair. Remember to use petroleum jelly each time to
seal the stem where the leaves have been removed before recording.
8
Place the leaves on graph paper and trace round the margin of each leaf, numbering the tracings
according to the order in which they were removed. Calculate the surface area of each leaf in cm2
and record your results in a suitable form.
9
Plot a graph of the rate of transpiration (mm3 min–1) against the total leaf area (cm2).
10 Describe and explain the effect of the successive removal of leaves from the twig on the rate of
uptake of water.
11 Use the potometer to plan an investigation to find out the effect of a named environmental factor
on the rate of transpiration of a leafy shoot.
(Hint: Is a hair drier or fan available?)
If there is time, you could carry out your investigation; remember that your investigation must be
controlled, and you must only change one factor at a time.
12 What are the limitations of using a potometer to measure the rate of transpiration?
(Hint: What is the definition of transpiration? Does a potometer measure this?)
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Practical 7.4
Preparing a slide of epidermal cells from a lettuce leaf
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
microscope
calibrated eyepiece graticule
scalpel
lettuce leaf (Lactuca sp.)
slides and cover slips
forceps
•
•
•
•
•
mounted needle
dropping pipette
distilled water
filter paper
eye protection
Introduction
In this practical, you will:
• make a temporary preparation of epidermal cells from a lettuce leaf
• set up a light microscope and use it to make observations and measurements
• make a drawing of the epidermal cell
• use an eyepiece graticule to measure the size of the cells.
Procedure
1
Identify the upper surface of a lettuce leaf. It is darker than the lower surface. With the lower
surface facing towards you, gently tear the leaf, as you would tear a piece of paper. You should be
able to see the thin, transparent upper epidermis along the edge of the tear. This takes practice.
2
Select an area where the upper epidermis is visible and place a small piece of the leaf containing
this area onto a clean microscope slide, with the outer surface of the leaf facing upwards. Trim
away any thick (green) parts of the leaf using a sharp scalpel. To avoid folding the epidermis,
gently ‘roll’ the scalpel blade on the specimen and remove the unwanted tissue using forceps. You
should be left with a piece of the transparent epidermis a few millimetres wide on the slide.
3
Add two drops of distilled water to the slide. Place a cover slip over the water and lower it gently
onto the specimen using a mounted needle. Use a piece of filter paper to absorb excess water.
4
Observe the epidermis under the microscope, first using first low-power and then medium-power
objectives. The cell walls of the epidermal cells are visible as wavy lines looking rather like a
jigsaw puzzle. Between the epidermal cells, pairs of guard cells and stomata should be visible.
5
Use the high-power objective to select a few epidermal cells and a guard cell pair. Make a large,
labelled drawing of these cells.
6
Use the eyepiece graticule that you calibrated in Practical 1.1 to measure the length of one of the
guard cells that you have drawn. Now measure the length of this cell in your drawing.
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7
Calculate the magnification of your drawing, using the formula:
magnification =
length of drawing of cell
actual length of cell
Remember that both lengths must be measured in the same units, e.g. micrometres (m). Write the
magnification underneath your drawing.
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Practical 7.5
Investigating stomatal density
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
shoot from plant with simple, non-hairy
leaves, such as privet (Ligustrum)
bottle of colourless nail varnish
fine forceps
•
•
•
•
paintbrush
slides
microscope
calibrated eyepiece graticule
Introduction
In this practical, you will:
• compare the density of stomata on the lower and upper surfaces of a leaf.
Procedure
1
Spread a thin layer of nail varnish to cover about 1 cm2 over the lower epidermis of a leaf. Spread
another layer over the upper surface of a different leaf.
2
Allow the varnish to dry (this takes at least 20 minutes).
3
Peel off the layer of varnish from the lower surface of the first leaf, using fine forceps. Lay the
varnish on a slide and gently flatten it on the slide using a paintbrush. Do not use a coverslip.
4
Examine the varnish under the microscope and locate the imprints of the stomata. Try both
medium-power and high-power objectives and choose the most suitable magnification for counting
the number of stomata in the field of view.
5
Count the stomata in the field of view. Repeat twice more at different locations on the leaf surface
and calculate the average number of stomata visible in the field of view.
6
Repeat steps 3–5 for the varnish from the upper surface of the second leaf.
7
Measure the diameter of the field of view, using a calibrated eyepiece graticule.
8
Determine the area of the field of view using the formula πr2 and record the mean number of
stomata per mm2 for both lower and upper leaf surfaces.
9
Which surface has a greater stomatal density?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 7.6
Investigating the leaves of xerophytes
Safety
There are no special safety precautions.
Apparatus and materials
•

microscope
hand lens
•
•
calibrated eyepiece graticule
prepared slide of TS of a leaf of marram grass
(Ammophila arenaria)
Introduction
In this practical, you will:
• observe some of the adaptations shown by the leaves of a xerophyte, marram grass.
Xerophytes are plants that have adaptations for survival in dry conditions.
Procedure
1
Use a hand lens to look at the prepared slide showing a transverse section of a leaf of marram grass
(Ammophila arenaria). Compare this section with Figure 7.22 on page 141 of the Coursebook.
Also observe fresh leaves of marram grass, if these are available.
2
Study the diagram below, which is a plan drawing of a transverse section of a leaf of marram
grass.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Use the low-power and high-power objectives of your microscope to find the areas that
correspond with the structures listed below:
•
outer epidermis
•
inner epidermis
•
epidermal hair
•
vein
•
thin-walled mesophyll cells containing chloroplasts
•
thick cuticle
•
area of thick-walled cells providing support.
Indicate the position of each on the plan drawing using label lines.
3
Read the following information about marram grass.
Marram grass, Ammophila arenaria, grows on sand dunes around the coasts of western Europe. It
is the dominant species on young dunes where sand is deposited by the wind. Small heaps of sand
build up around marram plants and fresh shoots grow up through the sand that collects over them.
In fact, marram does not grow well where sand is not frequently deposited by the wind.
Sand drains very well and retains very little water. This means that only plants well adapted to dry
conditions can survive on sand dunes that are often exposed to strong winds. Rates of transpiration
are very high. Marram grass is well adapted to such dry, windy conditions. Marram grass is a
xerophyte. It grows in a dry habitat and can decrease its transpiration to a minimum under
conditions of water shortage.
In common with other grasses, marram grass has long, narrow leaves. In wet weather, the leaf is
flat and looks like many other grass leaves. In dry weather, cells located at the base of each furrow
lose water and shrink. These cells are called hinge cells because, when they shrink, the furrows
become narrower, making the leaf roll up to form a tube. When fully rolled-up, there is just a
narrow slit along one side of the tube to allow the entry of air.
4
Study Figure 7.21a on page 140 of the Coursebook, which shows a scanning electron microscope
(SEM) micrograph of a transverse section through part of a rolled leaf of marram grass. The inside
of the leaf has deep furrows that are lined by thin mesophyll cells full of chloroplasts. Much of the
rest of the leaf is made of large, thick-walled cells, which support the leaf, and veins that carry
water, nutrients and sugars. When rolled like this, the leaf has a small ratio of external surface to
its volume. The outer (lower) epidermis has a thick cuticle and no stomata. All the stomata are on
the inner (upper) epidermis where additional protection is provided by stiff interlocking hairs that
reduce the flow of air inside the leaf.
5
Using your observations and the information above, answer the following questions.
a
Draw up a table to show the differences between the upper and lower epidermis of marram
grass.
b Explain how and why the epidermal cells from opposite sides of the leaf of marram grass are
different.
c State three ways in which leaves of marram grass are adapted to survive in dry weather.
Explain how each of the methods you describe is effective in helping the plant to survive.
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Chapter 8 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 8.1
Microscopy of blood vessels
Most commercially prepared slides of artery and vein tissue show a muscular artery and a vein, as in
Figure 8.6 on page 161 of the Coursebook. It is possible to obtain slides of elastic arteries to compare
with the muscular arteries. Slides of the aorta are suitable.
Practical 8.2
Observing mammalian blood
Small volumes (25 cm3) of defibrinated mammalian blood (e.g. horse blood) can be bought from
biological suppliers. It will keep in a refrigerator for up to 4 weeks. Used slides and cover slips should
be placed in disinfectant (5% chlorine-based bleach solution, labelled ‘irritant’). Wright’s stain
contains eosin and methylene blue dissolved in methanol. It should be labelled ‘highly flammable’ and
‘toxic’. It is safer to purchase the ready-made stain.
It is not essential for students to wear gloves when handling sterile blood samples, but they may prefer
to do so to avoid getting any blood on their hands. Any blood that does get on hands should be washed
off with soap and water. Gloves should be worn when handling stains.
Eukitt® mounting medium is available from various suppliers (e.g. BDH). It will allow the smears to
be kept for several days, and does not cause the stain to fade. However, it is possible to observe the
cells without using a cover slip.
Students may ask the meaning of eosinophil and basophil. Eosinophils stain with acidic dyes such as
eosin. Basophils stain with basic dyes.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 8.3
Heart dissection
At the end of the practical, the hearts and any material cut from them should be wrapped up and
disposed of safely. Normally, sheep (or goat or ox) hearts obtained from butchers do not have intact
atria or major blood vessels attached. Obtain hearts direct from an abattoir, or ask a butcher to order
some for you with vessels attached. Alternatively, ask for hearts with lungs attached.
Students may want to put their textbooks into clear plastic bags to keep them clean during the
practical. Students should also wear surgical gloves during the practical.
Domestic chlorine-based bleach can be used as disinfectant. This will usually have a concentration of
less than 5%, which has a lower hazard rating. It should be labelled ‘irritant’.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 8.1
Microscopy of blood vessels
Safety
There are no special safety precautions.
Apparatus and materials
•
•
•
•
microscope
prepared slides of elastic artery (aorta), muscular artery and vein
prepared slide of a section of an organ such as mammalian kidney or
thyroid gland, for observing capillaries
calibrated eyepiece graticule
Introduction
In this practical, you will:
• compare the structures of arteries, veins and capillaries as seen through the light microscope.
You are provided with prepared slides of transverse sections of an elastic artery, a muscular artery and
a vein from a small mammal, as well as a section of an organ in which to observe capillaries. The
diagram below shows the tissues in a transverse section of a generalised mammalian blood vessel.
You should also refer to Figure 8.5 on page 160 of the Coursebook, which shows sections through an
artery, a vein and a capillary.
Procedure
1
Use the low-power and medium-power objectives of a microscope to examine the elastic artery.
Note that the structure of the elastic artery differs from that of the generalised blood vessel in the
diagram above. (In each comparison here and below, consider the size of the lumen, and the
thickness and composition of each tissue layer.)
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
2
Make a plan drawing to show the shape of the elastic artery and the distribution of the tissues
within it. Annotate your drawing to show how the elastic artery differs from the generalised
diagram above.
3
Use the calibrated eyepiece graticule to measure the thickness of the wall of the elastic artery. Add
this information to your drawing and construct a linear scale for the drawing.
4
Examine the muscular artery with the low-power and medium-power objectives of your
microscope. Note how the structure differs from both the diagram above and the elastic artery.
5
Make a plan drawing to show the shape of the muscular artery and the distribution of the tissues
within it. Annotate your drawing to show clearly how the muscular artery differs from the elastic
artery.
6
Use the calibrated eyepiece graticule to measure the thickness of the wall of the muscular artery.
Add this information to your drawing and construct a linear scale for the drawing.
7
Examine the vein with the low-power and medium-power objectives of your microscope. Note
how the structure differs from the generalised diagram and the two arteries you have looked at.
8
Make a plan drawing to show the shape of the vein and the distribution of the tissues within it.
Annotate your drawing to show how the vein differs from the diagram above and from the two
arteries that you have drawn.
9
Use the calibrated eyepiece graticule to measure the thickness of the wall of the vein. Add this
information to your drawing and construct a linear scale for the drawing.
10 a
Construct a table to show the differences in appearance between an elastic artery, a muscular
artery and a vein.
b Explain how the features of the two types of artery that are visible under the microscope are
adaptations for their functions within the mammal.
c
Explain how the features of the vein that are visible under the microscope are adaptations for
its function within the mammal.
11 Examine a section of an organ such as kidney or thyroid gland. Under high power, locate a
capillary seen in cross-section. Use the calibrated eyepiece graticule to measure the diameter of the
capillary. How does your measurement compare with the value given in the coursebook?
12 The wall of a capillary consists of a single layer of squamous epithelium. How does the size and
structure of a capillary relate to its function?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 8.2
Observing mammalian blood
Safety
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Wear gloves when handling stains.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
microscope
slides
cover slips (large, oblong type)
clean, dry Pasteur pipette
sample of sterile mammalian blood
medical gloves
slide-staining jar (or small beaker) containing
Wright’s stain (a mixture of eosin and
methylene blue in methyl alcohol)
•
•
•
•
•
•
•
two slide-staining jars (or small beakers)
containing distilled water
temporary mounting medium (e.g. Eukitt®)
glass rod
paper tissues
prepared slide of mammalian blood stained to
show white cells
100 cm3 beaker containing saline solution
100 cm3 beaker containing disinfectant
Introduction
In this practical, you will:
• use a microscope to examine a smear of mammalian blood and make observations of different
types of blood cell.
If fresh blood is available, start at step 1 of the procedure. If you are using prepared slides, start at
step 9.
Procedure
The blood used is sterile and it is not essential to wear gloves when handling it, but you may wish to
do so to avoid getting any on your hands. Gloves should always be worn when handling the stain.
1
Use a Pasteur pipette to add a drop of sterile blood to one end of a clean, dry microscope slide.
Immediately put the pipette into a beaker of saline solution.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
2
Use the edge of a second microscope slide to draw the blood across the first slide (see diagram
below). This spreads the blood cells thinly along the surface, forming a blood smear.
3
Allow the slide to dry completely. This normally takes a minute or two.
4
Dip the slide in Wright’s stain for 30 seconds, then lift the slide out and allow most of the stain to
run off the slide by holding the end against the wall of the staining jar.
5
Place the slide in the first jar of distilled water for 20 seconds, agitating gently. This removes
excess stain.
6
Transfer the slide to the second jar containing distilled water, to rinse off any remaining stain.
Leave it in the water for 20 seconds, agitating gently.
7
Remove the slide from the staining jar and leave it to dry by standing it vertically against the side
of the staining jar. Rest the end of the slide on a tissue.
8
When the slide is completely dry, place it flat, with the smear uppermost, on a dry tissue. Using a
glass rod, draw a thin line of temporary mounting medium along the middle of the length of the
smear. Place the cover slip on the mounting medium and leave for 10 minutes before examining
the slide under the microscope.
If you are using prepared slides, start here
9
Use the low-power objective of a microscope to locate some cells on your slide. You will need to
adjust the illumination and the focus of the microscope carefully to see the blood smear. Under
low-power objective, cells will just be visible as tiny dots.
10 Switch to the medium-power then high-power objective to examine the cells in more detail. Even
under high power (×400), the cells are very small. Most common are red blood cells
(erythrocytes), which are stained reddish pink. Occasionally, among the red cells you will see
larger cells with nuclei. These are the white blood cells. Search the slide to see how many types of
white cell you can find. (See table on the following page – the details in the table are for
information only, you do not need to know the names of the different types of white cell, just that
white cells have roles in phagocytosis and antibody production.)
11 Make a drawing of some blood cells. Include three red cells and as many different types of white
cell as you can identify using the diagram on the next page. You should certainly be able to find an
example of each of the commoner types, neutrophils and lymphocytes. Label your drawings, and
add the names of the cell types. Annotate your drawings with cell functions.
12 A red blood cell has a diameter of approximately 7 µm. Use this information to construct a scale
bar for your drawing.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
13 After you have finished with them, place all used slides and cover slips in a beaker of disinfectant.
Summary of appearance and functions of white cells (note: the Cambridge International AS and A
Level Biology syllabus only expects you to have knowledge of the three commonest types –
neutrophils, lymphocytes and monocytes)
Type
Diagram
Appearance
(when stained
with Wright’s
stain)
Function(s)
Normal
relative
abundance in
humans / %
neutrophil
dark-purple, lobed
nucleus; reddish,
granular cytoplasm
phagocytosis
30–75
lymphocyte
large, round, darkpurple nucleus
almost filling cell;
little cytoplasm,
non-granular
B lymphocytes
make antibodies
20–45
monocyte
large, light-purple,
bean-shaped
nucleus; grey-blue
non-granular
cytoplasm
phagocytosis:
develop into
macrophages
0–10
eosinophil
pale, lobed nucleus;
bright-red
cytoplasmic
granules
involved in defence
against larger
parasites
0–6
basophil
dark-purple
nucleus; darkpurple cytoplasmic
granules
involved in
allergies
0–2
Cambridge International AS and A Level Biology © Cambridge University Press 2014
T lymphocytes
involved in cellmediated immunity
3
Practical 8.3
Heart dissection
Safety
Take care when using sharps.
Wear surgical gloves.
Dissection instruments and boards should be washed with disinfectant.
Disinfectant is an irritant. Spillages should be washed off immediately using plenty of water.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
sheep heart (hearts of other animals, such as a
goat or ox, can also be used)
dissection board
scissors
forceps
•
•
•
•
blunt seeker
scalpel
disinfectant
surgical gloves
Introduction
In this practical, you will:
• identify the structures visible on the surface of the heart
• trace the pathway taken by blood as it flows through the heart
• dissect the heart to show how its structure enables it to carry out its function as a pump.
Procedure
1
Place the heart on a dissection board with the coronary vessels on the upper side. You should be
looking at the heart as seen from the front of the animal (a ventral view), as shown in Figure 8.22
on page 173 of the Coursebook. Check that the left ventricle is on the right-hand side – it will feel
solid when pressed. The right ventricle feels softer.
2
Use the photographs and drawings on pages 173–177 of the Coursebook to help you identify the
following structures:
 left and right ventricles
 left and right atria
 aorta
 pulmonary artery
 vena cava
 pulmonary veins
 coronary arteries.
Cut away any surplus fat to expose the major blood vessels at the top of the heart.
3
Make a drawing of the heart to show the main external features. Draw a ventral view, with the
apex at the bottom of the drawing. Label and add a scale to your drawing.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
4
Cut into the vena cava and then through the right atrium. From there cut into the wall of the right
ventricle down to its base. Clear out any clotted blood. Open up the chambers and wash out with
water if necessary.
5
Examine the valve between the right atrium and the right ventricle. Measure the thickness of the
walls of the two chambers.
6
Trace the pathway taken by blood as it leaves the right ventricle through the pulmonary artery.
7
Cut down the pulmonary artery to expose the semilunar valves.
8
Make a drawing of your dissection of the right side of the heart. Label the drawing. Use
annotations to describe the appearance of the structures you have drawn and their functions. Add a
scale to your drawing.
9
Cut into the left atrium and then into the left ventricle as far as the apex of the heart. Open the
chambers and clean out as before.
10 Examine the valve between the left atrium and the left ventricle. Measure the thickness of the
walls of the two chambers.
11 Trace the pathway of blood as it leaves the left ventricle through the aorta.
12 Cut down the aorta to expose the semilunar valves. Find the origin of the coronary arteries in the
aorta.
13 Make a drawing of your dissection of the left side of the heart. Label the drawing. Use annotations
to describe the appearance of the structures you have drawn and their functions. Add a scale to
your drawing.
14 Record all your measurements of wall thickness in a suitable way.
15 If there are any spare hearts that have not been dissected, make some cross-sections by cutting
across the hearts at different depths from top to apex. This will help you to appreciate the
differences in the thickness of the chambers.
16 When you have finished, dispose of the dissected material as instructed, wash the dissection board
and place the instruments into disinfectant. Wash your hands thoroughly.
17 a
Draw up a table to compare the structure and appearance of the four chambers of the heart.
b In one complete circuit through the heart, how does the output of blood from any one of these
chambers compare with that from the other chambers?
c
Explain how the valves you have displayed in your dissection ensure that blood flows through
the heart.
d Calculate the ratio between the thickness of the wall of the left ventricle and that of the wall of
the right ventricle.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 9 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 9.1
Investigating the mammalian gas exchange system
It may be possible to obtain lungs with hearts attached from a butcher or abattoir.
At the end of the practical, the dissected material should be wrapped up and disposed of safely.
Students should place dissecting instruments in disinfectant solution and wash their hands thoroughly.
Domestic chlorine-based bleach can be used as disinfectant. This will usually have a concentration of
less than 5%, which has a lower hazard rating. It should be labelled ‘irritant’.
Students can put their textbooks into clear plastic bags to keep them clean.
Practical 9.2
system
Microscopy of the tissues of the mammalian gas exchange
Students will find it easier to understand the sections of trachea and lung that they see through the
microscope and in photographs if they can dissect the organs as described in Practical 9.1.
Activity 9.1
Smoking survey
Students should be encouraged to construct simple hypotheses that can be tested easily by the survey.
They will need to spend time discussing the methodology and how to present their results. It may be
possible for them to use chi-squared (χ2) test with a contingency table on categorical data such as
male/female versus smoker/non-smoker.
Groups could present their findings to the rest of the class, either as a short presentation in PowerPoint
format or using overhead transparencies.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 9.1
Investigating the mammalian gas exchange system
Safety
Take care when using sharps.
Wear surgical gloves.
Dissection instruments and boards should be washed with disinfectant.
Disinfectant is an irritant. Spillages should be washed off immediately using plenty of water.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
lungs and bronchial system of sheep or goat
dissection board
scissors
forceps
•
•
•
•
scalpel
blunt seeker
surgical gloves
disinfectant
Introduction
In this practical, you will:
• dissect the lungs and bronchial system of a mammal to investigate the structure of the mammalian
gas exchange system.
Procedure
1
Place the lungs on a dissection board and arrange them on either side with their curved surfaces
facing upwards and the tubes towards the top of the board. Straighten out the tubes. You should
now be looking at the dorsal surface of the lungs, i.e. from the back of the animal.
2
Examine the external appearance of the gas exchange system. In order to see all of the organs you
will have to turn over the lungs so you can see them from the ventral view. Identify the following
structures:
• lobes of the left and right lungs
•
trachea
•
larynx
•
diaphragm
•
oesophagus.
The heart may still be attached to the lungs. If so, trace the pathway taken by blood as it flows
from the heart to the lungs and from the lungs back to the heart.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
3
Address the following points and questions.
a
How many lobes of the lungs are there?
b Describe what happens to the trachea as it passes between the lungs.
c
How can you tell the difference between the trachea and the oesophagus?
4
Examine the outside of the trachea and then run your finger, or a blunt seeker, down the inside.
5
Cut open the trachea from the top with a pair of scissors and examine the inside surface.
6
Follow the trachea down as far as you can by cutting it open, and find the points at which it
branches into the lobes of the lung. Continue cutting into the lung tissue to follow these airways
into the lungs. Examine the surface of the airways.
7
Make two cuts across the middle of the trachea either side of one of the hard bands that you will
have found.
8
Address the following points and questions.
a
Describe the internal surface of the trachea.
b How many major branches of the trachea have you found?
c
Where do these branches go? What are they called?
d Describe the arrangement of the hard material in the wall of the trachea. What is it made of? Is
the material arranged in the same way in the airways inside the lungs as in the trachea? If not,
describe the differences.
9
Cut through the lung tissue and examine its appearance. Look for small tubes that have a different
appearance from the airways that you have been following. These are blood vessels. The small
white vessels are arteries; the small pink vessels are veins. If the heart is present, try tracing the
veins back to the heart. (You will find it easier to locate some veins on the surface of the lungs and
start from there.)
10 Address the following points and questions.
a
Describe the appearance of the lung tissue. Explain why it is like this.
b What is the name given to the arteries in the lungs? What type of blood flows through them?
c
What is the name of the veins in the lungs? What type of blood flows through them?
11 When you have finished, dispose of the dissected material as instructed, wash the dissection board
and place the instruments into disinfectant. Wash your hands thoroughly.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 9.2
Microscopy of the tissues of the mammalian gas exchange
system
Safety
There are no special safety precautions.
Apparatus and materials
•
•
microscope
prepared slide of section of mammalian trachea
•
•
prepared slide of mammalian lung tissue
calibrated eyepiece graticule
Introduction
In this practical, you will:
• examine prepared slides of tissues from the mammalian gas exchange system and make drawings
of the tissues
Procedure
1
Examine a slide of a cross section of a trachea, using the low-power objective of your microscope.
If you have carried out Practical 9.1, compare what you can see with your observations when you
dissected the trachea.
2
Make a plan drawing of the trachea to show the arrangement of the tissues. Use Figure 9.3 on
page 188 of the Coursebook to help you identify these. Do not draw individual cells, just areas of
different tissues. Label the drawing.
3
Remove the slide from the microscope and measure the diameter of the trachea. Add this
information to your drawing, as a scale.
4
Annotate the drawing to show how the structure of the trachea is related to its function.
5
Now examine the section of trachea under high power. Look carefully at the tissue closest to the
lumen (the central air space). You should be able to find two cell types. Make a drawing of a small
number of cells from this tissue. Use labels to identify the cells that you have drawn. Annotate the
drawing to show how these structures are stained.
6
Examine a prepared slide of lung tissue under low power. Search the slide for bronchioles, alveoli,
arteries and veins.
You may need to look at several slides to find all of these structures. Look at Figure 9.3 on
page 188 of the Coursebook to help you identify the structures.
7
Selecting an objective lens with a suitable power, observe a group of three or four adjacent alveoli
and make a drawing of these. Label your drawing and use annotations to explain how alveoli are
adapted for gaseous exchange.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
8
Use a calibrated eyepiece graticule to measure the distance between the air in an alveolus and a
blood vessel. If you cannot do this, then measure the distance using Figure 9.5 on page 189 of the
Coursebook. Note that the diameter of a red blood cell is 7 m. This should help you to calculate
the distance that gases have to diffuse between the air and the blood.
9
How can you tell the difference between arteries and veins in the lung tissue?
10 How can you tell the difference between sections of trachea, bronchi and bronchioles when they
are viewed through the microsope?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 10 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 10.1
Testing bacteria for antibiotic sensitivity
Suitable species of saprotrophic bacteria are Bacillus subtilis and Micrococcus luteus.
Chromobacterium lividum, which produces dark purple colonies, can also be used, although it needs
glycerol in the nutrient agar for growth.
The principles of aseptic technique should be explained to the students. It is not possible for this
resource to deal with the details of aseptic technique, but teachers should note that aseptic methods
must be used at all stages in preparation of materials and growth media, transfer of cultures and
disposal of used apparatus and plates.
All used or contaminated materials should be steam sterilized in an autoclave or pressure cooker.
Further information on aseptic technique and microbiological laboratory practice can be obtained from
various sources, e.g.
• www.nuffieldfoundation.org/practical-biology/aseptic-techniques
• www.microbiologyonline.org.uk/teachers/safety-information/good-microbiological-laboratorypractise
This practical investigates the effects of antibiotics, using harmless species of cultured bacteria. Do not
attempt to culture bacteria from other sources such as soil or hands, which could contain pathogenic
species of microorganism.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 10.1
Testing bacteria for antibiotic sensitivity
Safety
Wear surgical gloves.
Aseptic technique should be used throughout.
Disinfectant is an irritant. Spillages should be washed off immediately using plenty of water.
Plates should be autoclaved after the investigation.
Apparatus and materials
•
•
•
•
•
broth cultures of two species of bacteria
Mastring™ antibiotic paper rings
two Petri dishes containing nutrient agar
sterile forceps
sterile glass pipette
•
•
•
•
•
marker pen or labels
sticky tape
incubator set at 25 °C
beaker of disinfectant
surgical gloves
Introduction
In this practical, you will:
• test two species of bacteria for their sensitivity to a range of antibiotics.
When a patient has a bacterial infection, doctors need to know which antibiotic will be most effective
in killing the organism responsible.
Eight antibiotics are supplied on a paper ring called a Mastring™. The Mastring™ is coded as shown
in the table below.
Coding for antibiotics on Mastring™
Letter code
Antibiotic
Disc colour
C
chloramphenicol (25 μg)
green
E
erythromycin (5 μg)
red
FC
fusidic acid (10 μg)
dark green
MT
methicillin (10 μg)
gold
NO
novobiocin (5 μg)
lilac
PG
penicillin G (1 unit)
pink
S
streptomycin (10 μg)
white
T
tetracycline (25 μg)
brown
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Procedure
1
You are provided with broth cultures of two species of bacteria. Using a sterile glass pipette,
transfer a pipette full of one broth culture onto a nutrient agar plate.
2
Swirl the plate around so that the liquid covers the whole of the agar surface. Discard the excess
liquid into a beaker of disinfectant.
3
Using sterile forceps, transfer a Mastring™ ring to the centre of the agar.
4
Replace the lid and tape it on. Label the base of the plate with your name, the date and the species
of bacterium used.
5
Repeat steps 1–4 with the second species of bacterium.
6
Place the plates upside down (so the label is facing upwards) in an incubator at 25 °C until
bacterial colonies have grown (about 24 hours).
7
Examine the plates without opening them for translucent zones around each of the antibiotic discs.
Measure the diameter of the translucent zones, and record your results in a suitable format.
8
List the order of effectiveness of the antibiotics against each bacterium.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 11 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Activity 11.1
Vaccination
Teachers should warn students to take care to use reliable sources such as the World Health
Organization and other respected medical authorities when doing their research. There are sources of
misinformation about vaccination on the Internet.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Chapter 12 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 12.1 Investigating the effect of temperature on dehydrogenase
activity in yeast
Both TTC and yeast are low hazard.
Actively respiring yeast suspension can be made by mixing 10 g of dried yeast with 100 cm3 of water
and adding 5 g of glucose. The suspension should be prepared 1 hour before use.
It is difficult to produce an end-point colour standard, since the mixture becomes progressively pinker
with time. This is a limitation of the experiment and, at best, the end point can only be judged to
about ±15 seconds. A suitable piece of pale pink card could be used for comparison of the colour.
A colorimeter cannot be used with a cloudy suspension.
The optimum temperature is higher than students expect, usually about 50–60 °C, with some decrease
in dehydrogenase activity at 70 °C, as the enzymes start to denature. This may be due to the fact that
denaturation is a time-dependent process, so that the equilibration time does not fully denature the
enzymes. It may also be that brewer’s yeast has been selected to have a high optimum temperature.
Another limitation is that it takes different amounts of time to equilibrate the tubes at different
temperatures: it is not possible to control this variable. However, use of a thermostatic water bath
would improve the precision of temperature control.
Respiration of the yeast cells will be partly aerobic but, in a boiling tube with little exposure to air,
probably mainly anaerobic. Students could be prompted to suggest methods of oxygenating the
mixture to allow aerobic respiration to continue, so that the rates of respiration under aerobic and
anaerobic conditions might be compared.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Practical 12.2 Investigating the rate of respiration of small organisms using a
simple respirometer
Soda lime contains a mixture of sodium, potassium and calcium hydroxides, and is very corrosive:
label the container in which the muslin bags of soda lime are supplied.
Eye protection should be worn throughout the practical.
Ensure students wash their hands after handing the living organisms.
There is probably too much material in this practical to carry out in a single laboratory session. The
work will need to be spread over two sessions, or shared between groups.
Instead of using a marker pen and ruler, mm-scaled adhesive tape can be used. This is available from
educational suppliers.
An alternative is to use a graduated pipette of a suitable volume (e.g. 1 cm3) instead of a capillary tube.
This avoids the need for calculating the volume (step 7).
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 12.1
Investigating the effect of temperature on dehydrogenase
activity in yeast
Safety
Take care when using heating apparatus.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
boiling tubes
10 cm³ syringe
large beaker (water bath)
tripod
Bunsen burner
test-tube holder
stirring rod
100 cm³ of a 10% suspension of actively
respiring yeast
•
•
•
•
•
•
•
•
test-tube rack
1 cm³ syringe
thermometer
gauze
heat-resistant mat
stopwatch
distilled water
10 cm³ of 0.5% triphenyltetrazolium
chloride (TTC) solution
Introduction
In this practical, you will:
• carry out a procedure at different temperatures between room temperature and 70 °C, to investigate
the effect of temperature on the activity of dehydrogenases in yeast.
Actively respiring yeast contains dehydrogenase enzymes. Normally when the yeast respires,
hydrogens are removed from the respiratory substrates and passed to hydrogen acceptors such as
NAD. It is possible to use an artificial hydrogen acceptor called triphenyltetrazolium chloride (TTC)
to show the activity of these enzymes. TTC is a redox indicator. It is colourless when oxidised and
pink when reduced. If TTC is mixed with yeast cells in suspension, some hydrogens will be passed to
the TTC, causing it to be reduced and change from colourless to pink.
Procedure
1
Use syringes to place 10.0 cm3 of the yeast suspension into a boiling tube and add 1.0 cm3 of
distilled water. This tube will act as a starting-point colour standard for the reaction, so that you
can see when a colour change has taken place in the experimental tubes.
2
Prepare a water bath at room temperature. Measure and record the temperature of the water in the
bath.
3
Place 10.0 cm3 of the yeast suspension into a clean boiling tube. Place 1.0 cm3 of TTC solution
into a second clean tube.
4
Place the two tubes in the water bath for several minutes to allow them to equilibrate to the
temperature of the water. How can you check they have equilibrated?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
5
Mix the contents of the tubes by adding the yeast to the TTC. Shake the tube and return it to the
water bath. Start the stopwatch.
6
Note the time taken for a definite pink colour to develop, by comparison with the starting-point
colour standard. Shake the tube gently at intervals to prevent the yeast settling to the bottom of the
tube.
7
Repeat steps 1–6 at five more temperatures between room temperature and 70 °C. At each
temperature, be careful to maintain the temperature of the water bath as constant as possible.
8
Present your results as a table. An arbitrary measure of the rate of reaction can be found by
1
calculating the reciprocal of the time taken for the colour to develop (rate =
).
time
1000
If you calculate the values of
, this will give more manageable numbers.
time (in seconds)
Add these values to the table.
9
Plot a graph of the rate of reaction (arbitrary units) against temperature.
10 Address the following points and questions.
a
What are the roles of dehydrogenase enzymes in respiration?
b
Describe what the graph tells you about the activity of dehydrogenases in yeast between
room temperature and 70 °C.
c
Explain why the activity is affected in this way.
d
What are the main sources of error and limitations of this experiment?
e
How could you improve the design of the experiment?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 12.2
Investigating the rate of respiration of small organisms using a
simple respirometer
Safety
Wear eye protection.
Soda lime is corrosive. If if it is spilled on skin, wash it off immediately with plenty of water.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
two 10 cm3 plastic syringes with short lengths
of rubber tubing attached
two 20 cm lengths of glass capillary tubing of
known internal diameter
water bath (trough or wide beaker)
small beaker of manometer fluid (coloured
water containing a drop of detergent)
marker pen
stopwatch
aluminium foil
•
•
•
•
•
•
•
two bags of soda lime wrapped in muslin to fit
inside syringe
maggots, mealworms or germinating mung
beans
paper tissues
balance weighing to at least 0.01 g
ruler
blunt forceps
eye protection
Introduction
In this practical, you will:
• measure the rate of respiration of some organisms (part A of the procedure)
• find the effect of a change in temperature on the rate (part B)
• measure respiratory quotients of the organisms (part C).
A respirometer can be used to measure the rate of uptake of oxygen by small organisms, such as
insects or germinating seeds. There are several different designs of respirometer. Figure 12.19 on
page 280 of the Coursebook shows a respirometer with a built-in control.
In this simple version, the organisms are placed in the barrel of a syringe.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
As the organisms respire, they use up oxygen. Any carbon dioxide produced is absorbed by soda lime.
The rate of uptake of oxygen can be found from the rate of movement of the fluid in the capillary tube.
Procedure
A Measuring the rate of respiration
1
Select some small living organisms, such as maggots, mealworms or germinating mung beans.
Check that they will fit comfortably into the barrel of a syringe, as shown in the diagram.
Weigh the organisms on a piece of aluminium foil.
2
Place some soda lime wrapped in muslin in the bottom of the barrel of the syringe. Introduce the
organisms that you have weighed into the syringe. Now carefully insert the plunger about half way
down the syringe.
3
Attach a length of glass capillary tube to the rubber tubing on the syringe. Place the respirometer
on the bench for about two minutes. Dip the end of the capillary tubing into the beaker of
manometer fluid. A small amount of coloured water will enter the capillary tube to make the
respirometer shown in the diagram. This is your experimental respirometer.
4
Place the respirometer horizontally on the bench and mark the position of the meniscus in the glass
capillary tube. Start a stopwatch.
5
Prepare another respirometer with the same mass of soda lime but do not add any organisms.
This respirometer is your control. Mark the position of the meniscus in the same way as in step 4.
Note the time.
6
After five minutes, mark the new position of the meniscus in the experimental respirometer tube
(if five minutes is not sufficient, wait for a suitable length of time). Similarly record any
movement of the meniscus in the control respirometer.
7
From these readings, calculate the volume of oxygen used (in cm3) by the organisms. If the
internal radius of the capillary tube (r) is known, the volume is found from the formula:
volume of oxygen used = distance moved by meniscus × πr2
8
Now find the volume of oxygen used per unit mass of organisms per minute, using the formula:
volume of oxygen used =
9
a
total volume of oxygen used (from step 7)
mass of organisms ´ time
What are the advantages of using of this method of measuring rates of respiration, and what
are its limitations?
b How should you use the readings from the control respirometer?
B
Changing the temperature
1
Design an investigation using the respirometer shown in Figure 12.19 on page 280 of the
Coursebook to compare the rate of respiration of the organisms at two temperatures within the
physiological range, such as 20 °C and 30 °C.
2
What effect would an increase in temperature have on the rate of respiration of the organisms?
Explain why temperature would affect their rate of metabolism.
C
Measuring respiratory quotients
1
Measure the rate of uptake of oxygen by the organisms as in part A.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
2
Detach the capillary tubes and dismantle the syringes of the respirometers. Use forceps to remove
the soda lime bags. Wipe out the inside of the syringes with a tissue. Replace the organisms in the
experimental respirometer, replace the plungers and reattach the glass capillary tubing to the
syringes.
3
Place the respirometers horizontally on the bench for about two minutes. Mark the position of the
meniscus in the capillary tubes.
4
After five minutes mark the new position of the meniscus in the experimental respirometer. Also
record the movement of the meniscus in the control respirometer.
5
The respiratory quotient (RQ) =
volume of carbon dioxide produced per unit time
volume of oxygen used per unit time
You have measured the rate of uptake of oxygen in step 1. Let this = x cm3 min–1. In step 4, the
manometer contains no soda lime to absorb carbon dioxide, so if the volume of oxygen used up
equals the volume of carbon dioxide produced, the respiratory quotient (RQ) = 1.
When more carbon dioxide is produced than oxygen absorbed, the scale shows an increase in the
volume of gas in the respirometer. Let this = y cm3 min–1. The RQ can then be calculated from:
RQ =
x+ y
x
Conversely, when less carbon dioxide is produced than oxygen absorbed, the volume of gas in the
respirometer decreases (let this volume z cm3 min–1). The calculation is now:
RQ =
x– z
x
Using your values from steps 1 and 4, calculate the respiratory quotient of the organisms.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
3
Chapter 13 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 13.1
Investigating pigments in a leaf by paper chromatography
Both propanone (acetone) and hexane (petroleum ether) are highly flammable and harmful. Label
containers accordingly. Turn off the electricity supply to avoid sparks. The chemicals are irritating to
the skin and eyes and have a narcotic effect if inhaled. Prolonged exposure to the vapours should be
avoided.
It is important to use freshly prepared chlorophyll extract for this investigation.
Students could extract and separate pigments from different plant species for comparison. Algae,
mosses or ferns could be used, to see if there are differences between the pigments present in these and
flowering plants.
Practical 13.2
Investigating the light dependent stage of photosynthesis
All solutions are low hazard. The normal safety precautions associated with the use of chemicals
apply.
To make a standard 0.05 mol dm–3 pH 7.0 phosphate buffer solution, dissolve 4.48 g of disodium
hydrogenphosphate-12-water (Na2HPO4·12H2O) and 1.70 g of potassium dihydrogenphosphate
(KH2PO4) in about 300 cm3 distilled water. Make up to 500 cm3 with more distilled water, then store
in a refrigerator at 0–4 °C until needed.
To make the ‘isolation medium’, dissolve 24.23 g of sucrose and 0.19 g of potassium chloride in about
150 cm3 of the phosphate buffer solution. Make up to 250 cm3 with more of the buffer solution, then
store in a refrigerator at 0–4 °C until needed. Use at room temperature.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
To make the ‘reaction medium’, dissolve 0.20 g of DCPIP and 0.93 g of potassium chloride in about
150 cm3 of the phosphate buffer solution. Make up to 250 cm3 with more of the buffer solution, then
store in a refrigerator at 0–4 °C until needed. Use at room temperature.
The investigation can be tried with filters of other colours such as blue.
A further control for the experiment would be a tube containing isolation mixture alone (i.e. DCPIP
but no leaf extract) to determine whether or not leaf extract is needed for the reaction.
Light intensity and temperature were not controlled in this investigation.
Students should explain that the dye is reduced by electrons emitted from the chlorophyll in the
presence of light, and compare the rate of reduction in green, red and white light.
Practical 13.3 Investigating the effects of limiting factors on the rate of
photosynthesis
All solutions are low hazard. The normal safety precautions associated with the use of chemicals
apply.
In part B, the light intensity will decrease with distance following the inverse-square law. If the
distance (D) is measured in metres, a graph of rate of photosynthesis (rate of oxygen production)
1
against 2 produces a curve which is fairly linear at lower light intensities but starts to level off at
D
high light intensities (i.e. with the lamp close to the plant).
Students may suggest that heat from the lamp will affect temperature of the syringe. This is true – a
transparent heat shield or ‘cold’ light source could be used instead of a filament lamp.
In part C, the plant should be supplied with a high concentration of hydrogencarbonate ions, so that
the supply of carbon dioxide is not a limiting factor, and a high light intensity (lamp at 10 cm from the
plant). The two temperatures should be chosen from the physiological range, e.g. room temperature
(about 20 °C) and 30 °C. A water bath can be used to control temperature (although in practice this
would be difficult with the apparatus used here).
Another investigation is to find the effect of changing the wavelength of light, using suitable coloured
filters, with the lamp at a fixed distance from the plant. The rate should be high with either red or blue
light, but much lower with green light, as predicted from an absorption spectrum of chlorophyll
(Figure 13.16 on page 295 of the Coursebook).
Practical 13.4
Adaptations of the leaves of C4 plants
Prepared slides of leaf sections of maize and sorghum are available from suppliers such as Philip
Harris Ltd., as well as slides with two sections, comparing leaves of C3 and C4 plants.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 13.1
Investigating pigments in a leaf by paper chromatography
Safety
Wear eye protection.
Propanone and the solvent are both highly flammable; the electrical supply should be turned off to
avoid sparks. Both chemicals are irritating to the eyes and skin, and cause drowsiness if inhaled.
The solvent (hexane) is also harmful if inhaled or swallowed.
Prolonged exposure to the vapours should be avoided.
Wear gloves to avoid touching the chromatography paper.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
•
fresh leaves, such as spinach
(Spinacia sp.) or nettle (Urtica sp.)
fresh leaves from a different species
pestle and mortar
clean sand
two 100 cm3 beakers
filter funnel and filter paper
two boiling tubes with corks to fit
pins
test-tube rack
•
•
•
•
•
•
•
chromatography paper, approximately 15 × 2 cm
(Whatman No. 1)
surgical gloves
dropping pipette
scissors
fine glass microcapillary tubes or pins
20 cm3 of 90% propanone (acetone) in a stoppered
tube or bottle
10 cm3 of solvent (1 part propanone to 9 parts hexane)
in a stoppered tube or bottle
Introduction
In this practical, you will:
• separate and identify the pigments present in a leaf by paper chromatography
• compare the pigments present in the leaves of different plants.
Procedure
1
Cut the leaves into small pieces and place them in a mortar. Add some propanone and a little clean
sand. Grind the leaves with the pestle until you have a dark green solution of chlorophyll. Filter the
mixture and collect the solution in a beaker.
2
Pipette the propanone/hexane solvent into a boiling tube, to a depth of about 15 mm. Seal the tube
with a cork and leave it for about 10 minutes, so that the air inside becomes saturated with the
solvent vapour.
3
Cut a piece of the chromatography paper so that it fits inside an empty boiling tube without
touching the sides. Draw a pencil line 25 mm from one end of the paper. Fold the other end of the
paper and use the fold to attach the paper to a cork with a pin. Replace the cork and check that the
paper almost reaches the end of the tube.
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1
4
Remove the cork and, using a fine glass microcapillary tube or the head of a pin, place a small
drop of the pigment mixture on the centre of the line. Allow the spot to dry. Repeat this several
times, allowing successive drops to dry. This will build up a small but concentrated spot of
chlorophyll.
5
Suspend the chromatography paper inside the first boiling tube, so that the end of the paper is in
the solvent, but the solvent does not reach the level of the spot of pigment.
6
Stand the tube in dim light for about 30 minutes, until separation of the pigments has taken place.
Then remove the paper and quickly use a pencil to mark the position reached by the solvent (the
solvent front). Leave the paper (chromatogram) in dim light to dry. It is a good idea to draw
around the pigment spots in pencil and note their colours, before they fade.
7
Measure the distance moved by each pigment from the origin (distance a) and the distance moved
by the solvent front from the origin (distance b). You should measure to the middle of each
pigment spot. Dividing a by b gives you the relative front value (Rf). The expected Rf values for
different pigments, using this solvent, are shown in the table below.
Values for Rf of various leaf pigments in propanone/hexane solvent
Pigment
Colour
Rf value
carotene
yellow
0.95
phaeophytin
yellow–green
0.81
xanthophyll
yellow–brown
0.71
chlorophyll a
blue–green
0.65
chlorophyll b
green
0.45
8
Determine the Rf values for each spot of pigment that you can see. Compare your values to those
in the table, and identify the pigments (you may have more than 5).
9
Suggest three factors that might affect the Rf values.
10 Repeat the investigation, using leaves from a different species of plant. Compare the pigments
present in the leaves of the two species.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 13.2
Investigating the light dependent stage of photosynthesis
Safety
The normal safety precautions associated with the use of chemicals apply.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
•
about 50 cm2 of fresh leaves
such as spinach (Spinacia sp.), cabbage
(Brassica sp.) or lettuce (Lactuca sp.)
scissors
glass rod
five lengths of fine glass capillary tubing,
such as melting-point tubing
aluminium foil
dropping pipette
two flat-bottomed plastic tubes
red and green filters
white tile
•
•
•
10 cm3 of a solution labelled ‘isolation
medium’ (4 mol dm–3 sucrose and
0.01 mol dm–3 potassium chloride dissolved in
a standard pH 7.0 buffer solution). This
solution should be chilled before use.
10 cm3 of a solution labelled ‘reaction medium’
(0.003 mol dm–3 2,6-dichlorophenolindophenol
(DCPIP) and 0.05 mol dm–3 potassium chloride
dissolved in a standard pH 7.0 buffer solution).
This solution should be chilled before use.
bench lamp
Introduction
In this practical, you will:
• investigate the effect of different conditions on the light dependent reactions of photosynthesis,
using DCPIP as a redox indicator.
DCPIP is a redox indicator that is blue in its oxidised form and colourless when reduced.
The apparatus and solutions are maintained at a low temperature until the reaction is started in order to
reduce enzyme activity.
Procedure
1
Remove any midrib or large veins from the leaves supplied. Cut the leaves into small pieces and
place these in a cold flat-bottomed plastic vial. Add 2 cm3 of the chilled isolation medium and
grind the leaves with a glass rod.
2
Pour off the extract into a second plastic vial covered with foil to keep the extract in the dark.
Remove a sample of the extract by inserting a length of fine capillary tubing into it. Lay the
sample tube on a white tile and use this as a colour standard with which to compare the contents of
the subsequent samples.
3
Using a dropping pipette, add 10 drops of reaction medium to the leaf extract in the foil-covered
vial and gently shake the vial to mix the contents.
4
Take a second piece of fine capillary tubing and remove a sample of the mixture from the vial.
Quickly wrap this sample tube in foil to prevent any exposure to light, and lay it on the tile next to
the first sample tube.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
5
Take three more pieces of fine capillary tubing and stand them simultaneously in the extract
mixture. Quickly place one of the pieces of tubing on the tile under a green filter, one on the tile
under a red filter and one on the tile without any filter, next to the colour standard.
At this stage, the contents of the pieces of tubing should be dark blue–green. If they are not, start
the investigation again.
6
Direct a light at the pieces of tubing and note the colour of their contents every three minutes for
fifteen minutes. Record your observations in a table like the one below.
Record of results
Tubing
Contents
Colour at 3-minute intervals
0 min
7
a
1
colour standard
2
extract + DCPIP in dark
3
extract + DCPIP in green light
4
extract + DCPIP in red light
5
extract + DCPIP in white light
3 min
6 min
9 min
12 min
Explain the purpose of the colour standard (sample 1) and the control (sample 2).
b What further control should have been set up and why?
c
Explain your results in terms of the light dependent stage of photosynthesis. You should
compare the rate of reduction of the DCPIP in each sample with the control sample.
d In which organelle and precisely where in the organelle is the DCPIP reduced?
e
State one important variable which was not controlled in this investigation.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
15 min
Practical 13.3
Investigating the effects of limiting factors on the rate of
photosynthesis
Safety
Take care when using sharps.
Avoid touching electrical apparatus with wet hands.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
fresh, washed Canadian pondweed
scalpel
tile
large beaker of aerated distilled water
10 cm3 syringe
20 cm length of glass capillary tubing,
internal diameter 1 mm
rubber tubing to connect syringe to capillary
tubing
•
•
•
•
•
•
retort stand, boss and clamp
lamp with 60 W bulb
stopwatch
25 cm3 of 0.05 mol dm–3 sodium
hydrogencarbonate solution
25 cm3 of 0.10 mol dm–3 sodium
hydrogencarbonate solution
25 cm3 of 0.15 mol dm–3 sodium
hydrogencarbonate solution
Introduction
In this practical, you will:
• investigate the effect of different concentrations of carbon dioxide on the rate of photosynthesis of
a plant (part A)
• investigate the effect of light intensity on the rate of photosynthesis (part B)
• plan an investigation to find the effect of temperature on the rate of photosynthesis (part C).
Carbon dioxide concentration, light intensity, and temperature can each act as a limiting factor on the
rate of photosynthesis
The plant you will use is Canadian pondweed, so the carbon dioxide is supplied as hydrogencarbonate
ions in solution.
Procedure
A
Investigating the effect of carbon dioxide concentration on the rate of photosynthesis
1
Collect some pieces of Canadian pondweed (Elodea canadiensis) and cut them across internodes
into approximately 5 cm lengths. Transfer them to a well-illuminated beaker of distilled water for
a few minutes and select two pieces that are bubbling rapidly and regularly.
2
Transfer the pieces into the barrel of a 10 cm3 syringe. Fill the syringe with aerated distilled water
and replace the plunger. Invert the syringe and expel any air.
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
1
3
Using a short length of rubber tubing, fix a length of capillary tubing to the nozzle of the syringe
and clamp the apparatus vertically (see diagram below).
4
Gently push down the plunger to force the water into the top of the capillary tubing. Place a lamp
about 10 cm from the syringe. Allow the pondweed to equilibrate for a few minutes.
5
Find the rate of movement of the meniscus over a suitable period of time. Assuming that the
movement is due to oxygen production by the plant, this will be proportional to the rate of
photosynthesis. The volume of oxygen produced will be equal to the distance moved by the
meniscus multiplied by the internal cross-sectional area of the tubing. The area can be calculated if
the internal radius of the tubing (r) is known, using the formula πr2.
rate of oxygen production = rate of movement of meniscus × πr2
6
Repeat this procedure, using:
 0.05 mol dm–3 sodium hydrogencarbonate solution instead of the aerated distilled water
 0.10 mol dm–3 sodium hydrogencarbonate solution
 0.15 mol dm–3 sodium hydrogencarbonate solution.
Each time, allow the pondweed to equilibrate to the new conditions before taking readings.
You should use the same pieces of weed in each solution.
7
Present your results in a suitable form.
8
a
What effect did an increase in concentration of hydrogencarbonate ions have on the rate of
photosynthesis?
b As the molarity of the hydrogencarbonate solution was increased by equal increments did the
rate of photosynthesis increase by equal amounts too? If not, why not?
c
If you continued to increase the molarity of the hydrogencarbonate solution, would you expect
the rate of photosynthesis to continue increasing?
d What other conditions might affect the rate of photosynthesis?
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
2
B
Investigating the effect of light intensity on the rate of photosynthesis
1
Set up the apparatus as described in part A, steps 1–4, filling the syringe with 0.15 mol dm–3
sodium hydrogencarbonate solution instead of distilled water, to ensure that carbon dioxide is not a
limiting factor.
2
Dim the background lighting in the laboratory by closing any blinds and turning off the room
lights.
3
Place the lamp 10 cm from the syringe. Allow the pondweed to equilibrate for a few minutes, and
then measure the rate of movement of the meniscus.
4
Move the lamp so that it is 15 cm from the syringe, and measure the rate again.
5
Repeat the measurements, with the lamp at increasing distances from the syringe. Increase the
distance by 5 cm intervals, until there is no production of oxygen by the plant. Each time, make
sure you allow the plant to equilibrate to the new light intensity.
6
Calculate the volume of oxygen produced by the plant at different distances and record these
values in a table. Plot a graph of rate of oxygen production (= rate of photosynthesis) against
distance of the light source (D).
7
The intensity of light produced by a light source is proportional to the reciprocal of the distance
from the source squared (1/D2). Calculate 1/D2 for each distance of the lamp, and plot a graph of
the rate of photosynthesis against these values. The rate should level off at higher light intensities,
as shown in Figure 13.7, page 291 in the Coursebook. Does your graph look like this?
C
Planning an investigation to find the effect of temperature on the rate of photosynthesis
1
Design an investigation to find out how two different temperatures affect the rate of
photosynthesis of pondweed, using the apparatus above. In your investigation, you should consider
the choice of temperatures you will use, and how to control other variables.
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
3
Practical 13.4
Adaptations of the leaves of C4 plants
Safety
There are no special safety precautions.
Apparatus and materials
•
•
•
prepared slide of a transverse section of a leaf of a C4 plant such as maize (Zea mays)
or sorghum (Sorghum sp.)
microscope
calibrated eyepiece graticule
Introduction
In this practical you will:
• examine a cross-section through a leaf of a C4 plant and consider how its structure is adapted to its
function.
Tropical grasses such as maize, sorghum and sugar cane are known as C4 plants. The leaves of C4
plants have a specialised photosynthesis biochemistry and a structure that differs from the leaves of C3
plants. They fix CO2 into a 4C compound in specialised mesophyll cells, and pass this fixed carbon to
bundle sheath cells, where it is decarboxylated and the CO2 used in the Calvin cycle. This arrangement
of leaf tissues isolates the Calvin cycle from atmospheric oxygen, avoiding photorespiration.
Photorespiration is a particular problem for tropical plants that live in conditions of high temperatures
and high light intensities (see Coursebook, Chapter 13, pages 293–295).
Procedure
1
Examine a transverse section of a leaf of a C4 plant such as maize or sorghum through the
microscope. Find a vascular bundle in the leaf and identify the bundle sheath cells forming an
inner ring around the vascular bundle, as well as the outer ring of mesophyll cells around the
bundle sheath.
2
Make a low power plan drawing of the leaf section. Label the specialised tissues you have
identified in step 1. Annotate these tissues to explain how they are an adaptation for the plant’s
survival in a tropical habitat (see Figure 13.13, page 294 in the Coursebook). Add a title and scale
to your drawing.
3
Observe the tissue of the vascular bundle under high power. Draw one or two bundle sheath and
mesophyll cells. Label and annotate the drawing and add a title and scale.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Chapter 14 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 14.1
Investigating the structure of the kidney
The kidneys supplied by butchers do not usually have the ureters or blood vessels attached. You will
need to ask for this when ordering them.
20 volume hydrogen peroxide solution must be labelled ‘irritant’.
Students must wash their hands after handling biological material.
Practical 14.2
Investigating the structure of kidney tubules
Students will benefit from seeing whole, fresh kidneys before studying the histology of the kidney.
They could also be shown photomicrographs of kidney tissue to supplement Figure 14.7 on page 305
of the Coursebook.
Prepared slides of longitudinal sections of mammalian kidney should show cortex, medulla, pyramids
and pelvis. These are sometimes described as horizontal sections. Slides of vertical sections of kidney
show cortex and medulla. There are also slides of transverse sections of kidney, which show only the
cortex.
Step 8 asks students to construct a table of appearance and function of a number of kidney structures.
Students’ tables should look similar to the one on the following page.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Epithelium
Microvilli
Width /
m
capillary:
squamous
outer epithelium:
squamous

100
•
ultrafiltration of blood
120
•
collection of filtrate

45
•
•
•
selective reabsorption
active transport of Na+ and Cl–
co-transport of glucose
15
•
loss of water from tubule by
osmosis
25
•
active transport of Na+ and Cl–
out of tubule
45
•
•
selective reabsorption
active transport of Na+ out of
tubule
active transport of K+ into
tubule
reabsorption of water
Glomerulus
•
Bowman’s
capsule
•
Proximal
convoluted
tubule
•
cuboidal
Thin loop of
Henle
•
squamous
Thick loop
of Henle
•
cuboidal
Distal
convoluted
tubule
•
cuboidal
Collecting
duct
•



 (few)
Function
•
cuboidal or
columnar

50–60
Cambridge International AS and A Level Biology © Cambridge University Press 2014
•
2
Practical 14.1
Investigating the structure of the kidney
Safety
Wear eye protection.
Take care when using sharps.
Hydrogen peroxide solution can bleach clothing or skin and cause burns. Spillages should be washed
off immediately using plenty of water.
At the end of the practical, the kidneys and any material cut from them should be wrapped up and
disposed of safely. Dissection instruments and boards should be washed with disinfectant.
Wear surgical gloves if you wish.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
fresh whole lamb’s kidney with vessels and
ureter attached
dissecting scissors
forceps
blunt seeker
scalpel
surgical gloves
•
•
•
•
•
dissecting board
10 cm3 of 20 volume hydrogen peroxide in
labelled container
dropping pipette
hand lens
eye protection
Introduction
In this practical, you will:
• examine the structure of the kidney and the relationships between its different parts.
You are provided with a lamb’s kidney. Note that it does not have quite the same structure as a human
kidney, nor is it quite like the rodent kidneys that are normally used to make prepared microscope
slides (Practical 14.2).
Procedure
1
Look at a diagram showing the position of the kidneys in the body of a mammal (for example,
Figure 14.5, page 305 of the Coursebook).
2
Examine the whole kidney provided. This should have parts of the blood vessels and ureter still
attached. See if you can find and identify these structures – you will probably have to remove
some fat to locate them. The concave part of the kidney where they are attached is called the
hilum. Note the overall shape of the kidney and place it on a board with the hilum to the left.
3
Remove all of the fat surrounding the kidney and make a labelled drawing of the kidney to show
its shape and external features. If the ureter and blood vessels are very short, use dotted lines on
your drawing to show where they would be. Measure the length of the kidney and include a scale
on your drawing.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
4
Use a scalpel to make a cut along the edge of the kidney on the convex side opposite the hilum.
This should follow the line you drew to show the outer edge of the kidney. Do not cut all the way
through the kidney yet.
5
You should now be able to see into the slit that you have cut. Inside there should be some white
tissue visible. This is the pelvis. On each side of the slit you will see pink tissue, partly covering
the pelvis (Figure 14.6 on page 305 of the Coursebook). This is the medulla. The darker tissue
towards the outside of the kidney is the cortex.
6
Look for a hole in the pelvis. Push a blunt seeker through the hole and see where it emerges.
If you are successful, you should find that it comes out through the ureter at the hilum.
7
Now continue to cut all the way through the kidney to produce two longitudinal sections. Note the
colours of the pelvis, medulla and cortex. As far as possible, trace the path of the renal artery from
the hilum into the cortex. The renal vein follows the same path, but is much more difficult to see –
try using a hand lens. In the cortex, the artery branches to supply the glomeruli and kidney tubules.
8
Use a hand lens to examine the cut surface of one section. In the cortex, you will see tiny red spots.
These are glomeruli. In the medulla, you should be able to see striations that run from the cortex
towards the pelvis. These are loops of Henle and collecting ducts. Compare what you see with
Figure 14.7 on page 305 of the Coursebook.
9
Use a pipette to add some drops of hydrogen peroxide to one of the cut surfaces. After the
vigorous effervescence has cleared, you should be able to see the structures within the cortex and
the medulla more clearly.
10 Make a labelled drawing of one of the cut surfaces of the kidney, use the same scale as your first
drawing. Annotate the drawing to show the functions of the structures that you have labelled and
add a scale.
11 When you have finished, dispose of the dissected material and board as instructed, and place the
dissecting instruments into disinfectant. Wash your hands thoroughly.
12 Address the following points and questions.
a
Describe the differences in appearance of the cortex, medulla and pelvis in the kidney that you
dissected.
b Why was there vigorous effervescence when you added some drops of hydrogen peroxide to
the cut surface of the kidney?
c
The kidneys make up about 0.5 to 1.0% of the total mass of the body, but they receive about
25% of the output of the heart. Explain why the kidney has such a large blood supply.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
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Practical 14.2
Investigating the structure of kidney tubules
Safety
There are no special safety precautions.
Apparatus and materials
•
•
•
prepared slide of LS of mammalian kidney,
labelled slide 1
hand lens
microscope
•
•
prepared slide of VS of mammalian kidney,
labelled slide 2
eyepiece with calibrated graticule
Introduction
In this practical, you will:
• investigate the fine structure of the kidney.
Procedure
1
Use a hand lens to look at slide 1, which is a stained longitudinal section (LS) of the kidney. Make
a drawing of the section. Label your drawing to show the different regions of the kidney. Add a
scale to your drawing.
2
Use the hand lens to look carefully for renal capsules. Indicate on your plan drawing the region
where these structures are found.
3
Examine slide 2, which is a stained, vertical section of the kidney. Use the low-power objective of
your microscope and concentrate on the area with glomeruli.
4
Using Figure 14.7 on page 305 of the Coursebook to help you, make a high-power labelled
drawing to show the structure of one Bowman’s capsule and glomerulus. In your drawing, show as
much detail of the cells of the capsule and glomerulus as you can see.
5
Use a calibrated eyepiece graticule to measure the distance across the capsule. Calculate the
magnification of your drawing and add this to the drawing.
6
Make a high-power drawing of cross sections of proximal and distal convoluted tubules. These are
found in the same region as the glomeruli. Draw to the same scale as your drawing of the
Bowman’s capsule and glomerulus. Annotate your drawing, noting the colours of the nuclei and
cytoplasm in the cells that you have drawn.
7
Move the slide so that you are looking at the medulla. You should be able to see thick and thin
parts of loops of Henle, as well as collecting ducts and capillaries. Search the slide to find crosssections of these structures and make a drawing to show a representative area with one example of
each structure. Identifying these structures in the medulla is difficult.
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1
Look for the following:
 thin parts of loops of Henle – thin tubes made of thin cells with nuclei that project into the
lumen
 thick parts of loops of Henle – as above but with thicker cells that are cuboidal in shape
 collecting ducts – wide tubes with cuboidal or columnar cells forming the lining
 capillaries – small, thin-walled vessels; it should be possible to see red blood cells inside.
8
Construct a table to show the visible features of the following structures, and their functions:
 glomerulus
 Bowman’s capsule
 proximal convoluted tubule
 thin loop of Henle
 thick loop of Henle
 distal convoluted tubule
 collecting duct.
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
2
Chapter 15 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 15.1
Measuring human reaction times
This is a familiar investigation, but in this form it can be used to illustrate a number of points about
experimental design, significance testing and limitations of the method.
Reaction times generally improve with practice, although they may get longer again as familiarity
sets in.
A t-test could be used to compare the mean reaction times from Tables A.1 and B.1.
The method is very crude, and it has many limitations. For example, it is not possible to control the
positioning of the ruler very precisely, or the experimenter may influence the subject. A possible
improvement is to use one of the many free reaction-timing programs available on the internet.
Human nerves transmit impulses at speeds between about 10 and 100 m s–1. The value calculated from
these experiments will be an underestimate. It does not take into account the delay caused by synapses
and the time taken for muscles to contract.
Practical 15.2
Investigating the effects of substances on muscle contraction
The muscle fibres are best taken from fresh meat. The key thing is to use fine strands, less than 2 mm
wide and longer than 10 mm. The laboratory technician or teacher could prepare these before the
lesson in which case, they must be kept moist in Ringer’s solution (use a Ringer’s tablet dissolved in
the specified volume of distilled water).
ATP solution or powder can be obtained from some school suppliers, such as Griffin Education. It is
probably best for the teacher to add the ATP to the muscle fibres using a syringe.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
The muscle fibres should show little or no change in length with water or glucose solution, but a
marked contraction with ATP. The water is a control, which shows that it is the ATP that causes
contraction, rather than the water in which it is dissolved. Glucose does not stimulate contraction, but
the muscle cells, even in this in vitro condition, are still stimulated to contract by ATP.
The main limitation of the procedure is that some water or glucose may be remaining on the muscle
before the ATP is added, so that the contraction could be due to the combined effects of ATP with
glucose (and water). The procedure could be improved by treating pieces of muscle separately with
just one of the three liquids. The percentage changes should be averaged and compared as before.
Practical 15.3
Investigating the effect of auxin on the growth of coleoptiles
It is important that the coleoptiles used have not been split by the emerging leaves. The timing of
germination of the seedlings needs to be carefully organised. It is best to germinate batches on
different days to make sure that some are ready when required.
Teachers may wish students to prepare their own logarithmic serial dilutions of auxin. However, this
can obscure the main point of the investigation.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 15.1
Measuring human reaction times
Safety
There are no special safety precautions.
Apparatus and materials
•
metre ruler
Introduction
In this practical, you will:
• measure human reaction times.
Reaction time is the interval between the moment a stimulus is applied and the moment when the
response starts. Measurement of reaction time can give some indication of the speed at which
information is transmitted in the nervous system.
Reaction time can be estimated by timing how long it takes for a human subject to catch a falling ruler.
The distance the ruler falls, d (in metres) = 12 at2
Equation 15.1
where a = the acceleration due to gravity (9.8 m s–2) and t = time (in seconds).
Rearranging this equation, we get:
t=
d
4.9
Equation 15.2
Using this simple method, you can test two hypotheses:
• hypothesis 1: the response time to a stimulus improves (decreases) with practice
• hypothesis 2: the response to a sight stimulus is faster than the response to a touch stimulus.
Procedure
A
Reaction time to a sight stimulus
1 Work in pairs: one person is the experimenter and the other the subject. The subject rests his or
her dominant arm horizontally on a table, with the hand held over the edge. The experimenter holds
the metre ruler vertically so that the 10 cm mark is between the subject’s thumb and fingers, but not
touching either. The experimenter must not indicate when he or she will release the ruler.
2 With the subject watching, the experimenter releases the ruler. As soon as the subject sees the
ruler falling, he or she tries to catch it between thumb and forefinger. The experimenter records the
distance the ruler has fallen, in metres.
3 Repeat steps 1–2 ten times, each time recording the distance the ruler fell. From equation 15.2,
calculate the time taken between the stimulus (sight of ruler falling) and response (gripping the ruler),
and record your results in a table like the one below. Calculate the mean reaction time and add to the
table.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Table A.1 Reaction time to sight stimulus
Drop number
Distance / m
Time / s
1
2
3
4
5
6
7
8
9
10
Mean time:
B
Reaction time to a touch stimulus
1 The subject closes his or her eyes, and the experimenter places the ruler between the subject’s
fingers, lightly touching his or her thumb.
2 The experimenter releases the ruler. As soon as the subject feels the ruler falling, he or she tries to
catch it between finger and thumb. The distance the ruler has fallen (in metres) is recorded.
3 Repeat steps 1–2 ten times and record the distance the ruler fell. From equation 15.2, calculate the
time between the stimulus (feeling the ruler falling) and response (gripping the ruler), and record your
results in a table like the one below. Calculate the mean reaction time and add to the table.
Table B.1 Reaction time to touch stimulus
Drop number
Distance / m
Time / s
1
2
3
4
5
6
7
8
9
10
Mean time:
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
C
Interpretation of results
4
From the data in Tables A.1 and B.1, address the following points.
a
Plot a graph of reaction time against drop number. Do the results support hypothesis 1?
b Compare the results in Tables A.1 and B.1. Do they support hypothesis 2? The mean times will
probably be different, but do you think the difference is significant? Your teacher may be able
to suggest a statistical test to help you decide.
c
Compare your results with the mean values for other members of the class. Are they in
agreement?
d Suggest limitations of this method, and ways the method might be improved.
e
D
Estimate the distance from the touch receptors in your hand, into the central nervous system
and back out to the muscles of the hand. From this distance and the mean time value from
Table B.1, calculate the speed of the signal. Is it true to say that this is the speed of the nerve
impulses involved? Explain your answer.
Using reaction timing programs
There are many websites on the Internet that contain simple programs to measure reaction times. Carry
out a search for these, select a suitable program and repeat the investigation. Compare your results
with the ‘dropping the ruler’ method.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
3
Practical 15.2
Investigating the effects of substances on muscle contraction
Safety
ATP is an irritant. Spillages should be washed off immediately using plenty of water.
Apparatus and materials
•
•
•
•
•
microscope slide
piece of fresh lean meat (e.g. beef)
small beaker of 1% glucose solution
filter paper
1 cm3 syringe
•
•
•
•
forceps
distilled water
1% ATP solution
two Pasteur pipettes
Introduction
In this practical, you will:
• investigate the effect of glucose solution and ATP solution on muscle fibres.
Meat is composed mainly of muscle fibres.
Procedure
1
Take a clean, dry microscope slide. Remove a fine strand of muscle fibres about 15–30 mm long
and 1–2 mm wide from a piece of lean meat. Use forceps to place it on the middle of the slide.
Arrange the strand so that it is straight.
2
Add a drop of distilled water to the muscle fibres. Leave the water on the fibres for about two
minutes, and then measure the length of the fibres in mm.
3
Drain the excess water from the slide, and use a piece of filter paper to soak up the water from
around the muscle tissue.
4
Add a drop of glucose solution to the fibres. Leave for two minutes as before, then measure the
length of the fibres again.
5
Remove the glucose solution as you did with the water in step 3.
6
Now add a drop of ATP solution using a syringe (your teacher may wish to do this for you). Leave
for two minutes as before, then re-measure the length of the fibres.
7
Calculate the percentage change in length of the muscle fibres:
a
when the glucose solution was added
b when the ATP was added, comparing each with the length in distilled water.
8
Collect the group results for the percentage change in length with glucose and with ATP. Calculate
the mean percentage length change with each solution.
9
Address the following points and questions.
a
What was the effect, if any, of glucose solution and ATP solution on the muscle fibres?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
b How variable were the class results? You could give a value for the variability, either as the
range, or better still by calculating the standard deviation.
c
Explain the effects, if any, of glucose and ATP solutions on the muscle fibres.
d The procedure involved adding solutions one at a time to the same piece of muscle tissue. Why
might this method be criticised? Can you suggest a more reliable procedure?
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 15.3
Investigating the effect of auxin on the growth of coleoptiles
Safety
Take care when using sharps.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
wheat or oat seeds
cotton wool
cork mat (or large cork)
beaker containing about 30 cm3 of 2% sucrose
solution
six boiling tubes
test-tube rack
marker pen
•
•
•
•
•
•
two Petri dishes
forceps
scalpel with new blade
access to the following concentrations of
auxin in 2% sucrose solution: 100 ppm,
10 ppm, 1 ppm, 0.1 ppm, 0.01 ppm
six bungs to fit boiling tubes
incubator set at 25 °C
Introduction
In this practical, you will:
• find out the effects of different concentrations of auxin on the lengths of pieces of coleoptile.
The shoot that emerges from a cereal seed is covered by a sheath known as a coleoptile.
Procedure
1
Soak some wheat or oat seeds in water overnight, and then place them on well-watered cotton
wool in a Petri dish (fill the dish with water and pour off the excess). Place about 50–100 seeds in
each dish.
2
When they have grown to a length of about 15–20 mm, but before the leaves have broken through
the coleoptiles, remove a coleoptile by pulling it from the base of the seedling, using forceps.
3
Place the coleoptile on a cork mat. Cut about 2 mm from the tip, using a sharp scalpel, and discard
the tip. As precisely as you can, cut a further 10 mm length of the ‘decapitated’ coleoptile. Place
this piece in a beaker of 2% sucrose solution until required.
4
Cut sixty 10 mm lengths of coleoptile, as described in step 3.
5
You are provided with a logarithmic serial dilution of auxin (indoleacetic acid, IAA) in
2% sucrose solution. The concentrations are as follows:
 100 ppm (parts per million)
 10 ppm
 1 ppm
 0.1 ppm
 0.01 ppm
 0 ppm (2% sucrose solution with no auxin, acting as a control).
Place 10 cm3 of each auxin dilution into a different boiling tube and label each tube.
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
1
6
Now add 10 lengths of coleoptile from step 4 to each tube. Pick up the coleoptiles gently with
forceps to transfer them to the tubes. Make sure that the coleoptiles are in contact with the solution
and that they do not stick to the sides of the tube. Place a bung in each tube.
7
Place the tubes in the dark, in an incubator at 25 °C for 2 days.
8
Tip the contents of a tube into a Petri dish. Remove the coleoptiles one by one and measure their
length in millimetres. Record your results in a table.
9
For each auxin concentration, calculate the mean increase (or decrease) in length of the
coleoptiles. Plot a graph of the mean percentage change in length against the concentration of
auxin. Note that the horizontal axis of this graph has a logarithmic scale reading from 0.01 to
100 ppm, with no zero.
10 Draw a horizontal line on the graph at the value of percentage change you have calculated for the
control group (the coleoptiles in 2% sucrose solution). Values above this line indicate that growth
has been stimulated, below the line indicates an inhibition.
11 Answer the following questions.
a
Why were the tips of the coleoptiles removed (step 3)?
b Which concentration of auxin produced the greatest stimulation to growth of the coleoptiles?
c
How does auxin act on the coleoptile cells to cause an increase in length?
d When pieces of tissue are investigated in test tubes like this, it is called an in vitro experiment.
This is Latin for ‘in glass’, as opposed to in vivo experiments, which are carried out on intact
organisms. Can you think of any limitations of an in vitro experiment of this kind?
Cam brid ge International AS and A Level Biology © Cam brid ge University Press 2014
2
Chapter 16 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 16.1
Observing the stages of meiosis in the testis of a locust
A stock solution of acetic orcein contains 2.2 g of orcein dissolved in 100 ml of glacial ethanoic
(acetic) acid, and is corrosive. Dilute 10 ml of this solution with 12 ml of water before use. Wear eye
protection and gloves. Carry out the preparation and dilution in a fume cupboard. The diluted solution
should be disposed of at the end of the practical.
Students should wear gloves when handling acetic orcein stain.
Either the desert locust (Schistocerca gregaria) or the African migratory locust (Locusta migratoria)
can be used, although the testes are easier to locate in Schistocerca. The specimen should be a young
adult or late 5th instar just prior to its final moult.
Suppliers of living organisms for use in schools and colleges can supply batches of male locusts (e.g.
see Blades Biological Ltd (www.blades-bio.co.uk) who take international orders).
The locusts should be killed humanely, by placing them in a killing jar containing ethyl ethanoate
(acetate). The insects must not come into direct contact with the chemical (e.g. place them on a
perforated zinc platform above the ethyl ethanoate).
Students who have ethical objections to the killing of animals for dissection should be excused from
taking part in the practical – they can observe the prepared slides.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Activity 16.1
Using models to investigate genetic crosses
There are no specific safety issues.
Teachers may prefer to use poppit beads instead of paperclips, although large quantities of paperclips
are cheaper to obtain than the beads. Any different colours can be used to represent the different
alleles.
Activity 16.2
The lac operon
There are no specific safety issues.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Practical 16.1
Observing the stages of meiosis in the testis of a locust
Safety
Wear eye protection.
Take care when using sharps.
Take care when using mains-operated microscopes with water or solutions.
Take care when using heating apparatus.
Do not inhale the acetic orcein fumes. Wear gloves when handling the stain.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
•
•
•
•
microscope
cover slips
glass rod
scalpel (no. 3 handle with new no. 11 blade)
cork mat
filter paper
slide hotplate
prepared slide of locust testis squash
surgical gloves
pins
•
•
•
•
•
•
•
•
slides
hand lens
fine forceps
dissecting scissors
acetic orcein stain
freshly killed male locust, either a young adult
or a 5th instar
eyepiece with calibrated graticule
eye protection
Introduction
In this practical, you will:
• dissect the testes from a locust
• stain testis cells to show the chromosomes, and observe the stages of meiosis.
The stages of meiosis can be observed in cells from the testis of a locust in what is known as a testis
squash preparation.
Procedure
1
You are provided with a freshly killed male locust. Wearing surgical gloves use scissors to cut off
the insect’s wings and legs and pin the body to a cork mat, with the dorsal surface upwards.
2
Use the scalpel and scissors to make a longitudinal cut down the length of the abdomen as shown
in the diagram on the next page. Open the abdomen to expose the contents, and pin back the flaps
of the body wall onto the cork mat.
Cambridge International AS and A Level Biology © Cambridge University Press 2013
1
3
Using a hand lens identify the testes. They lie above the gut, over the 5th and 6th abdominal
segments. They are surrounded by yellow fat, and are a bunch of sausage-shaped tubules.
4
Separate two or three tubules from the fat and place them on a microscope slide. Gently squash
the tubules with a glass rod to spread out their contents.
5
Add a few drops of acetic orcein stain to the tissue, and place a cover slip on top. Place a piece of
filter paper on top of the slide and cover slip, and press down gently to spread out the cells.
6
Remove the filter paper and gently tap the cover slip with a blunt instrument such as the end of a
scalpel handle. This helps to further flatten the testis cells and spread out the chromosomes.
7
Warm the slide on a hotplate for 30 seconds. This helps to intensify the stain.
8
Locate the testis cells under the low-power objective of the microscope. Use the high-power
objective to identify cells undergoing meiosis. If you have difficulty identifying any suitable cells,
you can use a prepared slide of a locust testis squash instead.
9
Make drawings of any stages of meiosis that you can see. Label and annotate your drawings
(Figure 16.9 on page 370 of the Coursebook). Use a calibrated eyepiece graticule to measure your
drawings, and add a scale bar to your drawing.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
2
Chapter 17 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Practical 17.1
Measuring population growth in a culture of Chlorella
Ready-prepared culture solution, or a prepared mixture of salts to make the solution, may be
purchased from educational suppliers.
Alternatively make Sachs’ culture medium, by dissolving the following salts in 1 dm3 of distilled
water:
• 0.25 g of calcium sulfate CaSO4·2H2O
• 0.25 g of calcium phosphate CaH4(PO4)2·2H2O
• 0.25 g of magnesium sulfate MgSO4·7H2O
• 0.08 g of sodium chloride NaCl
• 0.70 g of potassium nitrate KNO3
• 0.005 g of iron(III) chloride FeCl3·6H2O
The made-up culture solution is low hazard. If teachers or technicians prepare the culture solution
from the solid salts, they should note that solid potassium nitrate is oxidizing and dangerous with some
metals and flammable substances. Solid iron(III) chloride is harmful, and irritating to skin. It can cause
serious damage to eyes.
Some experimentation will be needed to find a suitable dilution of the culture that provides a
reasonable starting concentration of cells. Students will need to practise with the haemocytometer to
find the grids.
The students will need to be organised into a rota in order to take samples each day over the 15-day
period
It is also possible to measure the cell density in a colorimeter, using a blue filter (wavelength 410 nm)
to find the transmission of the cell suspension against a blank of pure culture solution.
Cambridge International AS and A Level Biology © Cambridge University Press 2014
1
Relate the exponential growth phase to Darwin’s observation that populations in ideal conditions can
grow very fast, overproducing offspring until the carrying capacity of the environment is reached, due
to limiting factors.
Other microorganisms can be used, such as the alga Scenedesmus; or yeast cells, which do not need
illumination.
Practical 17.2
Measuring variation in a plant population
The nature of the area to be sampled should be considered in the risk assessment.
You should check the school grounds to select suitable species for investigation. A t-test or z-test
(for sample sizes >30) can be used to compare the means of two samples.
Activity 17.1
A simulation of natural selection
Coloured pieces of wool can be used as an alternative to toothpicks.
The size of the groups and the sampling area can be adjusted to suit class sizes; between 10 m  10 m
and 15 m  15 m is suggested. Selection from other habitats, such as bare soil, can be investigated.
With increasing predation, the selective advantage of the green ‘insects’ is reduced.
There are very many assumptions and limitations of this simulation. For example, the ‘insects’ don’t
move or hide; real birds may have a different way of locating insects, and so on.
Activity 17.2
Using a model to test the Hardy–Weinberg principle
Teachers may prefer to use Poppit beads instead of paperclips, although paperclips are cheaper. Any
two different colours can be used to represent the different alleles.
The expected frequencies of genotypes (as in the parental generation) are:
AA = p2 = 0.42 = 0.16
Aa = 2pq = (2 × 0.4 × 0.6) = 0.48
aa = q2 = 0.62 = 0.36
So the expected numbers of each genotype are:
AA = 0.16 × 50 = 8
Aa = 0.48 × 50 = 24
aa = 0.36 × 50 = 18
Calculation of the χ2 statistic is explained in the Coursebook (pages 386–387). The most likely
explanation for a rejected null hypothesis is the small size of the population, or a bias in the choice of
gametes (e.g. if the person selecting the paperclips can tell the difference between red and yellow in
some way – this would be equivalent to non-random mating).
To show the effects of natural selection against the recessive allele (step 12), the students should start
by carrying out the steps 1–5 as before.
Let’s say that the observed numbers in the next generation were as expected (i.e. 8 × AA, 24 × Aa and
18 × aa).
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The recessive homozygotes die, so students should now remove 36 yellow paperclips, but increase the
numbers to the original population size by adding more paperclips in the ratio of those remaining (i.e.
40 red and 24 yellow). In this case, it would need a total of:
 24 
 40 
  × 100 = 62 red and   × 100 = 38 yellow (to nearest whole numbers).
 64 
 64 
The students then repeat the exercise. After a few generations, the selection against the yellow allele
will result in a decrease in its frequency in the gene pool. However, it is unlikely to be completely
eliminated from the population, being maintained in the heterozygote. This nicely illustrates the effect
of selection, which disturbs the Hardy–Weinberg equilibrium. The students could plot a graph of the
frequency of allele a (starting at 0.6) against generation.
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Practical 17.1
Measuring population growth in a culture of Chlorella
Safety
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
microscope
haemocytometer slide and coverslip
Pasteur pipette
100 cm3 measuring cylinder
250 cm3 conical flask
•
•
•
•
•
cotton wool
10 cm3 syringe
fluorescent light bank or Gro-lux tubes
actively growing, dense culture of Chlorella
culture solution
Introduction
In this practical, you will:
• use a haemocytometer slide to count Chlorella cells
• investigate the phases of growth of a pure culture of Chlorella over a period of 15 days.
Chlorella is a unicellular freshwater alga which rapidly increases in numbers by asexual reproduction.
Procedure
A
Using a haemocytometer to count cells
1
Look at the haemocytometer. There are different types of these. Most have two small grids etched
onto the glass between an H-shaped drainage well (as shown in diagram (a) below) but some have
one central grid (as shown in diagram (b) below).
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2
Select the low-power objective lens of the microscope. Place the haemocytometer on the stage of
the microscope and line up one of the grids under the lens, by eye. Now look through the
microscope and adjust the focus until you can see a clear image of the grid (this takes practice).
The large middle square of the grid measures 1 mm by 1 mm, and consists of 25 medium-sized
squares (diagram (a) below). Switch to the high-power objective, and you will see that each
medium-sized square is surrounded by triple lines and is further divided into 16 small squares
(diagram (b) below).
When a cover slip is placed on the haemocytometer, the grids are 0.1 mm below the cover slip.
Therefore the volume above each of the 25 medium-sized squares is (0.2  0.2  0.1) mm3 =
0.004 mm3. If some solution containing cells is placed onto a haemocytometer, you can count the
cells and work out the number of cells per unit volume. Box P1.2, page 251 in the Coursebook
provides additional information
3
Using a measuring cylinder, transfer 100 cm3 of culture solution to a 250 cm3 conical flask.
4
Using a syringe, remove a 10 cm3 sample from an actively growing, dense culture of Chlorella.
Add this to the solution in the flask, which will dilute the culture about 100 times. Shake gently to
mix the cells and solution. Stopper the flask with cotton wool in between removing samples.
5
Using a Pasteur pipette, add a drop of dilute culture to the haemocytometer, placing the drop above
the grids. Place the cover slip on top and allow any excess liquid to run off into the drainage well.
6
Focus on one of the 25 medium-sized squares. Count the number of cells in the square. For cells
touching a line, count only those touching the top and right-hand sides of the square, and not those
touching the bottom or left-hand sides. Repeat the count for five squares (you can use the other
grid too).
7
Calculate the mean number of cells per square. If you now multiply this figure by 250 000 you will
get the number of cells in 1 cm3 of the culture solution.
B
Counting the number of Chlorella cells over 15 days
1
Place the culture flask under a ‘cold’ light source, such as a fluorescent light bank or Gro-lux
tubes, and maintain the flask at room temperature. The cells will use the light to carry out
photosynthesis, and reproduce asexually.
2
Take a sample of the culture and count the cells once a day, at about the same time every day.
Continue taking samples for about 15 days. Problems with sampling at weekends can be overcome
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by setting up several diluted cultures, with the same initial cell densities, over three days. This will
mean that there will be at least one sample for each of the 15 days.
3
Plot a graph of the cell density against time.
4
Describe the shape of the curve on your graph. Does it show any evidence of the Chlorella cells
undergoing an exponential growth phase? If a maximum cell density is reached, can you suggest
any abiotic factors which may be causing growth and reproduction of the cells to reach a limit?
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Practical 17.2
Measuring variation in a plant population
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
ruler
•
gloves
•
population of a suitable plant species
Introduction
In this practical, you will:
• measure continuous variation in a population of a plant species and present the results graphically
• use the t-test to compare the means of two sets of measurements.
Procedure
1
Select a common local plant species that shows variation such as:
 length of leaves, or leaf area
 length of stem or flower stalk
 mass of inflorescence.
2
Plants for measurement should be taken at random from the population rather than by selection as
this introduces bias in the sampling. You may be able to use random coordinates to locate the
plants to measure (see Practical 18.1)
Measure each of the variables in a large number of plants from the population (>40). What is the
best precision you can achieve? For example, it will probably not be possible to measure leaf
length to a precision of better than ±1 mm.
3
Collate the data in a tally chart, as shown below.
Tally of leaves of specific lengths
Leaf length/mm
Number of leaves
25
|
26
||
27
|||
28
||||
|||
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For some variables you may need to combine the categories as shown in the chart below. Note that
the categories do not overlap, and the length values reflect the precision of the measurements.
Tally of stems in length ranges
Stem length / mm
Number of stems
0–4
|
5–9
10–14
15–19
4
||||
||||
||
Plot a frequency histogram of the number of leaves (or other variables) against length categories.
Your histograms should look something like the diagram below.
Does each of your measured variables show a normal (bell-shaped) distribution?
5
Select two populations of plants of the same species, living in different habitats. From your
observations of the populations, identify a variable by which they may differ. For example, you
might suspect that a population of plants living in a shady habitat has smaller leaves than a
population growing in an open field where there is more light.
6
Measure a sample of 15 randomly selected leaves from each population of plants.
7
Carry out a t-test to establish the significance of the difference in the mean leaf length of the two
populations.
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Chapter 18 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers before they are presented to students. Additional notes
relating to each activity in this chapter are given below, but should not be regarded as risk
assessments.
The nature of the areas to be sampled should be considered in the risk assessments.
Practical 18.1 Investigating the distribution of plants in two habitats, using
random sampling
You should visit the field sites to identify suitable species to record and to select two areas for
comparison. Keys or identification sheets should be prepared that show all the common plant species
found in the area. A simple identification sheet can be made by photocopying leaves of the common
species. The names can then be written onto the photocopy paper, and the sheets laminated.
The Braun–Blanquet scale is a relatively simple abundance scale with a few large categories. It is
particularly suitable for use in species-rich communities.
Practical 18.2
Sampling animals from a terrestrial habitat
You should visit the field sites to select areas suitable for sampling. Keys or identification sheets
should be obtained that show all the invertebrates likely to be found in the habitat.
A beating tray can be made by attaching a sheet to a wooden frame about 0.5 m square. Commercially
made pooters are available, or they can be easily constructed using a small collecting tube, bung, glass
and plastic tubing. Pitfall traps can be made from a plastic cup or glass jar sunk in the ground. A cover
is needed to prevent rain collecting in the jar.
If a suitable aquatic habitat such as a pond or stream is available, sampling can be carried out using a
metal-framed pond net, which is made of a tougher material than a sweep net. Very fine-meshed
plankton nets can be used to sample plankton.
Students should be encouraged to think about the controls needed when comparing two habitats. The
sampling time and area from which the sample is taken must be the same for each habitat.
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Practical 18.3 Investigating the distribution of plants along an environmental
gradient, using a transect
Teachers should identify a suitable field site, such as a trampled path though the school field, or a
gradient from dry to marshy ground. Keys or identification sheets should be prepared that show all the
common plant species found in the area. A simple identification sheet can be made by photocopying
leaves of the common species. The names can then be written onto the photocopy paper, and the
sheets laminated. The Field Studies Council also publishes identification charts obtainable from:
www.field-studies-council.org/publications/fold-out-charts.aspx
Students could extend the investigation by measuring factors which might affect the plant distribution,
such as the degree of soil compaction, or the soil water content. Compaction can be measured using a
penetrometer – a simple instrument that measures the depth a spike is driven into the ground by a
standard force. The percentage water content can be measured by heating a known mass of soil in an
oven at 100 °C until the weight remains constant.
A point quadrat is easier to use than a frame (area) quadrat, avoiding subjective estimations of
percentage cover, or difficulties in counting numbers of plants in a field. However, the exercise could
be carried out with a frame quadrat if required.
The distribution of each species across the transect can be plotted as a histogram (Figure PG18.1).
Figure 18.1 Histogram showing the distribution of a species across a transect.
The histograms should be arranged one above the other for comparisons between species.
Alternatively, the distributions can be plotted as kite diagrams. These are normally used for rocky
shore transects, although there is no reason why they can’t be used for other habitats. The counts of
‘hits’ on the y-axis should be an ordinal variable; for example, an abundance scale of ranks, such as an
ACFOR scale (abundant, common, frequent, occasional, rare).
Figure PG18.2 on the next page shows how to plot a kite diagram.
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Figure 18.2
Kite diagram showing the distribution of a species across a transect.
Practical 18.4 Measuring the diversity of broad-leaved plant species in two
habitats, using Simpson’s Index of Diversity
You should visit the field sites to select two areas for comparison. Keys or identification sheets should
be prepared that show all the common plant species found in the area. A simple identification sheet
can be made by photocopying leaves of the common species. The names can then be written onto the
photocopy paper, and the sheets laminated.
Practical 18.5
Observing single-celled organisms
Students should wash their hands after handling cultures.
For drawing, Paramecium can be slowed down by using viscous solutions such as Protoslow available
from Blades Biological (www.blades-bio.co.uk/). Some species of Chlorella are free-living; others
exist symbiotically inside other protoctists such as Paramecium.
Practical 18.6 Using mark–release–recapture to estimate the size of a
population of snails
Students must wash their hands after handling biological material.
The nature of the area to be sampled should be considered in the risk assessment. The habitat must be
well defined.
Check the school or college grounds to select a suitable species for investigation. Snails are a good
species to use because they are easy to mark on the shells, and they are likely to remain within the
habitat. Other species such as woodlice can be used.
The ratio of the number of individuals caught and marked in the first sample (S1) to the total number in
the population (N) is equal to the ratio of the number marked and recaptured (R) to the total caught in
the second sample (S2).
S1 R
=
N S2
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Rearranging the equation gives:
N=
S1 ´ S2
R
The data supplied in step 7 gives (to nearest whole number):
N=
63´ 78
= 328
15
The snails should be collected at the same time of day in each sample, because they may change their
pattern of distribution at different times. For example, they may feed on leaves at night and hide under
stones during the day. Sampling at different times may bias the results.
The method will not give reliable results in a poorly defined habitat, or where there is a degree of
immigration or emigration, since the population size will vary. The marking technique must not harm
the snails and they must not be made more conspicuous to predators, which will reduce the size of the
population. The technique should not alter the habitat, which could also lead to a decrease in the
population.
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Practical 18.1
Investigating the distribution of plants in two habitats, using
random sampling
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
two 10 m measuring tapes
(or string with knots at 1 m intervals)
four strong pegs (if using string)
0.5 m × 0.5 m (0.25 m2) frame quadrat
•
•
random number table
(or calculator with random number function)
plant keys or identification sheets
Introduction
In this practical, you will:
• investigate the abundance of plant species in two habitats using a random sampling technique.
In a relatively uniform habitat such as a school field, the method of random sampling can be used to
investigate the abundance of plant species present. Quadrats are placed randomly in the area to be
sampled, using random numbers to generate coordinates. Note that throwing quadrats is not random.
Procedure
1
Mark out an area of ground 10 m × 10 m in the chosen habitat, using measuring tapes or string.
2
Use a random number table or a calculator with a random number function to generate a pair of
random numbers (from 0 to 9) to supply the coordinates for placing the first quadrat.
3
Locate the coordinates along the tapes and place the bottom left corner of the quadrat at the
intersection of the coordinates.
4
Count the number of different species of plants within the quadrat. It helps if you can identify the
species, using a plant key or identification sheets. If there are difficulties with identification, just
call them species A, species B, and so on. There is no need to count the total numbers of each
species.
5
Now select a common plant species. Let’s call the species that you have selected, species A.
6
Record the total numbers of individual plants of species A within the quadrat.
7
Estimate the percentage cover of species A within the quadrat, and convert the percentage to the
Braun–Blanquet scale using the table on the next page.
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Percentage cover and the Braun–Blanquet scale
% cover
8
Braun–Blanquet scale
<1
1
1–5
2
6–25
3
26–50
4
51–75
5
76–100
6
Repeat steps 2–7 another 9 times, and record your results in a table like the one below.
Record of results
Quadrat
No. of species
in quadrat
Presence / absence No. of individuals
of species A
of species A
Cover on the Braun–
Blanquet scale
1
2
3
4
5
6
7
8
9
10
9
Repeat the sampling in a different area. Try to select areas with visible differences in vegetation.
10 For each habitat, calculate the mean species richness (average number of species per quadrat).
11 For species A, calculate the species frequency, the mean species density and the mean cover on the
Braun–Blanquet scale in each habitat, using the following equations:
Species frequency = % of quadrats in which the species was found
Mean species density (per quadrat) =
sum of number of individuals in quadrats
number of quadrats
Mean cover on the Braun–Blanquet scale =
12 a
cover per quadrat
number of quadrats
Compare the differences in plant distribution in the two habitats. Can you think of any reasons
why the two areas might have different plant communities?
b What are the limitations of random sampling to measure the distribution of plants in the two
habitats?
c
What are the limitations of the different methods used to measure the distribution of species A?
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Practical 18.2
Sampling animals from a terrestrial habitat
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
beating trays
pooters
pitfall traps
hand lenses
keys or identification guides
small (artist’s) paintbrushes
•
•
•
•
•
trowels
trays
sweep nets
collecting jars
gloves
Introduction
In this practical, you will:
• use different pieces of apparatus to collect small invertebrates from a terrestrial habitat
• plan investigations using the apparatus.
Small terrestrial invertebrates can be sampled using various simple pieces of collecting equipment (as
shown in the diagram below).
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1
A beating tray is a large sheet supported by a frame. It is held underneath a bush or tree branch, which
is then shaken vigorously to dislodge organisms onto the sheet. The organisms are transferred to a
tray, where they can be collected and identified.
A pooter is used to suck small invertebrates into a collecting tube.
Sweep nets are passed back and forth through long grass to collect insects or other similar organisms.
A pitfall trap is a plastic cup or jar sunk into the ground to trap animals walking over it.
Take care not to harm the animals when handling them – an artist’s paintbrush is useful for this.
Return the organisms to their habitat after you have identified them.
Procedure
A
Using collecting equipment
1
Select a suitable terrestrial habitat from which to collect small invertebrates such as insects, for
example a field, hedge or tree.
2
Use the beating tray to collect small invertebrates from a hedge or tree branch. Collect the
organisms in specimen jars or using a pooter. Use keys or identification guides to identify the
animals you have caught.
3
Set up pitfall traps in a suitable spot, such as the edge of a field. The traps should have a cover to
keep out rain. Leave them for 24 hours before emptying their contents into trays and identifying
the animals.
4
Use a sweep net in a suitable habitat, such as long grass. Carefully remove any insects or other
invertebrates that you catch. Transfer the organisms to collecting jars and identify them.
B
Planning investigations
Plan investigations to test the hypotheses below. In your plan, state how any comparisons between two
habitats will be controlled – how will you ensure that sampling is fair and unbiased? If possible, carry
out the investigations.
Hypotheses
1 The invertebrate community living on one species of bush or tree is different from the community
living on another species of bush or tree.
2 Invertebrates living on the ground in a field are more active at night than during the day.
3 The invertebrate community living in a field of long grass is different from the community living
in a field of crop plants.
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Practical 18.3
Investigating the distribution of plants along an environmental
gradient, using a transect
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
20 m measuring tape (or length suitable for area of study)
point quadrat
plant keys or identification sheets
Introduction
In this practical, you will:
• use point quadrats to sample plant species across a terrestrial area.
A transect is a systematic sampling method, used to study the distribution of organisms across an
environmental gradient, where there is a change from one habitat to another.
An example of an environmental gradient is the change from a fully terrestrial habitat, such as a field,
to an aquatic one, such as a pond. Here the habitat gradually changes, passing through an intermediate
marshy area. Another example is across a path, where the habitat changes from grass to a trampled
area and back to grass again. A very clear environmental gradient is seen on a rocky seashore, where
the environment changes from marine at the bottom of the shore, to terrestrial at the top. In between
these two extremes, the rocky shore organisms live in habitats with different degrees of immersion.
Procedure
1
Select a suitable area for investigation, and place the measuring tape across the area.
2
Insert the point quadrat in the ground at the start of the tape, at right angles to the tape.
3
Drop a pin through the first hole in the point quadrat. Record any plant species that the pin
touches, using a suitable table. For each pin drop, record only one hit per species, regardless of the
number of times a pin touches that species.
4
Place the pin in the second hole and repeat step 3. Repeat this procedure until all ten holes have
been used. For each species, you will have a maximum of ten hits.
5
Move the quadrat further along the tape, to the next sampling station. (The distance will depend on
the size of the sampling area to be investigated.)
6
Repeat steps 2 to 5 at regular intervals along the tape.
7
Combine your results with those of other groups, in order to obtain a bigger sample size.
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8
Present the combined results graphically, to show the distribution of representative species across
the transect. You can do this as series of histograms, or as kite diagrams. Your teacher will help.
9
Address the following points and questions.
a
Describe any change in the plant community along the environmental gradient.
b Can you explain any trends that are apparent? What biotic or abiotic factors might be
important in affecting the distribution of plants across the transect? (See Figure 18.11 and
pages 431–434 in the Coursebook.)
c
What are the advantages and disadvantages of using a point quadrat, rather than an area
(frame) quadrat, when carrying out a transect?
d What are the limitations of an ecological investigation of this kind? How could you improve
the reliability of your sampling?
e
If time is available, you could measure one of the factors you have identified in b to find out if
there is a correlation between the factor and the distribution of plant species. Correlations can
be tested statistically, using the Spearman rank correlation coefficient (consult your teacher).
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Practical 18.4
Measuring the diversity of broad-leaved plant species in two
habitats, using Simpson’s Index of Diversity
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
two 10 m measuring tapes
(or string with knots at 1 m intervals)
stout pegs (if using string)
0.5 m × 0.5 m (0.25 m2) frame quadrat
•
•
random number table
(or calculator with random number function)
plant keys or identification sheets
Introduction
In this practical, you will:
• investigate the diversity of broad-leaved species in two habitats, using the random sampling
method described in Practical 18.1.
The diversity of plant species in a habitat can be measured using Simpson’s Index of Diversity. The
index requires the total numbers of each species of plant to be counted.
Procedure
1
Mark out an area of ground 10 m × 10 m in the chosen habitat, using measuring tapes or string.
2
Use a random number table or a calculator with a random number function to generate a pair of
random numbers (from 0 to 9) to supply the coordinates for placing the first quadrat.
3
Locate the coordinates along the tapes and place the bottom left corner of the quadrat at the
intersection of the coordinates.
4
Identify the different species of broad-leaved plants within the quadrat, using a plant key or
identification sheets. If there are difficulties with identification, just call them species A, species B
etc. Count the numbers of each species and record your results in a suitable table.
5
Repeat the sampling in a different area, such as a different part of the school field. Try to select
areas with visible differences in vegetation.
6
For each habitat, calculate Simpson’s Index of Diversity (D) from the following formula:
D=1–
n
  N 
2
where N = the total number of broad-leaved plants of all species, and n = the total number of
broad-leaved plants of one particular species.
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n
, square this number, and then sum (Σ) all the squares.
N
Values of D range from 0 to 1, where 0 represents a low diversity, and 1 a high diversity.
For each species, calculate
7
Which habitat has the higher species diversity?
8
Can you think of any reasons why the two habitats might have a different diversity?
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Practical 18.5
Observing single-celled organisms
Safety
Take care when using mains-operated microscopes with water or solutions.
Wash hands after handling biological material.
Apparatus and materials
•
•
•
•
•
•
dropping pipettes
microscope
hand lens
slides
cavity slides
cover slips
•
•
•
•
•
calibrated eyepiece graticule
live culture of Amoeba
live culture of Paramecium
live culture of Chlorella
sample of pond water, or water from an aquarium
Introduction
In this practical, you will:
• observe some species of single-celled organisms that live in freshwater; they are members of the
kingdom Protoctista.
All protoctists have eukaryotic cells. Some, such as Amoeba and Paramecium, have cells that are
similar in structure to cells of animals. These are sometimes called protozoa. Others, such as
Chlorella, have plant-like cells. They belong to a large group of protoctists commonly termed algae.
Multicellular organisms such as animals and plants are made up of many different kinds of cell, each
specialised to perform a particular function. In a single-celled (unicellular) organism, the one cell has
to perform all these different functions. Because of this, the cell of a protoctist often has a complex
internal organisation.
Procedure
A Observing Amoeba
1
An amoeba is a very large cell, just visible to the naked eye or through a hand lens. Observe a
culture of living Amoeba – they should be just visible as tiny grey specks on the bottom of the
culture bottle. Transfer a drop of water containing some cells onto a cavity slide. Place a cover slip
onto the slide.
2
Examine the slide using the low-power and high-power objectives and find a large specimen of
Amoeba. You may need to observe it for a while before it starts to move. When the Amoeba begins
to move, notice the changes in shape as extensions of the cell (pseudopodia) are formed.
3
Note the streaming of the cytoplasm in the direction of formation of the pseudopodia. The inner
cytoplasm is more granular, and called endoplasm. The clear, stiffer outer cytoplasm is called
ectoplasm. The mechanism by which pseudopodia form is still not fully understood.
4
Note the nucleus, and any food vacuoles in the cytoplasm. These vacuoles contain food particles
such as bacteria and small pieces of organic matter that the Amoeba has ingested by phagocytosis.
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You may also be able to see a contractile vacuole. This organelle fills with water and collapses at
intervals, removing water that has entered the cell by osmosis.
5
Make a labelled drawing of an Amoeba. Use a calibrated eyepiece graticule to measure the size of
the cell, and add a scale to the drawing.
B
Observing Paramecium
1
Place a drop of water from a pure culture of Paramecium on a microscope slide. Use forceps to
add a few fibres from a paper tissue or cotton wool. This restricts the movements of the
Paramecium so that they can be seen more easily. Place a cover slip over the specimen and gently
lower it into place.
2
Look at the slide using the low-power and then high-power objectives. Note the Paramecium
moving in the water. It moves quickly, using cilia that cover the surface of the cell. These beat in a
coordinated way, linked by their basal bodies through a network of protein filaments and
microtubules. What happens when the Paramecium hits an obstacle?
3
Observe the nucleus, and the stiff outer covering of the cell, called the pellicle. You may be able to
see the oral groove, where food particles are swept by the beating cilia into the ‘gullet’. If the
organism keeps still for long enough, you may be able to see the two contractile vacuoles filling
and emptying as they remove water from the cell.
4
Make a labelled drawing of a Paramecium. Use a calibrated eyepiece graticule to measure the size
of the cell, and add a scale to the drawing.
C
Observing Chlorella
1
Mount a drop of water from a pure culture of Chlorella on a microscope slide.
2
Look at the slide using the low-power and high-power objectives. This unicellular alga consists of
very small cells packed full of chloroplasts. Some single cells will be visible; others will be in
groups of three or four.
3
Make a labelled drawing of a few Chlorella cells. Use a calibrated eyepiece graticule to measure
the size of the cell, and add a scale to the drawing.
D
Observing single-celled organisms in pond water
1
Mount a drop of pond water on a microscope slide.
2
Look at the slide using the low-power and high-power objectives and see if you can find any other
unicellular organisms. Algal cells like Chlorella should be easy to identify from their green
chloroplasts. You may see Paramecium or other ciliates, or Amoeba. Some motile species use
flagella for movement, rather than cilia. A flagellum has a similar structure to a cilium, but is
much longer. There are often many cilia covering the surface of the cell, but there will be only one
or two flagella.
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Practical 18.6
Using mark–release–recapture to estimate the size of a
population of snails
Safety
Be aware of any possible dangers or biohazards in the fieldwork site. Discuss the risk assessment with
your teacher.
Wear gloves if required.
Wash hands after handling biological material.
Apparatus and materials
•
non-toxic waterproof paint or nail varnish
•
small paintbrush
•
bucket
Introduction
In this practical, you will:
• use the mark–release–recapture method to estimate the size of a population of terrestrial snails.
The mark–release–recapture method can be used to estimate the total population of a mobile animal
living in a well-defined area. A sample of the population is caught and marked in a non-harmful way.
The animals are then released back into their habitat and left to mix with the rest of the population.
After a suitable period of time, a second sample is taken and the numbers of marked and unmarked
individuals is counted. The proportion of marked individuals in the second sample is used to estimate
the size of the total population.
Procedure
1
Select a terrestrial habitat containing a sizeable population of an identifiable species of snail. The
habitat must be a well-defined one, such as a small walled garden.
2
Collect as many snails as possible from the habitat. Mark each snail with a small spot of non-toxic
waterproof paint or nail varnish applied to the shell. Keep the snails in a bucket until the paint has
dried.
3
Count the marked snails and return them to their habitat. Replace them evenly throughout the area.
4
After 3 days, collect a second sample of snails from the habitat. You should collect them at the
same time of day as you collected the first sample.
5
Record the number of marked and unmarked snails in the second sample.
6
Calculate the total population of snails from the formula:
Estimated population size = number in first sample × total number in second sample
number of marked individuals in second sample
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A student carried out this investigation. The results were:
 number of snails in first sample = 63
 number of snails in second sample = 78
 number of marked snails in second sample = 15
Calculate the population size from these data.
8
Why should both samples be collected at the same time of day?
Hint: think about the feeding habits of the snails.
9
Why is this method only suitable for a species living in a well-defined habitat?
10 Explain the importance of the following limitations of the sampling method:
a
the marking technique must not harm the snails or make them more easily seen
b the capture technique must not alter the habitat
c
there must be no immigration or emigration of snails from the habitat.
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Chapter 19 Practical guidance
These practicals are included to give ideas for activities to support teaching of the Cambridge
International AS and A Level Biology syllabus.
The practicals chosen relate closely to the learning outcomes, and may be used to develop students’
practical skills in preparation for practical assessment. However, they are not intended to form a
complete practical course.
Safety
Although great care has been taken in checking the accuracy of the information provided, Cambridge
University Press shall not be responsible for any errors, omissions or inaccuracies.
Teachers and technicians should always follow their school and departmental safety policies. You
must ensure that you consult your employer’s model risk assessments and modify them as appropriate
to meet local circumstances before starting any practical work. Risk assessments will depend on your
own skills and experience, and the facilities available to you. Everyone has a responsibility for his or
her own safety and for the safety of others.
The practicals should be carried out by teachers themselves before they are presented to students.
Additional notes relating to each activity in this chapter are given below, but should not be regarded as
risk assessments.
Activity 19.1
Investigating the primary structure of ribonuclease
Students will require internet access to get the most out of this activity; however, the Coursebook acts
as a useful reference and other reference books may assist.
This activity contains two sections, the first containing a number of worksheet-style questions and the
second making use of an online resource to investigate amino acid sequences.
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