Biology tcm4 123674 (1)

FURTHER PRACTICAL ACTIVITIES (AH BIOLOGY) 1
INTRODUCTION
Revised practicals for mandatory units
This pack contains revised practicals for the mandatory units as detailed
below. These practicals were first published in the HSDU support pack
Biology (Advanced Higher) Practical Activities, 7133, summer 2000.
• Staining a root tip and calculating its mitotic index
The concentration of disodium hydrogen phosphate (0.2 M) has been
added to the instructions in the technical guide for the citrate/
phosphate buffer used to dissolve toluidine blue.
• Gel electrophoresis of DNA treated with restriction enzymes
This practical based on the NCBE Plant DNA Investigation Kit has had
more detail and a new procedure for staining DNA added.
• Isolating and examining cysts of potato cyst nematodes
The text of this practical has had some minor amendments.
New practical for mandatory unit
A new practical for the Cell and Molecular Biology unit, The effect of
competitive and non-competitive inhibitors on the enzyme β-
galactosidase has been included.

FURTHER PRACTICAL ACTIVITIES (AH BIOLOGY)2
INTRODUCTION
Experimental work
One report of an experimental activity is required as evidence for the
assessment of Outcome 3 in each unit. The choice of experiment is not
prescribed in the unit specification and so Centres can select from the
activities included in the support materials, adapt them for individual
use, or use existing activities. The Student Activity Guides provide
guidance on the amount of detail and help students might expect to
receive. The experimental activity must allow for the collection and
analysis of information to meet the performance criteria of Outcome 3.
Outcome 3 performance criteria:
a. The information is collected by active participation in the
experiment.
b. The experimental procedures are described accurately.
c. Relevant measurements and observations are recorded in an
appropriate format.
d. Recorded experimental information is analysed and presented in
an appropriate format.
e. Conclusions drawn are valid.
f. The experimental procedures are evaluated with supporting
argument.
Purpose
A range of practical activities is provided that are suitable for Outcome 3.
The extension work in the teacher/lecturer guide provides ideas that
could be developed into investigations to meet the requirements of the
Biology Investigation unit.
Any hazards associated with the experiments have been identified and
suitable control measures included in the support material as a result of
risk assessment.
Structure
Teacher/lecturer guide
This indicates whether the experimental activity can be used to provide
evidence for Outcome 3 or for other purposes. A section on
background information includes the biology associated with the
experiment where necessary and any prior knowledge or skills students
will require before undertaking the activity. Advice on classroom
management for the teacher/lecturer will include advice on organising

INTRODUCTION
student groups, pooling results, time required and the supply of
materials to students. There will also be advice on possible extension
and follow-up activities that could be developed into ideas for
investigations.
Technical guide
This provides a list of materials required for each activity, including
sources and suppliers for items not generally available from major
suppliers. There is advice on the preparation of materials and risk
assessments. The supply of materials to students should allow for a
degree of planning and organising of experimental work. This does not
mean planning and designing in the sense of an investigation as often
the student will be presented with an experimental procedure. Rather it
should allow the student to plan how he or she will lay out equipment
and materials in preparation for carrying out the experimental activity
and planning the execution of the experimental procedures.
Preparing for the activity
This section is designed to make students think actively about their
experimental work and to plan and organise its execution. To that end
it includes an analysis of the activity which poses questions about the
experimental design. Students, although presented with experimental
procedures to follow, are expected to plan and organise carrying out
the experimental work. In practical terms this will involve reading
through the procedure, identifying and collecting the materials they
require and organising themselves to carry out the procedures and
record results either individually or as a group. For some experimental
activities ‘Preparing for the activity’ has been customised by adding
evaluation questions which will assist students in considering issues
which could be addressed in the experimental report.
This section presents a number of options for teachers and lecturers in
teaching experimental work. Students could be led through the stages
in preparing for the activity by their teacher/lecturer or it could be
presented to students as an individual or group activity. Alternatively
the different stages in preparing for the activity could be presented as a
mixture of these approaches as teachers and lecturers consider
appropriate for their students. Also different experimental activities
may lend themselves to different approaches, or as students’ skills
develop the approach may be changed to suit their experience.
A general section ‘Preparing for the activity’ is included as Appendix 1.
This should be used for each practical activity unless there are
customised questions on evaluation in which case a ‘Preparing for the

INTRODUCTION
activity’ section appears in the support material for that particular
activity.
Student activity guide
This includes an introduction, which provides background information
for the student on the biology of the activity or any other information
required. The experimental procedures for students are described in
the equipment and materials section and the instructions. The
instructions take the students through the steps required for the activity
as well as providing limited advice on the recording, analysis and
presentation of data.
Conditions required for practical work for Outcome 3
Arrangements documentation and Subject Guides refer to assessment
being carried out under controlled conditions to ensure reliability and
credibility. For the purposes of internal assessment, this means that
assessment evidence should be compiled under supervision to ensure
that it is the students’ own work.
It must be emphasised that the assessment for this outcome is not a
special assessment event but part of the ongoing learning and teaching
process. The experimental activity is likely to be performed by a small
group of students together. After collection of the experimental
information each student must complete a report individually under
supervision. A written report should be provided for evidence where
circumstances make that possible. For students with special needs for
whom written evidence is not appropriate alternative forms of report
can be used.
For Outcome 3 there is no specified time limit, but practical constraints,
such as the length of a class period, are likely to play a part. It is
appropriate to support students in producing a report to meet the
performance criteria. Thus redrafting of reports after necessary
supportive criticism is to be encouraged as part of the learning and
teaching process and to produce the evidence for assessment.
Redrafting should focus on the performance criteria concerned and, as a
general rule, should be offered on a maximum of two occasions
following further work by the student on the areas of difficulty.
Report writing
Students should receive an ‘Advice to Candidates’ page (Appendix 2)
which they can refer to during the experiment and the writing of the
report to aid clarity and ensure completeness of their report. This gives

INTRODUCTION
advice on structuring the report under specific headings making a blank
report booklet unnecessary. In some experiments where only one of
the items listed in the conclusion or evaluation is likely to be required
this can be indicated to the students.
Marking reports
The ‘Outcome 3: Teacher/Lecturer Guide’ in Appendix 3 summarises the
performance criteria together with suggested items which might aid the
professional judgement of the assessor. It is important to consider each
individual experiment and how the specific advice given in the Teacher/
lecturer guide for the experimental activity relates to the suggestions to
aid professional judgement. Centres may wish to produce customised
departmental marking schemes for the particular practical activities they
use to provide evidence of Outcome 3. The advice on marking reports
for Outcome 3 at Higher and Int 2 contained in the support material
Marking Advice for Assessing Outcome 3 (Int 2 and H), 5722, published
August 1999, applies equally to Advanced Higher Biology.
The final decision on achievement must be on the basis of the
performance criteria. Although poor grammar, poor sentence
construction and bad spelling would be drawn to the student’s
attention, these aspects are not in any of the performance criteria.
Definitive guidance on the assessment of students’ reports for
Outcome 3 is to be found in National Assessment Bank materials.

CELL AND MOLECULAR BIOLOGY
ACTIVITY 1
Unit: Cell and Molecular Biology (AH): Structure, function and
growth of prokaryotic and eukaryotic cells
Title: Staining a root tip and calculating its mitotic index
Type and purpose of activity
This experiment can be used to:
• provide evidence for the assessment of Outcome 3
• develop knowledge and understanding of the process of mitosis
• develop problem solving skills and in particular Outcome 2
performance criteria:
(b) information is accurately processed, using calculations where
appropriate
(d) experimental procedures are planned, designed and evaluated
appropriately.
Background information
In this activity students will prepare and stain root tips. To achieve an
Outcome 3 students must either have two different sources of root tips
or stain one type of root tip with two different stains. A comparison
between either the root types or the stains will then be possible.
Two recommended sources of roots are garlic and hyacinth. The garlic
cloves, bought normally for cooking purposes, will produce roots at any
time of year. Hyacinth bulbs can be bought at Garden Centres during
autumn and winter. Both garlic cloves and hyacinth bulbs will produce
ample roots for the experiment.
Suitable stains for studying the stages of mitosis in root tips are
lactopropionic orcein and toluidene blue.
The mitotic index is the fraction of cells in a microscope field which
contain condensed chromosomes. This index will be calculated for each
slide prepared.

Preparation of the plant materials and the stains is covered in the
Technical Guide.
To make this activity non-seasonal, it is possible to ‘fix’ the root tips
when available and then store them until required. Fixing of root tips is
only covered in the Technical Guide.
Classroom management
Students are asked to mark the root tip one or two days prior to
staining the root tips. This will enable them to link rate of growth with
mitotic index.
Microscopic examination of the slides:
Students should examine several slides and calculate the mitotic index
for each one. Prepared slides could also be available.
Supply of materials
In order to satisfy the core skill in problem solving, students will be
required to identify and obtain resources required for themselves.
Further advice on supply of material is given in the Technical Guide.
Advice on marking Outcome 3 report
Specific advice for performance criteria b–f
PC b: a description of the preparation of the root tip(s) and the
method(s) of staining should be included.
PC c: drawings or a description of some of the cells showing the
different stages of mitosis; the magnification used should also be
noted.
PC d: a table of results recording:
(i) the number of cells containing condensed chromosomes
in a particular field
(ii) the total number of cells in the field
(iii) the mitotic index for the field.
The results should include at least two different microscope
fields for each situation (i.e. two for each type of root tip or two
for each stain used).

PC e: either a conclusion is made about the rate of mitosis in the
different types of root tips (the higher the mitotic index the
greater the rate of mitosis) OR a conclusion is made about the
efficiency of each stain for detecting condensed chromosomes.
PC f: evaluation points include:
• the length of time the root tips were left in the acid: if too
short a time, maceration will be difficult; if too long a time the
tip will disintegrate when being handled
• the amount of cells unstained due to insufficient time in acid,
poor maceration or poor uptake of stain
• how efficient the stain is, e.g. the lactopropionic orcein
usually gives better definition of chromosomes while the
toluidine blue is stronger in colour
• the condition of the roots and their rate of growth prior to
using them for the experiment.
Extension work
Try to vary the mitotic index of the plant tissue, e.g. cutting the root tips
and keeping them at 0°C for 24 hours may increase the mitotic index.
The experimental method can be varied, e.g. varying the temperature or
concentration of acid; varying the time the root tip is in the acid;
squashing the root tip with a coverslip instead of macerating; varying the
age of the root used; preparing the stains differently (e.g. different
dilutions, different pHs); heating the lactopropionic orcein slide gently;
investigating a possible link between rate of growth of root and mitotic
index.
Acknowledgements
Information and advice from Dr Kwiton Jong, Royal Botanic Garden,
Edinburgh, is gratefully acknowledged. Information was also received
from Ashby Merson-Davies, Sevenoaks School, Kent.
This experiment was produced by the SAPS Biotechnology Scotland
Project. Funding for the project was provided by SAPS, Unilever and
The Scottish Office. Support was also provided by Edinburgh University,
Quest International, the Scottish CCC, the Higher Still Development
Unit and the Scottish Schools Equipment Research Centre (SSERC).

FURTHER PRACTICAL ACTIVITIES (AH BIOLOGY)1 0
Technical guide
The class will be varying either plant material or stain for this activity.
The list of materials required will vary depending on this decision.
Materials required
Materials required by each student/group:
gloves and eye protection
compound microscope (×100 – ×400 magnification)
small beaker of 1 M hydrochloric acid (2 will be required if plant material
is being investigated)
small beaker of water and dropper
microscope slides
coverslips
fine forceps
dissecting needle
scissors
soft tissue paper
ruler
fine thread
dropping bottle of lactopropionic orcein and/or (see below) dropping
bottle of toluidine blue
garlic clove with suitable roots and/or (see below) hyacinth bulb with
suitable roots
Materials to be shared:
waterbath at 60°C
marker pen
timer
dropping bottle of 50% glycerol
dropping bottle of 70% ethanol
lens tissue
Preparation of materials
If plant material is to be varied prepare both plant types below. If stain
is to be varied prepare just any one of the plant types.
To prepare hyacinth bulb roots: Place the bulb in a suitably sized
container with water so that the root end is just in contact with the
water. It is best to change the water daily if possible. Roots of a suitable
length (2–6 cm) will be available within a week and perhaps sooner.

FURTHER PRACTICAL ACTIVITIES (AH BIOLOGY) 1 1
Hyacinth bulbs can cause allergies. Wear gloves if handling the bulbs
regularly.
To prepare garlic clove roots: Carefully peel the clove and place it in a
suitably sized container with water, e.g. test tube/boiling tube so that the
root end is just immersed in the water. It is best to change the water
every 2–3 days. Roots of a suitable length (2–6 cm) will be available after
2–4 days).
If stain is to be varied prepare both stains, as detailed next. If plant
material is to be varied prepare just any one of the stains.
Wear gloves and eye protection when handling the stains.
Lactopropionic orcein should be prepared in a fume cupboard or well-
ventilated room. Dilute it to a 45% solution by volume with distilled
water.
Toluidine blue is harmful if swallowed. Prepare a 0.5% solution in a
citrate/phosphate buffer at pH4 (20 cm3
0.1 M citric acid + 10 cm3
0.2 M
disodium hydrogen phosphate + 8 cm3
distilled water).
Fixing the roots
This stage is required only if suitable roots are available but they are to
be stained at a later date.
Mix 6 cm3
absolute alcohol with 2 cm3
glacial acetic acid in a fume
cupboard. This mixture is called Farmer’s fluid and must be freshly
prepared. Once added to the Farmer’s fluid, the root tips can be stored
for many months in a refrigerator.
Supply of materials
It is not appropriate to provide all equipment and materials in, for
example, a tray system for each student/group. Equipment and
materials should be supplied in a way that students have to identify and
obtain resources. Normal laboratory apparatus should not be made
available in kits but should generally be available in the laboratory. Trays
could be provided containing one type of specialist equipment or
materials.

Read through the Student Activity Guide and consider the following
questions.
Analysis of activity
What is the aim of the activity?
Do you know if you are using two types of roots OR two types of stain?
What measurements are you going to make?
What safety measures are you required to take?
As a class, decide what a ‘nucleus’ should look like for it to be composed
of condensed chromosomes.
In your group, decide how the activity will be managed by allocating
tasks to each member. For Outcome 3 it is important that you play an
active part in setting up the experiment and in collecting results.
Recording of data
Prepare a table to record your results. You should use a ruler and
appropriate headings.
Evaluation
If varying plant material, was rate of growth of the two roots similar? If
not, is there a link between mitotic index and rate of growth?
If varying stain, was there a difference in the ability of the root cells to
absorb the stains? Were they absorbed too much/insufficiently?
Does the mitotic index vary much between different results? Account
for these differences, if possible.
Was the treatment in acid (step 4) sufficient to allow for both easy
handling of the root tip and easy maceration? (Insufficient acid
treatment results in easy handling but difficult maceration; too severe
acid treatment results in difficult handling but easy maceration.)

Introduction
You are going to stain root tips and examine them for signs of cells
dividing by mitosis. The chromosomes inside the nuclei of such cells
condense and become visible. You should know what condensed
chromosomes look like and how they move about inside a cell when
undergoing mitosis.
Equipment and materials
gloves and eye protection
compound microscope (×100 – ×400 magnification)
small beaker of 1 M hydrochloric acid (2 will be required if plant material
is being investigated)
small beaker of water and dropper
microscope slides
coverslips
fine forceps
dissecting needle
scissors
soft tissue paper
ruler
fine thread
dropping bottle of lactopropionic orcein and/or (see below) dropping
bottle of toluidine blue
garlic clove with suitable roots and/or (see below) hyacinth bulb with
suitable roots
waterbath at 60°C
marker pen
timer
dropping bottle of 50% glycerol
dropping bottle of 70% ethanol
lens tissue
Wear gloves and eye protection whilst carrying out this experiment.
Avoid skin contact with the stain(s) and avoid breathing in the fumes of
the stain, lactopropionic orcein, if used.

Instructions
Either two types of roots or two different stains will have been
prepared. Find out what is available.
1. One or two days before staining the root tips, remove the plant
material carefully from the water and blot dry gently. Use a
permanent marker pen to mark a small dot about 2 mm from the
end of some root tips. Replace the plant carefully in the water.
2. After one to two days, remove the plant material and use the
thread and ruler to measure how much the root tips have grown
since marked.
3. Preheat about 10 cm3
of 1 M hydrochloric acid in a small beaker to
60°C using a waterbath. Meanwhile, use a lens tissue and alcohol
to clean microscope slides and coverslips.
4. Using scissors, remove the last 2 mm from several young vigorously
growing root tips. Place them in the preheated acid and return to
the waterbath for 4–5 minutes.
5. Gently transfer each root tip to a clean microscope slide containing
a large drop of water.
6. Gently blot dry with a piece of soft tissue.
7. Using a dissection needle, thoroughly macerate the root tip and
spread over an area equivalent to the size of a 5p coin.
8. You are now ready to apply the stain.
If using toluidine blue – Add one drop to the macerated root tip
and immediately cover with a coverslip, invert the slide and blot
firmly several times on a wad of tissues.
If using lactopropionic orcein – Add one drop to the macerated
root tip and leave for 3–4 minutes. To speed up absorption of the
stain, warm the slide gently by holding it 30–40cm above a yellow
Bunsen flame (if your hand becomes uncomfortable you are
heating the slide too much). Cover with a coverslip, invert the
slide and blot firmly several times on a wad of tissues.

9. View under a microscope, ×40 – ×100 magnification initially. Scan
the slide to locate the region of mitosis.
10. View this area at a higher magnification (×400 should be sufficient)
and count:
(i) the total number of cells in the microscope field
(ii) the number of cells with condensed chromosomes which are
going through any of the four stages of mitosis. You will have to
decide where your cut-off point is when considering if cells in
prophase and telophase contain condensed chromosomes (consult
textbooks).
11. Repeat steps 9 and 10 for the various microscope slides prepared.
If you want to prevent the slides from drying out, mount them in
50% glycerol.
12. Calculate the mitotic index for each slide examined (the mitotic
index is the fraction or percentage of cells containing condensed
chromosomes).
13. Draw a table with suitable headings summarising your results.
14. Compare your results with other groups.

ACTIVITY 4
Unit: Cell and Molecular Biology (AH): Applications of DNA
technology
Title: Gel electrophoresis of DNA treated with restriction
enzymes
• develop knowledge and understanding of cutting DNA with
restriction enzymes
(c) conclusions drawn are valid and explanations given are
supported by evidence
appropriately.
This experiment is done with the help of the Plant DNA Investigation kit
obtained from the National Centre for Biotechnology Education
(NCBE), University of Reading, Whiteknights, PO Box 228, Reading RG6
6AJ. Tel: 0118 987 3743 Fax: 0118 975 0140. Cost £130.00 (2000
prices). SAPS offers sponsorship towards the initial cost of a kit
providing that a teacher from the school has attended a SAPS DNA
workshop. Contact SAPS at Edinburgh University (tel: 0131 650 7124) or
at Head Office (tel: 01223 507168) to obtain the appropriate form.
Refills and individual items can also be obtained from NCBE. Student
Guides and a Technical Guide are supplied with the kit and these supply
a great deal of relevant background information.
In this experiment it is assumed that a 4-tooth gel comb is used to
provide 4 wells in each gel. If using a 6-tooth gel comb the wells hold
less DNA.

In the experiment, a batch of DNA is digested by two different
restriction enzymes. Due to inappropriate buffer concentrations, the
activity of the second enzyme will be reduced. However, evidence of its
activity should still be apparent.
Following these instructions, this experiment requires three separate
days to be completed.
Day 1 – Practising with the microsyringe and digesting the DNA requires
30–40 minutes followed by a 40 minute incubation at 37°C. After the
incubation the small tubes should be stored in the freezer until the next
day.
Day 2 – Separating the DNA fragments requires about 30 minutes to set
up. Ensure the gels are loaded close to the electricity supply so they do
not have to be moved once loaded. As long as the electric current has
been applied long enough for the DNA to have moved out of the wells
(40–50 minutes at the lowest voltage) the electricity can be switched off
and on as required (however, when switched off, loading dye will diffuse
out of the gel making it difficult to see how far the DNA fragments have
travelled).
Day 3 – Staining the gels requires only 5–10 minutes but the gel can take
another 15–20 minutes to identify any visible bands and measure the
distance each band has travelled.
The following table is a guide to suitable power supplies, voltages and
total lengths of time to apply voltage to obtain good separation of DNA
fragments.
Students hands should be dry when carrying out the electophoresis.
Type of power supply Maximum safe voltage Time to run
battery (4 × 9 V) 36 V about 2 hours
*regulated power pack 30 V about 2.5 hours
unregulated power pack 16 V 5–6 hours
*regulated power packs can be identified either by labels on the
apparatus or from their accompanying technical information.
Supply of materials

PC b: an outline of procedure being carried out each day, e.g. Day 1 –
each restriction enzyme cutting up the DNA at specific points;
Day 2 – the electricity causing the DNA fragments to migrate
through the gel, the rate of movement being linked to the size
of the fragment; Day 3 – the DNA is stained and the number of
base pairs in any visible band identified by referring to the table
supplied.
PC c: a table of results with appropriate headings and units showing
the size of each visible DNA fragment and the distance it has
travelled.
PC d: a graph of the results. It is probably best with the size of DNA
fragment (number of base pairs) on the x-axis and the distance
travelled (mm) on the y-axis. (If the log of the size of DNA
fragment is plotted against the distance travelled a straight line
should be formed.)
PC e: a conclusion stating that the smaller the DNA fragment the
further it will travel; however, the relationship is not linear, e.g.
a small fragment half the size of another fragment will travel
more than twice the distance of the larger fragment.
(Alternatively, the conclusion could state that there is a linear
relationship between the log of the size of DNA fragment and
the distances moved through the gel.)
• was the DNA mixed enough each time it was transferred? If
too much DNA is in a well ‘streaking’ of the bands will occur;
too little DNA in a well will result in faint bands.
• was the electricity switched on the correct length of time and
an appropriate voltage used? DNA bands should be spaced
out over the entire gel; appropriate voltage is 1–5 volts per
centimetre (the distance between the two electrodes).
• corrosion may occur at the anode; despite this, the
electrophoresis should not be affected.
• if the gel is blank then either the DNA has not been
adequately rehydrated or the stain has not been left in
contact with the gel for long enough.

• why are the smaller DNA fragments not visible? What size
must the fragments in your gel be before they are visible?
• why have some fragments not separated sufficiently to be
seen as separate bands?
References
Investigating Plant DNA – Student Guide and Technical Guide. These
booklets accompany the DNA kit available from NCBE.
Micklos, D. and Freyer, G. (1990), DNA Science. A first course in
recombinant DNA technology, Cold Spring Harbour Laboratory Press/
Carolina Biological Supply Company.
Miller, M. B. (1993), ‘DNA technology in schools: a straightforward
approach’, Biotechnology Education, 4(1), 15–21.
Miller, M. B. (1994), ‘Practical DNA technology in school’, Journal of
Biological Education, 28(3) 203–211.
Miller, M. B. and Russell, G. A. (1996), ‘Practical DNA technology in
school – 2: Computer analysis of bacteriophage lambda base sequence’,
Journal of Biological Education, 30(3) 176–183.
http://www.ncbe.reading.ac.uk
Acknowledgement
Unit and SSERC.

Technical guide
Materials required
Day 1 – 2 pink tubes containing the restriction enzyme EcoR1
2 green tubes containing the restriction enzyme Hind111
1 yellow tube (empty)
1 white tube of DNA suspension
1 microsyringe and 6 tips
1 float
1 vial of loading dye
1 piece of parafilm
1 marker pen
Day 2 – electrical supply (see Teacher/Lecturer Guide)
2 electric wires with crocodile clips
enzyme tubes in the float from the previous lesson
vial of loading dye
gel in a plastic tank with comb, covered in buffer solution
microsyringe and 4 tips
piece of black card
2 pieces of carbon fibre tissue
Day 3 – tank containing your gel from previous lesson
stain (10 cm3
)
gloves
eye protection
Day 1 – waterbath at 37°C
Day 2 – bottle of TBE buffer
Preparation of materials supplied by the kit
Rehydrating the DNA – The λ DNA in the narrow white tubes provided in
the Plant DNA kit must be rehydrated with distilled water shortly before
the experiment is carried out. Follow the instructions on page 10 of the
Student Guide provided with the kit. One tube of DNA is required per
group of students.
Preparing the agarose gel – If necessary, this can be done a few days
before the experiment is carried out. Follow the instructions on page
12 of the Student Guide. One gel is required per group of students.

Two pieces of carbon fibre electrode tissue (approximately 42 mm ×
22 mm) are required per group. Wear gloves when handling the carbon
fibre tissue.
Dilute 1 volume of the TBE buffer concentrate with 9 volumes of
distilled water. About 35 cm3
will be required per group (11–12 cm3
to
dissolve the agarose and form the gel and the rest to cover the gel once
it is set). The liquid can be reused for 3–4 ‘runs’ after which it should be
discarded.
Dilute the concentrated stain for DNA with an equal volume of distilled
water. About 10 cm3
of stain is required per group. This diluted stain
can also be reused several times. Wear gloves and eye protection when
handling the stain.
Recipes for the various buffers and dyes used in the experiment are
given in the Technical Guide supplied with the kit.
Preparation of materials not supplied by the kit
Making a float – Make 4–5 holes in a plastic petri dish lid or base using a
small hot rod. The holes should be about 8 mm in diameter. This will
allow the pointed end of the enzyme microtubes through but will hold
their top end. Alternatively, the holes can be made in a thin piece of
foam such as a camping mat.
Pieces of Parafilm (about 5 cm × 5 cm) are required for the microsyringe
exercise. However, any non-absorbent paper such as benchcoat will be
suitable.
9 volt PP3 batteries can be obtained very reasonably (70p each – 2000
prices) from Middlesex University Services Ltd, (Teaching Resources),
Trent Park, Bramley Road, London N14 4YZ.
Tel: 0208 4470342 Fax: 0208 447 0340.
Supply of materials
materials.

Disposal of materials
All microtubes and gels can be safely disposed of in the bin. Buffer,
loading dye and stain can be diluted and washed down the drain. A
fuller account of safety is covered in the Technical Guide accompanying
the kit.

questions.
Are you familiar with how the restriction enzymes act on DNA?
Are you aware of what is happening during electrophoresis?
Getting organised for experimental work
Are you familiar with the microsyringe and how to deliver a set volume
using it?
Recording of data
Prepare a table with suitable headings and units to record the number of
base pairs in each identified DNA fragment and the distance it has
travelled through the gel.
Evaluation
Why are some DNA fragments not visible?
Why have some DNA fragments not separated sufficiently to be seen as
separate bands?
Is there evidence that the DNA was not evenly distributed in its original
tube? What can be done to prevent this?
How long should the electric current be passed through the gel so that
DNA bands will be separated as much as possible?
Can you account for some lanes of the gel being blank?

Introduction
This experiment uses most of the basic techniques involved in genetic
fingerprinting. The DNA is digested or ‘cut up’ using restriction
enzymes. The resulting fragments of DNA are then separated into bands
using an electric current and made visible by staining.
DNA source DNA fragments DNA fragments
of varying size separated and stained
If the order of bases in the DNA used is different each time then the
DNA fragments produced each time after digestion will also be different.
Thus, DNA from different organisms (except clones) will give a unique
result in this experiment – hence the term genetic fingerprinting.
DNA from a certain bacteriophage will be used in this experiment as
only one, short chromosome is present in the organism. This will result
in only a few different fragments being formed, thus making their
separation into distinct bands more likely.
Nuclear DNA from animals or plants consists of many large
chromosomes. After digestion, a very large number of fragments are
formed. If all these fragments were stained, a smear would result. To
obtain distinct bands (a fingerprint) with this complex DNA, only certain
fragments are selected using probes.
The simple, bacteriophage DNA is going to be digested in 3 different
ways:
– by mixing one sample of DNA with a restriction enzyme called EcoRI
– by mixing another sample of DNA with a different restriction enzyme
called HindIII
– by mixing a third sample of DNA with both of these enzymes.
DNA cut with
restriction
enzymes
electric
current

Each restriction enzyme will cut the DNA only when a certain sequence
of bases occurs, e.g. the enzyme EcoR1 cuts the DNA between bases G
and A only when the sequence GAATTC is present in the DNA. The
other restriction enzyme used cuts the DNA at a different sequence of
bases. Thus, each restriction enzyme is specific.
restriction enzyme EcoRI
The number of DNA fragments formed after digestion by an enzyme will
depend on the number of times the particular sequence of bases which
the enzyme acts on is present, e.g. the sequence GAATTC occurs 5 times
in the bacteriophage DNA used in this experiment. The DNA will
therefore be cut into six fragments when digested by the enzyme EcoRI.
Day 1 – 2 pink tubes containing the restriction enzyme EcoRI
2 green tubes containing the restriction enzyme HindIII
1 yellow tube (empty)
1 white tube of DNA suspension
1 microsyringe and 6 tips
1 float
1 vial of loading dye
1 piece of Parafilm
1 marker pen
Day 2 – electrical supply
2 electric wires with crocodile clips
enzyme tubes in the float from the previous lesson
DNA
double
helix
DNA cut
into
fragments
G
C
T
A G C T T A A
C G A A T T
G
C G A C
C T G
G
C
T
A G C T T A A
C G A A T T
G
C G A C
C T G

vial of loading dye
gel in a plastic tank with comb, covered in buffer solution
microsyringe and 4 tips
piece of black card
2 pieces of carbon fibre tissue
Day 3 - tank containing your gel from previous lesson
stain (10 cm3
)
gloves
eye protection
Day 1 – waterbath at 37°C
Day 2 – bottle of TBE buffer
Instructions
Preliminary exercise
This experiment requires you to transfer very small volumes
of liquids. A microsyringe is provided for you to do this. The
tips that fit on the end of the microsyringe have small ‘ridges’
on them. When the tip is filled to the upper ridge 10 µl will
be delivered. The lower ridge is for delivering volumes of
2 µl.
Follow the hints below when using a microsyringe.
• Before loading the microsyringe, pull the plunger out a little. This
gives some extra air with which to expel the last drop of liquid.
• When emptying the microsyringe tip, hold it vertically and at eye
level.
• To remove the last droplet from the tip, touch it against the inner wall
of the container.
• Do not touch the point of the microsyringe tip with your fingers.
There are enzymes in sweat which may contaminate and result in
unwanted digestion of DNA samples.
• A tip must only be used once to prevent any cross-contamination
occurring.
← 10 µl
← 2 µl

Microsyringe exercise
You may find this useful to become familiar with the microsyringe.
i) Draw in 2 µl of dye and deposit as drop 1 on the Parafilm.
ii) Repeat Step 1 until you have 5 separate drops of dye.
iii) Draw in 10 µl of dye and deposit it alongside the smaller drops.
iv) Now draw all five 2 µl drops into the micropipetter tip and deposit
them alongside the 10 µl drop.
v) Are the two drops the same size?
Day 1 – Digesting the DNA
1. Sit the 4 tubes containing restriction enzymes in the float on the
bench.
2. With a new microsyringe tip draw the DNA suspension into and
out of the microsyringe tip several times. This results in the DNA
being evenly distributed. Now transfer 20 µl of DNA to each of the
two pink tubes containing a restriction enzyme.
3. Again with a new tip, transfer 20 µl of DNA to one green tube
containing a different restriction enzyme. Remember to mix the
DNA thoroughly before transferring it.
4. Again with a new tip, transfer 20 µl of DNA to an empty yellow
tube. This tube will act as a control as here the DNA will be
undigested.
5. Cap the tubes and flick the sides of the tubes with one finger until
the blue colour is evenly spread throughout the liquid.
6. Place the float with the 4 tubes in a
waterbath at 37°C (leaving the one
remaining green tube on your bench).
7. After 10 minutes the restriction
enzymes will be in solution. This will
allow you to transfer the entire
contents of one of the pink tubes to

the remaining green tube again using a new tip on the
microsyringe.
The DNA in this green tube will now be digested by both
restriction enzymes. Mark the tube with a D – for double digest.
8. Flick each tube several times to mix the contents. Put the four
tubes (one pink, one unmarked green, one green marked D and
one yellow) in the float back into the waterbath to incubate at 37°C
for at least another 30–40 minutes.
N.B. The tubes can be left until next lesson as the restriction
enzymes will become denatured after a few hours. To prevent
further DNA breakdown, the tubes should be stored in a freezer
overnight.
Day 2 – Separating the DNA fragments
1. If not already done, cover the gel with about 20 cm3
of buffer
solution (to a depth similar to that shown in the diagram below).
Buffer solutions keep the pH stable and thus prevent unwanted
breakdown of unstable molecules such as DNA.
2. Remove the comb gently from the gel to expose the wells.
3. Ensure your tank is close to your electricity supply and place a
piece of black card under it to make the wells more visible.
*4. Using a new tip, draw in 2 µl of loading dye and mix this
thoroughly with the undigested DNA in the yellow tube by drawing
the mixture up and down in the tip several times.
*5. Draw up all the contents of the tube into the microsyringe tip and
load well 1 by emptying the syringe slowly when the end of the tip
is in the buffer solution and directly above the well.
N.B. The tip does not actually need to be in the well as the dense
dye will make the DNA solution sink.
loading dye
and DNA
buffer solution
gel
microsyringe tip

6. Repeat the last two steps marked * and load each well as follows,
using a new microsyringe tip each time:
Well 2 – DNA digested by restriction enzyme EcoRI (pink tube)
Well 3 – DNA digested by restriction enzyme HindIII (green tube)
Well 4 – DNA digested by both restriction enzymes (green tube D)
7. Put a piece of carbon fibre tissue at either end of the tank.
8. Connect the carbon tissue to the electricity supply using wires and
crocodile clips. Once the electricity is switched on the negatively
charged phosphates in the DNA are attracted to the positive
electrode. So, make sure the positive electrode is furthest away
from the DNA in the wells.
9. Check with your teacher what voltage you will be using and set up
the electricity supply accordingly. Switch on the electricity. The
TBE buffer can evaporate during electrophoresis, so periodically
check the depth of the buffer and top up as required (to a depth
similar to that shown in the diagram in Step 5).
As well as helping the DNA sink into the wells, the loading dye also
allows us to judge how long the electric current should be on by
moving in front of all but the smallest DNA fragments.
10. After an appropriate time (e.g. 12 hours at 9 volts; 6 hours at 18
volts) switch off the electricity, disconnect the crocodile clips and
remove the pieces of carbon fibre.
carbon fibre
wells
buffer solution

Day 3 – Staining the DNA
1. Return the buffer solution covering the gel to its original
container.
2. Pour about 10 cm3
of staining solution (Azure A) onto the surface
of the gel and leave it for at least 4 minutes.
3. Pour off the stain into a bottle labelled ‘reused stain’.
4. Wash excess stain from the surface of the gel with tap water.
5. Do not leave any water on the gel after rinsing. If you do the stain
will move out of the gel into the water.
If the staining solution has been used on a previous occasion you may
need to repeat the above procedure. If this is necessary allow at least 10
minutes for instruction 2.
Purple bands of stained DNA will appear shortly. The smaller the
fragments of DNA the further it will have travelled through the gel.
However, the smallest fragments will also take up less stain and may
therefore be difficult to see. Also, fragments of similar size will move
similar distances in the gel, resulting in little separation between them.
On the next page is a table showing the number and size of DNA
fragments formed during the experiment. This is possible as the entire
base sequence of the DNA in the bacteriophage used has been worked
out.
Lanes
1 2 3 4
DNA
bands
largest
smallest
loading dye
wells

Lane 1 Lane 2 Lane 3 Lane 4
Contents Undigested DNA digested DNA digested DNA digested
DNA by restriction by restriction by both
enzyme, enzyme, restriction
EcoRI HindIII enzymes
No. of DNA
fragments formed 1 6 8 13
No. of log of 48,502 4.685 21,226 4.327 23,130 4.364 21,226 4.327
base fragment 7,421 3.870 9,416 3.974 5,148 3.712
pairs in size 5,804 3.764 6,557 3.817 4,973 3.697
each 5,643 3.752 4,361 3.640 4,268 3.630
fragment 4,878 3.688 2,322 3.366 3,530 3.548
3,530 3.548 2,027 3.307 2,027 3.307
564 2.751 1,904 3.280
125 2.097 1,584 3.200
1,375 3.138
947 2.976
831 2.920
564 2.751
125 2.097
6. Examine your gel and try to connect the DNA fragments listed
above with the bands that have appeared in each lane. For each
identifiable band measure the distance it has travelled. Measure
from the bottom of each well to the front end of each band.
7. Make a table with appropriate headings and units showing the
number of base pairs, the log of the fragment size and the distance
travelled for each band.
8. Present your results as a graph with suitable scales and axes
labelled with quantities and units (put fragment size or log of
fragment size on the x-axis and distance travelled on the y-axis).

Unit: Cell and Molecular Biology (AH): Molecular interactions in
cell events: Catalysis
Title: The effect of competitive and non-competitive inhibitors
on the enzyme β-galactosidase
• develop knowledge and understanding of the effect of competitive
and non-competitive inhibitors on enzyme activity
(c) conclusions drawn are valid and explanations given are
supported by evidence
appropriately.
The enzyme β-galactosidase catalyses the following reaction:
LACTOSE GLUCOSE + GALACTOSE
The chemical ONPG (o-nitrophenyl β-D-galactopyranoside) is also
degraded by the enzyme:
ONPG ONP + GALACTOSE
The ONP produced is yellow, allowing the rate of this reaction to be
followed colorimetrically.
Galactose acts as a competitive inhibitor, competing with ONPG for the
active site of the enzyme. At a sufficiently high concentration, it will
inhibit the reaction by preventing ONPG making contact with the active
ACTIVITY 8
β-galactosidase
β-galactosidase

site. The enzyme, however, is still capable of activity. Thus, when the
ONPG concentration is increased it will eventually overcome the
inhibition.
Iodine solution on the other hand is a non-competitive inhibitor. When
it combines with the enzyme the shape of the active site is altered
sufficiently to prevent the substrate combining with it. Increasing
substrate concentration will therefore not overcome the inhibition.
Students can work individually or in pairs for this experiment.
If there are several groups of pupils requiring to use the colorimeter, a
rotation system could perhaps be employed, i.e. each group could start
the reaction (by adding the enzyme) 20–30 seconds apart. The
colorimeter would just require to be zeroed once for each ‘run’. In this
way 4–6 groups could carry out the experiment at about the same time.
Estimated time: 50–60 minutes should be sufficient to collect all the
data.
The enzyme solution must be kept in crushed ice. If allowed to reach
room temperature its activity will rapidly decrease.
Supply of materials
Specific advice for peformance criteria b–f
PC b: to include a description of the contents of the various cuvettes
set up; preparation of the enzyme solution.
PC c: a table of results for each inhibitor with appropriate headings
(volume of stock ONPG solution present (cm3
) and absorbance/
transmission after two minutes); a table of results using the ×20
diluted ONPG without inhibitor at the beginning and end of the
experiment.

PC d: results for each inhibitor are graphed with volume of stock
ONPG added on the x-axis and absorption/transmission after two
minutes on the y-axis; an appropriate scale is used and axes are
labelled with units; the points are correctly plotted and lines of
best fit are drawn.
PC e: a conclusion is made as to the type of inhibitor galactose and
iodine solutions are.
• evidence that the activity of the enzyme has remained about
constant throughout the duration of the experiment
• suitable precautions have been taken to prevent cross-
contamination
• the importance of keeping the concentration of each
inhibitor constant while increasing the ONPG concentration
• the suitability of the concentration of inhibitor used (did it
inhibit the ×20 diluted ONPG completely?) and the range of
ONPG concentrations used (did enzyme activity recover to its
initial level when ONPG concentration was high?)
• why it is more difficult to obtain complete inhibition with
galactose than with iodine solution.
Extension work
Substitute galactose for glucose (the other product of the reaction) to
see if it has a similar effect on enzyme activity.
Investigate the rates of reaction in the above experiment by regularly
measuring absorbance/transmission over 5–6 minutes.
Investigate the nature of the inhibition using the enzyme phosphatase
and the inhibitors phosphate and iodine.
The rate of reaction (V0
) at low substrate concentrations can be
calculated. If 1/V0
is plotted against 1/[substrate] then the maximum
velocity and the Michaelis constant for the reaction can be calculated.
See Hames reference on enzyme kinetics (or any good biochemistry
textbook).

References
Adds, Larkcom and Miller (eds.), (1996) Cell Biology and Genetics,
Nelson Advanced Modular Science.
Hames, B.D., Hooper, N.M. and Houghton, J.D. (1997), Instant Notes in
Biochemistry, Bios Scientific.
Russo, S. E. and Moothart, L. (1986), ‘Kinetic study of the enzyme
lactase’, Journal of Chemical Education, 63(3), 242–243.

Technical guide
Materials required
6 cuvettes (or test tubes if suitable colorimeter is used)
2 boiling tubes
beaker of crushed ice
6 × 1 cm3
droppers
10 cm3
syringe
6 cm3
ONPG stock solution (3 × 10-2
M in buffer)
40 cm3
buffer (0.1 M potassium phosphate, pH 8)
15 cm3
20% galactose in buffer
5 cm3
I2
/KI solution in buffer
25 cm3
distilled water
eye protection
gloves
colorimeter (420–440 nm filter)
1 cm3
dropper
distilled water
β-galactosidase stock solution
The buffer: 0.1 M K2
HPO4
adjusted to pH 8 with 0.5 M HCl. Each
student/group will require 80–100 cm3
. About half the volume made up
will remain as plain buffer. The rest will be used to make up other
solutions. Avoid direct skin and eye contact, wear eye protection and
gloves.
ONPG stock solution: 3 × 10-2
M in buffer. Each student/group will
require 6 cm3
. For every 10 cm3
required, weigh out 0.09 g and dissolve
in 10 cm3
buffer. Shaking for 5–10 minutes will be required for the
powder to be completely dissolved. The ONPG stock solution is best
made up fresh (or no more than 2 days in advance and stored in the
fridge). ONPG available from Sigma Aldrich, Fancy Road, Poole, Dorset
BH12 4QH. Catalogue no. N1127, 1 g for £9.70 (1999 prices).
Galactose solution: 20% in buffer. Each student/group will require
10–15 cm3
. To make up 50 cm3
, dissolve 10 g galactose in 50 cm3
buffer.
It dissolves readily.

I2
/KI solution: Each student/group will require about 5 cm3
. Dissolve
0.3 g iodine and 1.5 g potassium iodide in 100 cm3
water to make a stock
solution (this will keep for months stored in a dark glass bottle). Take
1 cm3
of this stock solution and make up to 80 cm3
with buffer. This
diluted I2
/KI solution is the solution to be used by the students in the
experiment.
Iodine is classified as harmful. Wear gloves when preparing the
solution.
N.B. The diluted I2
/KI solution must be made up immediately before
the experiment is carried out (it will remain effective as an inhibitor for
1 hour).
β-galactosidase is available as ‘Lactozym’ from NCBE, University of
Reading, Whiteknights, PO Box 228, Reading RG6 6AJ. Tel: 0118 987
3743. Fax: 0118 975 0140. Cost £12.50 (2000 prices) for 100 cm3
.
Avoid direct skin and eye contact, wear eye protection and gloves.
Enzyme powder can cause allergies. Do not allow any spillages to dry
up. Wipe up spillages immediately and rinse cloth thoroughly with
water.
For guidance on sources of colorimeters see SSERC Bulletin No. 198,
Winter 1999/2000, pages 20–27.
Supply of materials
materials.

questions.
What is being varied in the activity?
What variables must be kept constant?
Why should the enzyme activity be measured without either inhibitor
both at the beginning and at the end of the experiment?
In your group decide how the activity will be managed by allocating
active part in setting up the experiment and in collecting results.
Recording of data
Prepare tables to record your group results.
You should use a ruler, correct headings and appropriate units.
Evaluation
Has the activity of the enzyme remained about constant for the duration
of the experiment?
Cross-contamination will seriously affect the results. Have sufficient
measures been taken to avoid cross-contamination?
Why is it more difficult to completely inhibit the enzyme with galactose
than with iodine solution?
Is the range of ONPG concentrations used suitable to show clearly if the
inhibitor is competitive or non-competitive?

Introduction
Inhibitors are substances that reduce the activity of enzymes.
When the inhibitor binds reversibly to the active site of the enzyme it is
known as a competitive inhibitor. Often a competitive inhibitor is a
similar shape to the substrate. Its association with the active site of the
enzyme reduces the rate of binding between the substrate and the
enzyme, thus lowering the rate of reaction. However, this type of
inhibition can be overcome by increasing the substrate concentration as
this will decrease the chances of enzyme and inhibitor binding.
When a non-competitive inhibitor combines with an enzyme, the
active site may still be free. When it combines with the enzyme the shape
of the active site is altered sufficiently to prevent the substrate
combining with it. Increasing substrate concentration will therefore not
overcome the inhibition.
In this experiment you will use the enzyme β-galactosidase. Its normal
substrate is lactose but you will use a synthetic substrate, ONPG. When
the enzyme is active, it breaks down the ONPG to a yellow substance.
Thus, the rate of reaction is proportional to the intensity of the yellow
colour formed.
ONPG YELLOW SUBSTANCE + GALACTOSE
(ONP)
The reaction will firstly be carried out without an inhibitor, using a low
concentration of substrate. An inhibitor will then be used at a
concentration that prevents this enzyme/substrate mixture from
reacting. While keeping the inhibitor concentration constant, the
substrate concentration will be gradually increased. If the inhibition is
overcome by this action, the inhibitor is competitive. If the inhibition is
unaffected, the inhibitor is non-competitive.
β-galactosidase

6 cuvettes (or test tubes if suitable colorimeter is used)
2 boiling tubes
beaker of crushed ice
6 × 1 cm3
droppers
10 cm3
syringe
6 cm3
ONPG stock solution (3 × 10-2
M in buffer)
40 cm3
buffer (0.1 M potassium phosphate, pH 8)
15 cm3
20% galactose in buffer
5 cm3
I2
/KI solution in buffer
25 cm3
distilled water
eye protection
gloves
colorimeter (420–440 nm filter)
1 cm3
dropper
distilled water
β-galactosidase stock solution
Instructions
Wear eye protection and gloves throughout this experiment to avoid
direct skin and eye contact with some of the chemicals used.
1. Put 20 cm3
of distilled water in a boiling tube. Surround the tube
with crushed ice and add 4 drops of β-galactosidase.
This is the enzyme solution you will use throughout the
experiment. Do not allow it to reach room temperature as this
will reduce the enzyme’s activity considerably. Ensure the stock
β-galactosidase is returned to the refrigerator as soon as possible.
Enzyme powder can cause allergies. Do not allow any spillages to
dry up. Wipe up spillages immediately and rinse cloth thoroughly
with water.
2. Mix 0.5 cm3
of the stock ONPG solution with 9.5 cm3
of 0.1M buffer
(pH 8). Label ×20 dilution.
3. Put 2 cm3
of buffer and 1 cm3
of this ×20 diluted ONPG solution
into a cuvette. Mix by inverting the cuvette 2–3 times. Zero the
colorimeter with this solution.

4. Add 0.5 cm3
of the diluted enzyme to the cuvette. Start the
stopclock and invert the cuvette 2–3 times.
5. Record the absorbance/transmission two minutes after adding the
enzyme. This should be between 0.3 and 0.5 absorbance units
(50–32% transmission).
If the absorbance is above 0.5 units, dilute the enzyme solution
with distilled water and repeat steps 2–4 until an appropriate
absorbance is obtained after 2 minutes. If the absorbance is below
0.3 units, add 1–2 drops of the stock β-galactosidase to your
diluted enzyme.
You are now going to investigate:
(i) the effect of galactose (an inhibitor) on the activity of the enzyme
(ii) the effect of increasing the ONPG concentration (the substrate) in
the presence of galactose.
6. Mix the solutions, as shown in the following table, in different
cuvettes.
cuvette no. 20% galactose ONPG stock buffer (cm3
) *ONPG ×20
in buffer (cm3
) solution (cm3
) dilution
(cm3
)
1 2 - - 1.0
2 2 0.25 0.75 -
3 2 0.5 0.5 -
4 2 0.75 0.25 -
5 2 1.0 0 -
* Note: the volume of ONPG stock solution in the ×20 dilution is 0.05 cm3
7. Treat each cuvette in turn as follows:
Invert 2–3 times, put in colorimeter and zero the instrument.
(Care! If you are sharing the colorimeter with other groups, only
the first group should zero it for each ‘run’.)
Add 0.5 cm3
of the diluted enzyme solution. Start the stopclock
and invert cuvette 2–3 times.
Take an absorbance/transmission reading 2 minutes after adding
the enzyme. Record your results in a table with suitable headings.
Rinse out the cuvettes several times with water and dry.

You are now going to investigate:
(i) the effect of iodine solution (another inhibitor) on the activity of
the enzyme
(ii) the effect of increasing the ONPG concentration in the presence of
the iodine solution.
Care! Iodine is harmful. Wear gloves and eye protection.
8. Again, using the following table as a guide, mix the solutions in
different cuvettes.
cuvette no. I2
KI solution ONPG stock buffer (cm3
) *ONPG ×20
(cm3
) solution (cm3
) dilution
(cm3
)
1 1.0 - 1.0 1.0
2 1.0 0.5 1.5 -
3 1.0 1.0 1.0 -
9. Treat each cuvette in turn as follows:
Invert 2–3 times, put in colorimeter and zero the instrument.
(Care! If you are sharing the colorimeter with other groups, only
the first group should zero it for each ‘run’.)
Add 0.5 cm3
of the diluted enzyme. Start the stopclock and invert
cuvette 2–3 times.
Take an absorbance/transmission reading 2 minutes after adding
the enzyme. Record your results in a table with suitable headings.
Rinse out the cuvettes several times with water and dry.
10. To ensure that enzyme activity has remained constant, repeat steps
3–5. These results should be similar to the ones obtained initially.
11. Present your results for both investigations as a graph with suitable
scales and axes labelled with quantities and units.

ENVIRONMENTAL BIOLOGY
ACTIVITY 5
Unit: Environmental Biology (AH): Symbiotic relationships
(Parasitism)
Title: Isolating and examining cysts of potato cyst nematodes
• develop knowledge and understanding of parasitism and more
specifically of the relationship between potato cyst nematodes (PCN)
and potato plants
(b) information is accurately processed using calculations where
appropriate
appropriately.
An outline of the life cycle, transmission and control of the potato cyst
nematode (PCN) is covered in the Student Activity Guide.
This is a good example of parasitism to study as:
(i) it affects a common and economically important food crop
(ii) cysts containing the parasite remain viable for many years and can
be collected and examined at any time of year
(iii) controlling PCN is expensive, complicated and an ever increasing
problem.
There are two species of PCN; Globodera rostochiensis and Globodera
pallida. Although both are troublesome, G. pallida is the more serious
pest and becoming increasingly difficult to control. Some varieties of
potato are resistant to G. rostochiensis. A few varieties are partially

resistant to G. pallida. Varieties susceptible to both are: Arran Comet,
Desiree, Estima, King Edward, Maris Bard, Maris Peer, Pentland Dell,
Record, Wilja, Golden Wonder and Kerr’s Pink. Resistant varieties to G.
rostochiensis include: Cara and Maris Piper. Nadine and Sante are
resistant to G. rostochiensis and partially resistant to G. pallida.
Obtaining suitable soil samples is covered in the Technical Guide. The
initial extraction of PCN using sieves should take only 15–20 minutes.
However, filtering the water/soil mixture must be completed before
proceeding to the next stage of the experiment. The filtering will take
about 30 minutes and, of course, longer if the water/soil mixture is
filtered a second time.
Ideally the moist filter papers should be kept overnight in a humid
environment. The cysts will then burst more readily. However, it is
possible to complete the entire experiment on the same day if necessary,
although cyst bursting may be less successful.
Examination of the cysts will take 30–60 minutes. The filter papers are
first examined under a low power binocular microscope (×10 – ×20).
Cysts are transferred to a microscope slide and then burst whilst viewing
under a compound microscope (×100). Identifying PCN cysts and
distinguishing between viable and non-viable PCN is covered in the
Student Activity Guide.
N.B. PCN are a serious pest of a common food crop and as such
are subject to statutory control measures to limit their spread and
population increase. It is therefore essential that good laboratory
practice is followed at all times during this procedure. This
includes autoclaving all possible sources of viable cysts once the
experiment is completed. All possible precautions should also be
followed to prevent soil infected with viable cysts from being
washed down the sink, especially if sludge from local sewage
treatment plants is spread on agricultural soil. Care must also be
taken to avoid cross-contamination of samples.
Supply of materials
In order to satisfy the core skill in problem solving, students will
be required to identify and obtain resources required for
themselves. Further advice on supply of material is given in the
Technical Guide.

PC b: a description of the method used to extract PCN from a soil
sample; a description of a viable and non-viable PCN.
PC c: a table with suitable headings showing the total number of cysts
per 100 g of at least two soil samples.
PC d: a table with suitable headings showing the percentage of viable
cysts in at least two soil samples.
PC e: a conclusion on how suitable each soil would be for producing a
crop of seed potatoes.
• possible ways of losing PCN cysts during the extraction
method
• the possibility of mistaking a viable PCN for a non-viable one
• the reliability of the method used in taking the soil sample
from a field.
Extension work
Make exudates from resistant and non-resistant potatoes. Mix these
with viable cysts and note any differences in number of PCN released
from cysts. A method for making exudate and inducing hatching of
cysts is included in the Technical Guide.
As above but vary the exudate, e.g. temperature of mixing, previously
boiled, vary pH and concentration.
Examine a variety of soils for PCN.
Test the efficiency of the extraction method by adding a known
number of cysts to a soil sample, follow the method given and
calculate the percentage recovered. The extraction method can be
varied and the percentage of cysts recovered monitored.

References
Atkinson H. (1997), ‘The worm in the root!’, Biological Sciences
Review, 9(5), 36–38.
Council Directive of 8 December 1969 on control of potato cyst
eelworm (69/465/EEC), Official Journal of the European Communities
Number L323/3 24/12/69.
Evans K. A., Harling R. and Dubickas A. (1998), ‘Application of a PCR-
based technique to speciate potato cyst nematodes and determine the
distribution of Globodera pallida in ware growing areas’, Aspects of
Applied Biology, 52, 345–350.
Evans F. and Haydock P. (1999), ‘Control of plant parasitic nematodes’,
Pesticide Outlook, 10(3), 89–128.
Marks R. J. and Brodie B. B. (Editors), Potato Cyst Nematodes – Biology,
Distribution and Control.
Acknowledgements
The original protocol for this experiment was obtained from the Scottish
Agricultural College (SAC), West Mains Road, Edinburgh. This
information and advice from A. Evans and C. Kasperak of SAC are
gratefully acknowledged.
Information and advice were also obtained from D. Trudgill and A. Holt,
Scottish Crop Research Institute (SCRI), Invergowrie.
Acknowledgements also to J. Pickup, Scottish Agricultural Science
Agency (SASA), East Craigs, Edinburgh.
Unit and SSERC.

Technical guide
Materials required
large filter paper (185 mm diameter)
set of compasses with pencil
ruler
filter funnel (top internal diameter about 100 mm)
washing bottle
glass rod
large beaker, e.g. 400 cm3
binocular microscope (×10 – ×20)
compound microscope (×100)
piece of acetate
large conical flask, e.g. 250 cm3
pair of fine forceps
microscope slides
coverslips
dried soil, gently crushed or rolled
balance
weighing boats
soil sieves with large mesh (550 µm – 850 µm) – mesh no. 30 or 20
soil sieves with small mesh (250 µm) – mesh no. 60
Obtaining a suitable soil sample containing viable PCN may present a
problem in some areas. A garden or allotment with a history of growing
susceptible varieties of potatoes (see Teacher/Lecturer Guide) is usually
a good source. In rural areas a local farmer may be willing to provide
suitable soil.
If taking soil samples from any land you must ensure that all equipment
used and boots worn are clean and could not be contaminated with
cysts from a prior sampling site. The distribution of cysts is unlikely to
be uniform. ‘Hot spots’ will occur and so it is important to take several
samples of about 100 g at intervals throughout the field. Sampling
points should be chosen randomly and small soil samples lifted using a
trowel or the widest cork borer (no. 6 – each bore will give about a 10 g
sample).
SAPS may be able to supply a limited number of non-viable cysts.

Soil samples should be dried at room temperature before use. This
increases the chances of PCN cysts floating during their extraction from
soil. If the soil is not fine, it may also need to be passed through a
riddle or lumps broken up gently.
To make exudate:
1. Grow susceptible potato in sand (or sandy soil) for 2–3 weeks.
2. Collect and wash roots.
3. Cover roots with water and leave for 4–6 hours or overnight in a
refrigerator.
4. Filter and collect exudate.
To induce hatching of cysts:
1. Put about 10 cysts in water for 5–7 days.
2. Remove all the water and cover with exudate.
3. Cysts will start to hatch within 5 days. Remove a few drops of
exudate to a dimpled microscope slide to view nematodes.
N.B. New cysts may need to be stored at 4°C for 3–6 months before they
will hatch.
Supply of materials
materials.
Disposal of materials
It is most important that good laboratory practice is carried out during
this experiment. All materials containing cysts must be autoclaved or
soaked in bleach before being disposed. Suitable precautions are listed
in the Student Activity Guide.

questions.
What measurement are you going to take?
Are you aware of the size of potato cyst nematode cysts and what they
look like?
Are you aware of the differences between viable and non-viable potato
cyst nematodes?
Are you aware of the precautions you must follow to prevent further
spread of this parasite?
active part in carrying out the experiment and in collecting results.
Recording of data
Prepare a table to record:
(i) the total number of cysts in each soil sample
(ii) the percentage of viable cysts.
You should use a ruler, correct headings and appropriate units when
necessary.
Evaluation
Are there possible flaws in the extraction process where PCN can be lost
from the sample, leading to unreliable results?

Do you think the procedure involved in taking the soil sample is
reliable? Is the sample size (50 g) large enough? (A 500 g sample is used
when this procedure is carried out professionally.)
Has the filter paper been examined sufficiently or is it possible that cysts
on it could be overlooked?

Introduction
Potato cyst nematodes (PCN), also known as potato cyst eelworms
(PCE), are world-wide parasites of potato plants. They originated in
South America where the Incas practised a seven-course rotation to
control them. Being parasites the PCN receive all their nutritional
requirements from the potato plant, resulting in reduced root and foliar
growth and a reduction in tuber yield. The cost of damage caused by
PCN is estimated to be about £43 million each year in the UK alone
(1990–1995). This annual cost is increasing as is the incidence of PCN.
Like many parasites, PCN have a highly specialised life cycle. The cysts
you are going to isolate are only about 0.5 mm in diameter and may
contain up to 200–600 eggs initially which have larvae coiled up inside
them.
0.5 mm
0.5 mm
every year a small number
of eggs are released
spontaneously. This
number increases when a
susceptible potato variety
is grown in infected soil.
Female becomes attached to
potato plant. When fertilised
by male its body swells
and develops into a cyst.
Larva emerges from egg, invades
root and establishes a feeding site. If
no host plant is available, the larva
dies within days.
Cyst - light brown in colour.
Contains 200-600 eggs.
Can remain dormant in soil
for up to 30 years.
Egg containing
coiled up larva.
Cyst – light brown in colour.
Contains 200–600 eggs. Can
remain dormant in soil for up
to 30 years.
Every year a small number
of eggs are released
spontaneously. This
number increases when a
susceptible potato variety
is grown in infected soil.

Infection of potato plants by PCN has several effects:
(i) Even moderately low population densities (about 5 eggs per gram
of soil) will reduce yields and high populations (200–2000 eggs per
gram of soil) may result in complete crop loss.
(ii) As a result of infection, plants have a stunted root system making
them more susceptible to drought.
(iii) Secondary invaders, e.g. fungi, can enter the root system more
readily.
The main means of passive transmission of PCN are through the planting
of infected potatoes, i.e. potatoes grown on infected land, and by the
movement of contaminated soil, e.g. that adhering to farm machinery.
They are mainly controlled by using a combination of the following:
(i) Crop rotation (long rotations allow natural population decline)
(ii) Use of resistant varieties which inhibit PCN multiplication
(iii) A type of pesticide known as nematicides (affect the nervous
system of juveniles which prevents juveniles locating a host plant).
large filter paper (185 mm diameter)
set of compasses with pencil
ruler
filter funnel (top internal diameter about 100 mm)
washing bottle
glass rod
large beaker, e.g. 400 cm3
binocular microscope (×10 – ×20)
compound microscope (×100)
piece of acetate
large conical flask, e.g. 250 cm3
pair of fine forceps
microscope slides
coverslips
dried soil, gently crushed or rolled
balance
weighing boats
soil sieves with large mesh (550 µm – 850 µm) – mesh no. 30 or 20

soil sieves with small mesh (250 µm) – mesh no. 60
detergent with dropper
Precautions required to be taken
As potato cyst nematodes are a serious pest to an economically
important food crop, the precautions listed below must be followed.
1. If taking soil samples from any land you must ensure that all
equipment used and boots worn are clean and could not be
contaminated with cysts from a prior sampling site.
2. Find out if sludge from your local sewage treatment plant is spread
on agricultural soil. If so, all possible precautions should be
followed to prevent viable cysts from being washed down the sink.
3. After use, all apparatus such as sieves and glassware should be
autoclaved or soaked in bleach overnight before being washed.
Such treatment will kill viable cysts.
4. Wipe up spillages with a paper towel and place in a bin.
5. Care must be taken to avoid cross-contamination of samples.
Instructions
N.B. For successful extractions, cysts must be clean and previously
dried in the soil at room temperature.
1. Weigh out 50 g of the dried soil. The soil sample has a history of
being used for growing potatoes. Break up any small lumps gently
with the end of a glass rod.
2. Collect the two soil sieves, fitting the one with the larger mesh size
on top. Place the sieves above a bucket or polythene bag and add
the soil sample to the top sieve.
3. Sift the dry soil for 3–4 minutes.
4. Wash the sieves under a fast running tap. Cysts will not pass
through the finer sieve so it can be washed on its own under the
tap. When washing the larger mesh sieve always place the finer
mesh sieve beneath it.

In the instructions that follow, treat the contents of each sieve
separately. Each group of students should therefore form two
smaller groups, one working with the soil in the large mesh sieve,
the other with the soil in the small mesh sieve.
5. Away from the sink, wash out
the contents of your allocated
sieve into a beaker with the help
of a wash bottle.
To do this, hold the sieve almost
at right angles above the beaker
and with a wash bottle project a
stream of water on to what was
the lower side of the sieve.
Slowly rotate the sieve while
doing this. Then, turning it the
right way up, wash final
contents from the sieve. Do
not now wash sieves in the sink
– see precautions.
6. Allow the soil/water mixture to settle until little movement of
material is occurring (10 minutes).
7. Meanwhile, using a pair of
compasses and a pencil, draw four
concentric circles on a large filter
paper (as shown in the diagram).
Ensure the circles drawn are
complete and prominent. Draw a
straight line from the centre to the
edge of the filter paper.
8. Fold this filter paper twice and fit it
into a filter funnel. Sit the funnel
on top of a large conical flask.
9. Once the contents of the beaker have settled, decant quickly into
the filter paper without disturbing the sunken soil. While
decanting, rotate the beaker slowly so that any floating debris stuck
to the sides gets washed into the filter paper.
wash
bottle
sieve
soil sample
185 mm
15 mm

10. Add a drop of detergent to the soil/water mixture while it is
filtering. This encourages any cysts present to migrate to the sides
and stick to the paper.
11. Using a high pressure flow of water, add about 200 cm3
to the
beaker containing the soil. Allow to settle and decant as before
into the filter paper.
12. Once filtration is complete remove the filter paper from the
funnel, unfold it and place overnight in a humid, airtight container.
This ensures that the cysts will burst easily.
13. On the next day, place the filter paper on a suitable surface (e.g. a
piece of acetate) and examine under the binocular microscope.
Starting at the straight line in the outermost circle, examine this
circle for cysts. Repeat this procedure for the other circles on the
filter paper.
Potato cyst nematode cysts are only 0.5 mm in diameter
on average. However, they are easily detected by their
shape and colour – perfectly spherical apart from a
small ‘neck’ (rather like a gourd or a spherical
decoration commonly put on a Christmas tree). They
vary from being orange and copper coloured to a dull dark brown.
Warning: Other cysts may be present, e.g. cereal cyst nematode
(these are lemon shaped).
14. With a pair of fine forceps remove any cysts from the filter paper
and place in a droplet of water on a microscope slide. The
concentric circles drawn previously should help to ensure the
entire filter paper is scanned although most cysts should be found
in the outermost circle. Count the total number of cysts found on
the filter paper. Add this to the number found on the filter paper
from the other sieve of the same soil sample.
15. Select at random several cysts and place them far enough apart on
a few microscope slides so that each can be covered by a separate
coverslip. Add a drop of water to each cyst and cover each one
with a coverslip.
16. Examine each cyst in turn under a microscope (×100 total
magnification). Whilst viewing a cyst press down gently on the
coverslip. This will cause the cyst to burst and release its contents.
Look in particular at any larvae whose egg case has burst. If the

egg case does not burst you will see capsule-shaped objects as in
the diagram of the life cycle. Determine the number of cysts
containing viable larvae.
N.B. Do not attempt to burst open all the eggs. A cyst just needs to
contain ONE viable larvae for it to be scored as viable. If cysts are
completely empty, assume they are non-viable.
17. Calculate:
(i) the total number of cysts per 100 g of your soil sample (you
started with a 50 g sample).
(ii) the percentage of viable cysts in your random sample of cysts.
18. Compare the soil sample you have just examined with one with a
different history for growing potatoes.
19. Present your results in a table with suitable headings. Draw a bar
chart with the axes labelled appropriately to show the results
graphically.
The experiment you have just done is a simplified, scaled-down version
of a test carried out routinely on fields intended for the production of
seed potatoes. If even one viable potato cyst nematode is found in a
500 g sample then the field cannot be used to provide seed potatoes.
N.B. 1. This experiment is done for educational purposes only and
should not be used as a basis for any agronomic decisions
due to the relative inexperience of the testers.
2. Soil and any equipment used in the experiment must now be
autoclaved to kill any PCN cysts. Do not dispose of any soil
samples by returning them to land from which they did not
originate.
Viable larvae will uncoil completely
when the egg case bursts. Their
‘skin’ will be smooth and free of
any sudden indentations. Non-viable larvae will have folds
and ‘kinks’ in their ‘skin’.

BIOTECHNOLOGY
ACTIVITY 9
Unit: Biotechnology (AH): Use of Micro-organisms: Stages of
growth
Title: Growth curve: Determination of doubling time and growth
rate constant
• develop knowledge and understanding of the stages of growth of
microbes in culture, turbidometric measurement of cell growth and
growth rate constants
(a) relevant information is selected and presented in the
appropriate format
(b) information is accurately processed using calculations where
appropriate.
In industry, it is important to be able to determine the growth rate of a
given micro-organism and understand the factors that affect it in order
to generate maximum product by the most economic means. The
product may be a metabolite produced at a given stage of the growth
cycle or it may be the organism itself, e.g. the production of yeast
biomass to be used as starter cultures for brewing or baking, or as the
starting point for autolysis which produces a huge variety of food
flavourings.
As the number of cells in a microbial culture increases, the turbidity
(cloudiness) of the culture increases. Turbidity is caused by the
suspended cells scattering light and it may be measured using a
colorimeter. Absorbance increases as the cell concentration increases

BIOTECHNOLOGY
giving a convenient, rapid and accurate method of measuring cell
growth rates.
This method cannot, however, distinguish between live and dead cells.
A growth curve generated by this method over the time suggested will
not demonstrate the death or senescent phase. Viable counts would
have to be carried out.
Absorbance is plotted on a graph against time. Doubling of absorbance
indicates doubling of the number of cells and the time taken for this to
occur can be read from the graph.
In this experiment students will create a growth curve of absorbance
versus time, then use it to calculate doubling time and growth rate
constant using absorbance as the measure of growth.
Obtaining results for a growth curve cannot be managed in one lesson.
This practical has been designed with the aim of ease of collection of
data and production of a classic growth curve shape showing lag phase,
exponential phase, stationary phase and eventually death phase if the
culture is left long enough or viable counts are measured.
Medium is inoculated with a very small quantity of yeast late in the
afternoon then samples are taken three times per school day for the
next three or four days (early morning, lunchtime and late afternoon).
Timing is not critical but time of sampling should be recorded so that
hours of incubation can be calculated.
Samples do not have to be read immediately – they can be placed in
sterile Bijoux bottles, tubes or universals, refrigerated and the
absorbance read when the time is convenient, although preferably
within 24 hours. The yeast cells will settle so it is very important to
shake gently to suspend the cells before reading the absorbance.
Students working as part of a group could arrange a rota for removal of
samples.
A number of factors are important.
• Very small inoculum – to allow good demonstration of lag phase.
• Timing – lag phase has been best observed by inoculating the
medium late in the afternoon.

BIOTECHNOLOGY
• Yeast type – do not use fast acting yeasts as they show a very short lag
phase, if any.
• Cultures must be gently agitated before removing a sample to ensure
that the cells are in suspension.
• Samples must be well suspended before taking a reading.
Supply of materials
PC b: to include description of how the low inoculum concentration is
achieved; method of sampling; method of measuring
absorbance.
PC c: a table of results with appropriate headings and units showing
the time and date of sampling, hours of growth and absorbance.
PC d: a graph of absorbance on the y-axis and hours on the x-axis. Lag
phase, log phase and time of stationary phase should be
labelled. Indication of measurement of generation (doubling)
time should be made on the graph (i.e. the time taken for the
absorbance to double).
PC e: growth rate constant is calculated using the generation time
determined from the graph.
• accuracy of inoculum concentration
• mixing of culture before removal of samples
• suspension of cells before reading absorbance
• control of temperature
• determination of doubling time from graph
• usefulness of semi-logarithmic graph paper in plotting and/or
analysing results.

BIOTECHNOLOGY
Extension work
The effects on the growth curve and the growth rate constant of varying
the growth media.
The effects on the growth curve and growth rate constant of varying the
incubation temperature.
The effects of different concentrations of starter culture on the length of
the lag and log phases of growth and initiation time of stationary phase.
The effects on the growth curve of keeping the inoculum under
different conditions before inoculation, e.g. in the fridge.
Comparison of growth curves and growth rate constants for different
micro-organisms or types of dried yeast in the same media.
Comparison of different methods of enumerating micro-organisms (e.g.
haemocytometer and viable count) to generate a growth curve.
References
Iain S. Hunter (2000), Biology: Biotechnology Student Monograph
(Advanced Higher), Learning and Teaching Scotland
Acknowledgment
Quest International, Learning and Teaching Scotland, the Higher Still
Development Unit and SSERC.

BIOTECHNOLOGY
Technical guide
Materials required
5 cm3
sterile yeast glucose broth as blank
99 cm3
sterile yeast glucose broth in flask
dried yeast (not fast acting)
weighing boat
spatula
10 cm3
sterile water (if balance is accurate to 0.01 g)
100 cm3
sterile water
sterile 1 cm3
pipette
discard jar containing 2% stericol
semi-log graph paper
waterbath or incubator at 30°C
balance (accurate to 0.001 g preferably, or 0.01 g)
colorimeter (440 nm)
Yeast glucose broth (for 1 litre medium)
20 g glucose
20 g bactopeptone
10 g yeast extract
0.1 M sulphuric acid or 0.5 M sodium hydroxide
distilled water
Instructions
1. Wear a lab coat.
2. Weigh glucose, bactopeptone and yeast extract into a beaker.
3. Add distilled water to 1 litre mark.
4. Stir thoroughly and adjust to pH 6.
5. Dispense volumes into required containers for autoclaving.
6. Autoclave for time and temperature appropriate to medium.

BIOTECHNOLOGY
Notes
• Medium for blanks can be kept in sterile Bijoux bottles or small sterile
test tubes (plugged or covered) and can be refrigerated and used
over the four days taken to generate the growth curve.
• Media should be made up, dispensed into Bijoux bottles, test tubes or
flasks (covered or plugged) then sterilised immediately by
autoclaving.
• Tins of traditional dried yeast are better as sachets of yeast tend to be
of the fast-acting variety and do not demonstrate lag phase so well.
• When samples have been read, the yeast suspension should be
disposed of into a discard jar containing 2% stericol and the cuvette
washed with detergent and hot water.
• Digital colorimeters, e.g. WPA CO75 or Harris S-Range colorimeter,
are best used for this experiment. Older colorimeters may not be
sensitive enough.
Supply of materials
materials.

BIOTECHNOLOGY
questions.
How will you record these measurements?
How will you determine the information you require to make the final
calculation?
What constant will you calculate?
tasks to each member. It is very important that samples are removed at
least three times per day.
Recording of data
Prepare tables and semi-logarithmic graph paper to record your group
results.
You should use a ruler, correct headings and appropriate units.
Evaluation
How effective were the methods which you used?
What were the limitations of the equipment?
What were the sources of error?
What possible improvements could be made to the experiment?
What is the benefit of plotting results on semi-logarithmic graph paper?
What is the economic importance of the process which you are studying
and the calculations which you will make?

BIOTECHNOLOGY
Introduction
Growth is the process during which living organisms synthesise new
chemical components for the cell and as a result they usually increase in
size. In unicellular organisms, such as bacteria and yeast, growth leads
to cell division and consequently an increase in population size. The
growth of a population of single-celled micro-organisms grown in a
closed environment typically shows four stages: lag phase; exponential
phase; stationary phase; death phase.
The lengths and characteristics of these phases will depend upon
factors such as the nature of the growth medium and temperature of
incubation.
In industry, it is important to understand the factors which affect the
growth rate of a given micro-organism in order to generate maximum
product by the most economic means. For example, if the desired
product is a secondary metabolite such as an antibiotic which is
produced when the organism has stopped growing, the manufacturer
will want to provide optimum conditions for the culture to reach
maximum numbers in stationary phase in the shortest time possible.
In some cases, the product is the organism itself, e.g. the production of
yeast biomass to be used as starter cultures for brewing or baking, or as
the starting point for autolysis which produces a huge variety of food
flavourings.
Growth of a population can be measured using the following methods:
Cell counts: total numbers of cells are counted directly using a
microscope and a special slide called a haemocytometer.
Dilution plating: the culture is serially diluted and a known volume of
each dilution plated out and incubated. Resulting colonies are counted
giving a measure of viable numbers of cells in the original population.
Turbidometric methods: Cell density is measured using a colorimeter.
This is a photometric method which measures the light scattered by the
cells in suspension. Increase in cell density is an extremely accurate
method of measuring cell growth rates.

BIOTECHNOLOGY
In this practical, you will produce a growth curve of absorbance against
time for a culture inoculated with a known dry mass of Saccharomyces
cerevisiae (bakers’ yeast) then grown over several days. From this you
will be able to calculate generation time and a growth rate constant.
5 cm3
sterile yeast glucose broth as blank
99 cm3
sterile yeast glucose broth in flask
dried yeast (not fast acting)
weighing boat
spatula
10 cm3
sterile water (if balance is accurate to 0.01 g)
100 cm3
sterile water
sterile 1 cm3
pipette
discard jar containing 2% stericol
semi-log graph paper
water bath or incubator at 30°C
balance (accurate to 0.001 g preferably, or 0.01 g)
colorimeter (440 nm)
Instructions
1. Start this experiment late afternoon at the start of a week.
2. Using aseptic technique, add 0.025 g dried yeast to 100 cm3
sterile
distilled water at 30°C. Shake gently to ensure that the cells are
evenly distributed and suspended.
(Note: if your balance is not sensitive enough to measure out such
a small quantity, add 0.25 g yeast to 10 cm3
sterile distilled water,
mix well then aseptically withdraw 1 cm3
and add to 99 cm3
sterile
distilled water.)
3. Using aseptic technique, dilute 100 times by adding 1 cm3
to
99 cm3
sterile broth in a flask. This should give a starting
concentration of 0.0025 g/l for your growth curve.
4. Using sterile medium as the reference, calibrate the colorimeter
(i.e. set it to zero).

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