EXPERIMENTS
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SEPARATING MICRO-ORGANISMS
Source: NQ Curriculum Support Intermediate 2 Biotechnology
(Unit 2 Student Materials)
When professional microbiologists isolate micro-organisms from the
environment or an infected person, it is extremely rare to obtain a pure
culture. It is therefore necessary to separate micro-organisms. Plating or
streaking can be used to achieve this. Using isolated single colonies as
inocula for further streak plates, pure cultures can be obtained.
Streaking out a mixed broth culture on an agar plate and incubating it to
obtain single colonies of different types of bacteria or yeasts can simulate this.
Streak plate showing individual colonies
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Instructions
These instructions are for right-handed people. If you are left handed, please
reverse the instructions accordingly.
1 Wear a lab coat.
2 Prepare your work space on the bench, collect the materials and set them
out correctly on the bench.
3 Label the bases of the Petri dishes containing the appropriate sterile agar
with initials, date and name of culture.
4 Using the mixed broth culture as the inoculum, streak out applying the
method you have used previously.
5 Incubate the plates at room temperature for 48 hours.
If you have obtained well isolated single colonies, use these as inocula for
further streak plates using the method you have used previously to obtain
pure cultures.
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CONJUGATION IN MUCOR (ZYGOSPORE PRODUCTION)
Source: James Watt College

Label an agar plate as shown here:
+
-

Sterilise a platinum stab wire or a scalpel blade.

Transfer a piece of mycelium from a Mucor (+) strain to one half of the
agar plate.

Resterilise the wire/blade.

Transfer a piece of mycelium from a Mucor (-) strain to the other half of the
agar plate

Use sellotape tabs to secure lid to base of plate

Carefully invert the plate and store at 25C for 3 – 7 days.

Note the growth which is found after this period of incubation.
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EFFECTS OF TEMPERATURE ON MICROBIAL GROWTH
Source: James Watt College

Stock cultures (24 hours) of the following microbes:
Bacillus subtilus
Escherichia coli
Bacillus stearothermophilus
Saccharomyces cerevisiae

5 nutrient agar plates.

Divide each plate into four quadrants and label appropriately with microbes
name or initials.

Inoculate each quadrant with a different microbe by means of a single line
of inoculation of each microbe in its appropriately labelled section.

Secure lids to plates.

Incubate in inverted position at appropriate temperature (10C, 20C,
37C, 50C and 60C) for 24 hours.

Record amount of growth:
(-)
(1+)
(2+)
(3+)

=
=
=
=
Absence of growth
Scant growth
Moderate growth
Abundant growth
Comment on the temperature ranges the 4 microbes grow best at.
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ANTIBIOTIC SENSITIVITY: KIRBY-BAUER METHOD
Source: James Watt College
This technique is used to determine the sensitivity of various microbes to
antibiotics present on discs.

Inoculate agar plates with 0.1 ml of test microbe:
E. coli
Klebsiella pneumoniae
Staphylococcus epidermis albus
i.e. set up spread plates

Allow culture to soak in for 10 minutes.

Add antibiotic disks using sterile forceps.
Do not move the disk once it is placed on the agar

Secure lids to plates.

Incubate at 30C for 24 hours. Do not invert.

Take a note of the specific antibiotics used here.

Determine whether the bacteria are resistant, intermediate or susceptible
to each antibiotic by comparing ‘zones of inhibition’ ie areas of clearing
around the antibiotic discs.

Deduce which antibiotic(s) would be best to treat each individual species
of bacteria.
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BIOCHEMICAL TESTS
Source: James Watt College
Oxidase Test
This identifies cytochrome c oxidase, an enzyme found in obligate aerobic
bacteria. Soak a small piece of filter paper in a fresh solution of 1% (w/v) NN-N1-N1-tetramethyl-p-phenylenediamine dihydrochloride on a microscope
slide. Rub a small amount from the surface of a young, active colony onto the
filter paper using a glass rod or plastic loop: a purple-blue colour within 10 s
is a positive result
Note: Performing the oxidase test – never use a nichrome wire loop, as this
will react with the reagent, giving a false positive result.
To avoid false negatives ensure you use sufficient material during oxidase
and catalase testing, otherwise you may obtain a false negative result: a
clearly visible ‘clump’ of bacteria should be used.
Catalase Test
This identifies catalase, an enzyme found in obligate aerobes and in most
facultative anaerobes, which catalyses the breakdown of hydrogen peroxide
into water and oxygen. Transfer a small sample of your unknown bacterium
onto a coverslip using a disposable plastic loop or glass rod. Invert onto a
drop of hydrogen peroxide on a slide: the appearance of bubbles within 30 s
is a positive reaction. This method minimizes the dangers from aerosols
formed when gas bubbles burst.
The oxidase and catalase tests effectively allow us to sub-divide bacteria on
the basis of their oxygen requirements.



Obligate aerobes will be oxidase and catalase positive;
Facultative anaerobes will be oxidase negative and catalase positive;
Microaerophilic bacteria, aerotolerant anaerobes and strict (obligate)
anaerobes will be oxidase and catalase negative – the latter group will
grow only under anaerobic conditions.
Once you have reached this stage (colony characteristics, shape, Gram
reaction, oxidase and catalase status) it may be possible to make a tentative
identification, at least for certain Gram-positive bacteria, at the generic level.
To identify Gram-negative bacteria further tests are usually required.
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SERIAL DILUTION
Source: HSDU Biology and Biotechnology Microbiological Techniques
Intermediate 1-Advanced Higher Folder
Materials
Lab coat
Eye protection
Benchkote if necessary
Disinfectant and paper towels
Discard jar with disinfectant
Bunsen burner
Labels
7 sterile 1 cm3 syringes or pipette tips
7 sterile test tubes or bottles each
containing 9 cm3 diluent
Broth culture of yeast or bacteria to
be diluted
Instructions
1 Wear a lab coat and use eye protection.
2 Label the test tubes or bottles with the appropriate dilution, 10 -1 – 10-7.
3 Make sure that the lid of the culture tube is firmly attached then shake the
culture vigorously to separate clumps of cells and to distribute the
organisms evenly throughout the liquid.
4 Remove a sterile syringe or pipette tip from its pack/container. Do not
touch the parts which will come in contact with organism. If using a pipette
tip, carefully attach to the dispenser.
5 Using aseptic technique (i.e. flaming the neck of the tube or bottle after
removal and before replacement of its lid), remove exactly 1 cm 3 of the
fluid and transfer to the dilution tube next in the series.
6 Place syringe or pipette tips into discard jar.
7 Mix the dilution well.
8 Repeat steps 4 – 7 until the last dilution tube is reached.
This process dilutes the organisms in the original sample to a countable
number.
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COUNTING YEAST CELLS USING A HAEMOCYTOMETER
Source: HSDU Biology and Biotechnology Microbiological Techniques
Intermediate 1-Advanced Higher Folder
Materials
Haemocytometer and coverslip
Tissue
Alcohol
Lens tissue
Water
Suspension of yeast cells
Capillary tube
Petri dish with moist tissue
Microscope
Instructions – setting up the slide
1 Clean the haemocytometer with alcohol,
then wipe with lens tissue.
2
Using a damp tissue, moisten the slide as shown in diagram.
3 Push the special coverslip on to the slide as shown in diagram, pressing
down on the outside edges of the coverslip at the same time until you can
see Newton’s rings (see diagram). If you push the centre of the
coverslip, it is likely to break.
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Instructions – loading the haemocytometer
4 Shake the cell suspension gently.
5 Insert the end of the capillary tube into the suspension. The liquid will rise
into the tube.
6 Run the end of the capillary tube along the edge of the coverslip between
the arms of the ‘H’. The suspension should fill the area between the
coverslip and the top half of the ‘H’ (shaded in diagram below). If the
suspension flows into the troughs (the lines of the ‘H’), clean the
slide and start again.
7 Turn the slide through 180 and repeat for the opposite edge of the
coverslip.
8 Place the haemocytometer on a damp tissue in a Petri dish for at least two
minutes to equilibrate.
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Instructions – counting the cells
The haemocytometer has two grids situated as shown in the diagram:
1
Place the haemocytometer on the microscope stage
2
Using the instructions for use of the microscope, examine the
haemocytometer using the 4x objective lens. You should be
able to view one whole grid as shown in the diagram.
3 Increase the magnification to the 10x objective lens.
You should be able to see the 25 central squares, each of
which is divided into 16 smaller squares (see diagram at
instruction 4).
4 Increase the magnification to the 40x objective lens.
You will see one of the 25 central square made up of 16
small squares.
5 Count the cells in each of the four corner squares and the central square;
(see shaded squares in instruction 3). Note that you will count five groups
of 16. Include in the count those cells touching the top or right side of the
square; do not count those cells touching the bottom or left side. This
takes account of cells which are half in and half out the square.
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Instructions – calculation
Length of side of grid
= 1 mm
Area of grid
= 1 mm2
Depth between coverslip and slide
= 0.1 mm
Volume under squared area (25
squares) of grid
= 1 mm2 x 0.1 mm = 0.1mm3
Volume under 5 squares (the
number counted)
=
0.1/
5
mm3 = 0.02mm3
You have therefore counted the number of cells in 0.02 mm 3 and can use the
following calculation to estimate the cell concentration of your original
suspension.
Number of cells (total in 5 squares) in 0.02 mm3
= n
Number of cells in 1 mm3
= n x 50
Number of cells in 1 cm3
= (n x 50) x 1000
If the cell suspension counted has been diluted, then the above result must be
multiplied by the appropriate dilution factor to give the concentration of the
original culture.
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VIABLE COUNT: POUR PLATE METHOD
Source: HSDU Biology and Biotechnology Microbiological Techniques
Intermediate 1-Advanced Higher Folder
Materials
Lab coat
Eye protection
Benchkote if necessary
Disinfectant and paper towels
Discard jar with disinfectant
Bunsen burner
Dilution series of organism
Sterile pipettes or syringes (0.1 cm3)
Sterile Petri dishes
20 cm3 volumes of sterile molten
nutrient agar at 45C
Glass spreader
Alcohol
Instructions
1 Wear a lab coat and use eye protection.
2 Label the underside of the plates with initials, date, sample and dilution.
For greatest reliability/precision, each dilution should be plated in triplicate
and the average of the three counts used.
3 Remove a sterile 0.1 cm3 syringe or pipette tip from its pack/container. Do
not touch the parts which will come in contact with organism. If using a
pipette tip, carefully attach to the dispenser.
4 Start with the highest dilution (i.e. 10-7).
5 Using aseptic technique (i.e. flaming the neck of the tube or bottle after
removal and before replacement of its lid), remove exactly 0.1 cm 3 of the
sample and transfer to the base of a sterile Petri dish.
6 Using aseptic technique, pour 20 cm3 sterile nutrient agar over the sample
and mix gently.
7 Place syringe or pipette tip into discard jar.
8 Repeat steps 5 – 7 for dilutions 10-6, 10-5 and 10-4.
9 When the plates are solidified and dry, incubate upside down at the
appropriate temperature for the appropriate time.
10 After incubation, select plates for counting that contain 30 – 300 colonies
(samples which contain <30 colonies/0.1 cm3 diluent are subject to large
fluctuations in numbers or sampling errors, plates which contain >300
colonies are likely to have overlapping colonies).
11 Count accurately and record the number of colonies on each plate.
12 Calculate the concentration of viable cells or colony forming units (cfu) in
the original suspension.
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VIABLE COUNT : SPREAD PLATE METHOD
Source: HSDU Biology and Biotechnology Microbiological Techniques
Intermediate 1-Advanced Higher Folder
Materials
Lab coat
Eye protection
Benchkote if necessary
Disinfectant and paper towels
Discard jar with disinfectant
Bunsen burner
Labels
Dilution scrics of organism
Sterile pipettes or syringes (0.1 cm3)
Sterile nutrient agar plates
Glass spreader
Alcohol
Instructions
1 Wear a lab coat and use eye protection.
2 Label the underside of the plates with initials, date, sample and dilution.
For greatest reliability/precision, each dilution should be plated in triplicate
and the average of the three counts used.
3 Remove a sterile 0.1 cm3 syringe or pipette tip from its pack/container. Do
not touch the parts which will come in contact with organism. If using a
pipette, carefully attach to the dispenser.
4 Start with the highest dilution (i.e. 10-7).
5 Using aseptic technique (i.e. flaming the neck of the tube or bottle after
removal and before replacement of its lid), remove exactly 0.1 cm 3 of the
sample and transfer to the surface of an appropriately labelled sterile
nutrient agar plate.
Note: Keep remainder of all samples
6 Using aseptic technique, spread the sample evenly across the plate with
the glass spreader.
7 Place syringe or pipette tip into discard jar.
8 Repeat steps 5 – 7 for dilutions 10-6, 10-5 and 10-4.
9 When the plates are dry, incubate upside down at the appropriate
temperature for the appropriate time.
10 After incubation, select plates for counting that contain 30 – 300 colonies
(samples which contain <30 colonies/0.1 cm3 diluent are subject to large
fluctuations in numbers or sampling errors, plates which contain >300
colonies are likely to have overlapping colonies).
11 Count accurately and record the number of colonies on each plate.
12 Calculate the concentration of viable cells or colony forming units (cfu) in
the original suspension.
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YEAST GROWTH CURVE
Source: Adapted from SAPS ‘Growth Curve: Determination of Doubling Time’
http://www-saps.plantsci.cam.ac.uk/worksheets/scotland/double.htm
Technical guides are also available from the same source.
Read through the Student Activity Guide and consider the following questions.
Analysis of Activity
What is the aim of the activity?
What measurements are you going to make?
How will you record these measurements?
How will you determine the information you require to make the final
calculation?
What constant will you calculate?
Getting organised for experimental work
In your group decide how the activity will be managed by allocating tasks to
each member. It is very important that samples are removed at least three
times per day: ideally early-morning, lunchtime and late afternoon. This will
happen over 3 days. [A rota for removing samples may help].
Recording of data
Prepare tables and a graph 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 ideas do you have for further work?
What is the economic importance of the process which you are studying and
the calculations which you will make?
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STUDENT ACTIVITY GUIDE
Introduction
Stages of Growth
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.
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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.
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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.
Equipment and materials
Materials required
Materials required by each student/group:
1 x 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 0.01g)
100 cm3 sterile water
sterile 1 cm3 pipette
discard jar containing 1% Virkon
semi-log graph paper
Materials to be shared:
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 Draw a table showing date, time, hours of incubation and absorbance.
3 Using aseptic technique, add 0.025 g dried yeast to 100 cm 3 sterile
distilled water at 30C. Shake gently to ensure that the cells are evenly
distributed and suspended.
4 Using aseptic technique, dilute 100 times by adding 1 cm 3 to 99 cm3 sterile
broth in a flask. This should give a starting concentration of 0.0025 g/l for
your growth curve.
5 Using sterile medium as the reference, calibrate the colorimeter (i.e. set it
to zero).
Note: keep this reference medium (the blank) in the refrigerator
throughout experiment.
6 Shake the flask containing the yeast culture gently to distribute the cells
evenly. Using aseptic technique, withdraw a 5 cm 3 sample.
7 Measure the absorbance of the sample you have just withdrawn. Record
date, time and absorbance in the table.
8 Incubate at 30C.
9 Repeat instructions 5 – 8 three times per day for the next three days (early
morning, lunch-time and late afternoon if possible). If it is not possible to
measure the absorbance at the time of taking the sample, place it in a
sterile container, label with initials, date and time and refrigerate until
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10
11
12
13
convenient to do so, preferably within 24 hours. Make sure that the yeast
is fully suspended before reading the absorbance.
Plot absorbance vs time.
Identify on your graph the lag, log and exponential phases of the growth
curve.
From the exponential (log) phase of growth curve, work out the time in
hours taken for the absorbance and hence the population size to double.
Calculate growth rate constant.
SUPPLEMENTARY STUDENT INFORMATION
Calculation of growth rate constant
Growth rate constant, k, is a measure of the number of generations (the
number of doublings) that occur per unit of time in an exponentially growing
culture.
k=
n 2
g
where n 2 is the natural log of 2 (determine this from your calculator) and g is
the time in hours taken for the population to double during the exponential
phase of growth.
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DETERMINING DRY WEIGHT USING TURBIDITY: VERSION 1
Source: Adapted from SAPS ‘Growth Curve: Determination of Dry Weight
from Standard Curve’.
http://www-saps.plantsci.cam.ac.uk/worksheets/scotland/curve.htm
Technical guides are also available from the same source.
PREPARING FOR THE ACTIVITY
Read Through the Student Activity Guide and consider the following
questions.
Analysis of activity
What is the aim of the activity?
What measurements are you going to make?
How will you record these measurements?
How will you determine concentration of yeast cells in a growing culture?
Getting organised for experimental work
In your group decide how the activity will be managed by allocating tasks to
each member.
Recording of data
Prepare tables and 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 is the significance of using dry mass as the measurement of
concentration?
What were the limitations of the equipment?
What were the sources of error?
What possible improvements could be made to the experiment?
What ideas do you have for further work?
What is the economic importance of the process which you are studying and
the calculations which you will make?
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STUDENT ACTIVITY GUIDE
Introduction
In industry it is often important to determine the actual concentration of cells in
a growing culture. This may involve counting total numbers of cells, numbers
of viable cells or cell concentration in terms of dry mass.
In this experiment you will use commercially available dried yeast to produce
a standard curve of absorbance at 440 nm against concentration (dry mass in
grams/litre). You will then use your standard curve to determine the
concentration of yeast in each of a series of diluted samples. [These can be
from a previous experiment.
Equipment and materials
Materials required
Materials required by each student/group:
100 cm3 sterile yeast glucose broth in flask or bottle
11 large test tubes (or universals)
test tube rack
dried yeast (not fast acting)
weighing boat
spatula
4 x 10 cm3 syringes (or pipettors with tips)
1 cm3 pipette
cuvettes
discard jar containing 1% Virkon
graph paper
marker pen
Materials to be shared:
Balance (accurate to 0.01 g)
Colorimeter (440 nm)
Crushed ice (optional)
Instructions
In order to make up the appropriate concentrations of yeast you will weight out
0.2 g dried yeast and add it to 20 cm3 sterile broth to give a concentration of
dry mass of 10 grams/litre. You will then use this suspension (the standard
dilution) to make further dilutions giving you yeast concentrations in g/l of 5,
4, 3, 2.5, 2, 1.5, 1.0, 0.5, 0.25, 0.05. Look at the table carefully and make
sure you understand it before you start.
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You may need crushed ice to keep your tubes cool while you carry out the
experiment.
1 Withdraw 20 cm3 broth from a flask of sterile yeast glucose broth and add
to a test tube (or universal).
2 Add 0.2 g dried yeast to the broth in the test tube. This will give you a
concentration of 10 g/litre. Shake gently occasionally until the cells are
fully resuspended. This is your standard dilution. Use it to make further
dilutions.
3 Draw a table with test tube number, yeast concentration in g/l and
absorbance at 440 nm.
4 Label clean test tubes 1 – 10. Add the volume of sterile broth shown in
Row A in the table below to each tube. Pay close attention as you are
required to add different volumes to the tubes.
5 From your standard dilution add the volume shown in Row B to tubes 1 –
5. Shake the standard dilution tube gently each time before removing the
sample. Make sure that you understand how the yeast concentration in g/l
is worked out.
6 Add the volumes from the numbered tubes shown in Row B to tubes 6 –
10.
Note: Pay careful attention to tube numbers and volumes.
Remember to suspend cells by shaking gently before taking a sample and
use a fresh syringe for each tube.
Dispose of used syringes and pipettes in the discard jar.
Make sure that you understand how the yeast concentration in g/l is
worked out.
A
B
Tube no
1
Volume
5
sterile
broth (cm3)
Volume
5
yeast
suspension
(cm3)
2
6
3
7
4
7.5
5
8
6
5
7
5
8
5
9
5
10
9
4
3
2.5
2
yeast conc. 5
(g/l)
4
3
2.5
2
5
cm3
from
tube
3
1.5
5
cm3
from
tube
5
1
5
cm3
from
tube
7
0.5
5
cm3
from
tube
8
0.25
1
cm3
from
tube
8
0.05
7 Using sterile broth as the reference, calibrate the colorimeter (i.e. set it to
zero) at 440 nm.
8 Starting with tube 10 shake the test tube to distribute the cells evenly and
use a pipette to transfer about 3 cm3 into a cuvette.
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9 Measure the absorbance at 440 nm of the sample you have just
withdrawn. Record it in the table.
10 Return the sample to its original tube and repeat steps 7 and 8, using the
same pipette and cuvette, to obtain readings for tubes 9 – 1.
11 Draw a graph of absorbance at 440 nm vs. yeast concentration
(grams/litre). This is the standard curve which you will use to determine
yeast concentration from absorbances of serially diluted samples in earlier
experiments e.g. viable count experiment.
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Using the standard curve to determine concentration in grams/litre from
known absorbance
1 Use the absorbances measured in the previous experiment.
2 Draw a table showing time of sample, absorbance and concentration.
3 Fill in the time and absorbance rows.
4 Carry out the following for each absorbance:




Mark the value on the absorbance (vertical) axis of your standard
curve
Draw a horizontal line till it meets the standard curve (a). Mark the
point
Draw a vertical line from there to meet the yeast concentration axis (b)
Read the value and complete the concentration row on the table
5 Determine the dry weight of yeast (in g/litre) present in the original sample.
6 Write a report on your practical placing particular emphasis on evaluation
of the equipment and methods used with respect to the resulting accuracy
and reliability.
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DETERMINING DRY WEIGHT USING TURBIDITY: VERSION 2
Source: James Watt College

Take stock culture of micro-organism & serial dilute to 10-7.
e.g.
1 ml
1 ml
1 ml
9 ml
diluent
9 ml
diluent
9 ml
diluent
= 10-1
= 10-2
= 10-3
1.0 ml
stock
10-7
This will give 8 samples in total (including original stock).

Measure the turbidity of each of these samples using a spectrophotometer
set to 600 nm.

Plot graph of absorbance against dilution factor: is there a direct
relationship between dilution and turbidity?

Given the following piece of reference information, convert your
absorbance readings to quantities of actual micro-organism (a sample
calculation is provided to help).
Absorbance at 600nm value of 0.5 is equivalent to a total cell mass of 1
mg dry cells per ml.
Example of calculation
A600 = 0.298 from machine for 10-1 dilution
0.298 x 1.0 mg/ml = 0.596 mg/ml for 10-1 dilution
0.500
Correcting for dilution factor, this becomes
0.596 x 10  5.96 mg/ml.
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CARBOHYDRATE FERMENTATION
Source: James Watt College
Carbohydrate utilization tests
Some bacteria can use a particular carbohydrate as a carbon and energy
source. Acid end-products can be identified using a pH indicator dye while
CO2 is detected in liquid culture using a Durham tube (inverted small test
tube). Aerobic breakdown is termed oxidation while anaerobic breakdown is
known as fermentation. Identification tables usually incorporate tests for
several different carbohydrates.
Durham tube in carbohydrate utilization broth. Air within the Durham tube is
replaced by broth during the autoclaving procedure.







Collect the following materials:
 24 hour broth cultures of E. coli, Staphylococcus epidermis
albus, Erwinia carotovora and Micrococcus luteus.
 Phenol red dextrose, lactose and sucrose peptone broths in
test tubes also containing Durham tubes.
Label one tube of each type of medium and inoculate with E. coli.
Label one tube of each type of medium and inoculate with Staphylococcus
epidermis albus.
Label one tube of each type of medium and inoculate with Erwinia
carotovora.
Label one tube of each type of medium and inoculate with Micrococcus
luteus.
Label one tube of each type of medium and do not incubate i.e. control
tubes.
During incubation do not tip the fermentation tube as this may accidentally
force a bubble of air into the Durham tube to give a false positive result.
Mix tubes by rolling them back and forth between the palms of the hands.
Incubate tubes at 30C for 24 – 48 hours.
Examine all cultures for evidence of acid (pH change), and/or acid and gas
production for each sugar.
Determine type of fermentation occurring.
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DIGESTION OF CELLULOSE
SOURCE: “Microbial Friends & Allies” BBSRC publication (originally
developed by NCBE http://www.ncbe.reading.ac.uk/ )
Cellumonas bacteria secrete cellulase enzyme so could potentially be used to
utilise cellulose waste (eg paper) as feedstock for their fermentation process,
converting low cost start materials into products of greater value.
1 Use sterile forceps to place a disc of sterile filter paper on the surface of a
nutrient agar plate. Use the tips of the forceps to smooth the paper onto
the agar, to ensure that there is a good contact.
2 Take the lid off the Cellulomonas culture bottle. Keep the bottle top in your
hand – do not place it on the bench. Briefly pass the neck of the bottle
through a Bunsen burner flame.
3 Dip a sterile cotton wool bud into the culture. Do not allow the culture to
drip on the bench. Flame the neck of the bottle again and replace the lid.
4 Quickly ‘paint’ a message or picture on the filter paper with the culture (see
diagram below). Dispose of the cotton wool bud into a beaker of
disinfectant.
5 Seal the Petri dish diagonally with a small amount of tape. Label the base
of the Petri dish with your initials, the date, and the name ‘Cellulomonas’.
6 Incubate the Petri dish at 25 - 30C in an inverted position: the filter paper
should stick to the surface of the nutrient agar.
2 or 3 weeks later
After incubation, the sealed plates may be examined for digestion of the filter
paper under a binocular microscope or using a hand lens. You should be able
to read the message/see the picture as the paper should have been digested
where the Cellulomonas bacteria were painted.
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TESTING VIABILITY OF YEAST AT DIFFERENT STAGES OF THE
AUTOLYSIS PROCESS
SOURCE: SAPS
http://www-saps.plantsci.cam.ac.uk/worksheets/scotland/yeast.htm
Teacher and technical guides can also be found at this site.
Preparing for the Activity
Read through the Student Activity Guide and consider the following questions.
Analysis of activity
What is the aim of the activity?
What is being varied in the activity?
What measurements are you going to make?
Getting organised for experimental work
What precautions you must be taken to prevent contamination of the agar
plates?
Can you successfully examine material under a microscope at x400
magnification?
In your group decide how the activity will be managed by allocating tasks to
each member. It is important that you play an active part in setting up the
experiment and in collecting results.
Recording of data
Prepare summary tables to record your group results.
You should use a ruler, correct headings and appropriate units.
Evaluation
How effective were the tests which you used?
What were the limitations of the equipment?
Were there any possible sources of error?
What possible improvements could be made to the experiment?
What is the biological importance of the process which you are investigating?
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Student Activity Guide
Introduction
Yeasts are versatile micro-organisms which have been used for centuries by
man to produce bread and alcoholic drinks.
In more recent years they have been used to produce flavourings for the food
industry. The yeast is grown in huge fermenters to produce biomass –
Upstream Processing and then it is treated in different ways to produce
different flavourings. These flavourings are found in most of the savoury
snack foods which we eat – crisps, soups, snacks etc.
The yeast goes through a series of different treatments to develop the huge
variety of different end products. All of these treatments involve the process
of autolysis and are examples of what is known as Downstream Processing.
The yeast products may be powders, granules or pastes and they are then
incorporated into processed foods to provide natural flavourings.
This process of AUTOLYSIS (auto-self; lysis-splitting) involves killing the
yeast and encouraging the breakdown of the cells by enzymes. These may
be the cells own endogenous enzymes or enzymes may be added. It is these
products of enzyme degradation which produce the specific flavour molecules.
Autolysis usually begins with the addition of salt to the cells, causing water to
leave the cells by osmosis and beginning the process of cell breakdown. The
cells are then heated encouraging further breakdown of the cells.
In this practical you are going to carry out the process of autolysis and try to
find at what point in the process the cells actually die. You will salt and heat
yeast and then test the viability by plating out the treated yeast to see if it will
grow. You will also test the autolysed product to see if the dehydrogenase
enzymes are active and to see if the cells take up methylene blue dye.
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Yeast Autolysis
Equipment and materials
Materials required by each student/group:
10 g fresh yeast
salt
5x 200 cm3 beakers
stirring rod
Materials to be shared:
Ovens at different temperatures
Instructions
1 Autolyse the yeast in 4 different ways. You could alter the temperature at
which the yeast is autolysed or the amount of salt which is added to the
yeast. One possible regime is suggested below:




2 g yeast 1 g salt at room temp overnight
2 g yeast 1 g salt at 40C overnight
2 g yeast 1 g salt at 60C overnight
2 g yeast 1 g salt at 80C overnight
2 Collect dried yeast samples and rehydrate by slowly adding water to the
dried sample, stirring constantly. Continue to add water to the samples to
make them up to 100 cm3.
3 Autolyse a fresh sample of yeast. Mix 2 g yeast with 1 g salt and add
water to make this sample up to 100 cm3.
You can now test the viability of these yeast samples by 3 different methods.
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Method 1: Plating Out Autolysed Yeast Samples
Streaking an agar plate
Equipment and materials
Materials required by each student/group:
Disinfectant and cloth
3 Yeast agar plates
Metal inoculating loop
Rehydrated yeast samples
Marker pen
Sellotape
Materials to be shared
Incubator at 30C
Instructions
1 Make sure that you are working in an area which has been swabbed with
disinfectant.
2 Turn your petri dish upside down and use a pen to mark the base, as
shown, and label it with the yeast samples to be used, the date and your
initial.
1
2
3 Open your petri dish and using a sterile inoculating loop, place your
sample of yeast onto one side of the petri dish. Make a shape like this.
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4 Repeat with other yeast samples and plates.
5 Seal your plates with tabs of sellotape and place it in an incubator at 30C.
6 Record any growth of yeast over the next 5 days.
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Method 2: Comparing the activity of the dehydrogenase enzymes
present in autolysed yeast samples.
Background Information
During a metabolic pathway such as aerobic respiration glucose is gradually
broken down and energy is released. Hydrogen is released from the glucose
in a process knows as oxidation. This hydrogen binds to a co-enzyme and
each reaction is catalysed by an enzyme known as a dehydrogenase.
Although it would not be possible to detect this reaction in a test-tube some
chemicals such as resazurin dye change colour when they gain hydrogen.
It changes colour in the following ways:
BLUE
LILAC
Unreduced
MAUVE
PINK
Partially
reduced
COLOURLESS
Reduced
You can use this reaction to compare the activity of the dehydrogenase
enzymes present in each of your autolysed yeast samples.
The time it takes for the dye to change colour will indicate the activity – the
faster the colour change takes place the greater the activity of the
dehydrogenase enzymes.
Activity of enzymes such as dehydrogenases would indicate that the yeast is
likely to be viable.
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Equipment and materials
Materials required by each student/group:
Rehydrated yeast samples
1 test-tube rack with 5 test tubes
5 labels
1 stop clock
1 pair safety spectacles
1 syringe/measuring cylinder
16 cm3 Resazurin dye
20 cm3 5% glucose solution
colour chart
Materials to be shared:
Waterbath at 35C
Instructions
1 Collect the materials indicated above.
2 Label 5 test-tubes and add 3 cm3 of resazurin dye to each tube.
3 Add 3 cm3 of the appropiate yeast suspension to the labelled tubes.
4 Shake each tube and place in a water bath at 35C.
5 Using the colour chart, record the colour of each tube every 2 mins for 20
mins.
If you do not get a reaction in any of the tubes after 10 minutes add 3 cm 3
of 5% glucose solution to each test-tube and shake.
6 Record the results in a table with suitable headings.
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Method 3: Testing the yeast samples with methylene blue
Methylene blue dye will diffuse into the yeast cells. If the cells are living they
will pump the blue dye out but if they are dead they will remain blue.
Equipment and materials
Materials required by each student/group:
rehydrated yeast samples
microscope
microscope slides and coverslips
dropper
0.1% methylene blue solution
50 cm3 beaker
stirring rod
distilled water
Instructions
1 Place a drop of one of the diluted yeasts onto the microscope slide and
add a drop of methylene blue dye and wait 5 minutes.
2 Place the slide onto the stage of your microscope and focus.
3 Count all the blue cells and clear cells in your field of view.
4 Repeat steps 1-3 for each of the yeast samples.
You may find that there are too many cells to count, if this is the case then you
can dilute your samples and start again.
5 Record your results in a suitable table.
6 Calculate the percentage of viable ie unstained cells.
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TISSUE CULTURE: EFFECT OF M&S SALTS ON EXPLANT GROWTH
SOURCE: LTScotland National Curriculum Support Materials
Intermediate 2 Biotechnology, Unit 3 Biotechnological Processes, Student
Materials
Background information
This tissue culture technique uses (cotyledon) explants taken from the top of a
seedling. Micro-organisms grow faster than plant cells. It is important to keep
seeds, medium and equipment sterile. Follow aseptic technique
throughout.
Equipment and materials required by each student/group:
Part 1
Eye protection
Disinfectant and paper towels
10 mustard seeds
30 cm3 10% bleach in lidded container
forceps
clingfilm
new, clean, block shaped sponge
germination vessel, half filled with sterile water
Part 2 (after 2 – 3 days)
All equipment must be very clean
Disinfectant and paper towels
Bunsen burner
Eye protection
Forceps
Scissors
6 small glass containers:
2 half filled with 1% plain agar
2 half filled with 1% plain agar plus 2.2 g litre M&S salts
2 half filled with 1% plain agar plus 4.4 g litre M&S salts
clingfilm
Materials to be shared
Light bank
Hazard bags for disposal of plates
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Instructions Part 1: Sterilising seeds for germination
1 Prepare work space: clear area, swab bench with disinfectant.
2 Collect materials and equipment for part 1.
3 Place 10 mustard seeds in lidded container of bleach. Swirl seeds and
leave for 10 minutes in the bleach.
Remember your seeds are now sterile. Do not touch with your
hands.
4 Pour off bleach.
5 Rinse three times with fresh, sterile water, leaving them covered with a
little water.
6 Sprinkle the seeds onto the clean sponge, making sure the seeds are
spaced out.
7 Place sponge in germination vessel, with water level half way up sponge.
8 Label vessel with your name date and seed type.
9 Place under a light bank in a warm place (ideally 20 - 26C), for 2 to 3
days.
10 Leave until germinated and cotyledons (young leaves) have just started to
unfold.
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Instructions Part 2 (after 2 – 3 days)
1 Collect germination vessel and equipment for part 2 (see Student Guide).
2 Select the 6 straightest and longest seedlings for your experiment.
3 Label six glass containers with date and check there are 2 each of three
medium types:
You should have 6 containers:
2 with agar and no M&S salts
2 with agar and 2.2 g/l M&S salts
2 with agar and 4.4 g/l M&S salts
4 Cut off six cotyledons as shown below. These are your explants.
5 Use cooled, sterile forceps to transfer one explant into each of the six
containers. Push cut end of explant into the agar, making sure the
cotyledons are not touching the agar’s surface.
6 Cover containers with cling film and place in a warm area under a light
bank for 1 to 2 weeks.
7 Observe and note the number of explants with roots after 3, 6 and 9 days
approximately.
8 Collect class results and calculate % explants with roots for the three
different media on the three different days.
9 Present your results as three lines on a single graph (one line for each
medium) with suitable scales and axes labelled with quantities and units.
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CLONING CAULIFLOWER
Source: NCBE Practical Biotechnology at
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PRACBIOTECH/PDF/
cauli.pdf
A similar protocol, which includes colour photographs, can be found at
http://www-saps.plantsci.cam.ac.uk/docs/tissue.pdf
Materials required for each student or group of students:
Fresh cauliflower curd (the white part)
Sterile distilled water (100 cm3)
70% ethanol (50 cm3)
20% Domestos solution (100 cm3)
Boiling tubes, each containing 2-3 cm3 of plant tissue growth medium
Sterile Petri dish
Metal forceps and scalpel
Non-absorbent cotton wool and aluminium foil
White tile or suitable cutting surface
Practical details
1 Swab the working area with 70% ethanol. Keep the ethanol away from
exposed flames!
2 Cut out a small piece of cauliflower curd; roughly the size of a cherry.
Working on a clean Petri dish, divide the curd into three.
3 Place the pieces (explants) in bleach e.g. Domestos solution for 10
minutes to surface sterilize the tissue.
From this point on, quick, aseptic operations are important to prevent
contamination.
4 Rinse the explants in three successive beakers of sterile distilled water.
Use flamed, cooled forceps to do this – the correct way to flame forceps
and other instruments is to dip them in alcohol, then to pass them briefly
through a flame to ignite the ethanol. As the ethanol burns off, it heats the
surface of the instruments to 70C, killing any contaminating organisms.
Do not heat forceps and scalpels until red hot, and remember to keep
ethanol away from exposed flames.
5 The explants can be left in the final beaker of sterile water (covered with a
Petri dish lid) until required.
6 Take the first tube of growth medium withdraw the cotton wool plug, then
briefly flame the neck. Use flamed, cooled forceps to pick up an explant
and quickly drop it into the tube. Return the forceps to the ethanol beaker.
Flame the neck of the tube once more, then replace the plug.
7 Repeat Step 6 with the two remaining explants and two fresh tubes of
growth medium.
8 The tubes should be kept in a warm, light place. Growth should be visible
within 10 days. Contamination, if it has occurred, should also be visible by
this time. Failure of anything to grow usually indicates that the bleach has
not been rinsed from the plant tissue.
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Safety
Students should wear safety goggles when using bleach solution.
Ethanol used for sterilizing working surfaces should be kept away from naked
flames.
Results
Observe periodically throughout the 10 day incubation process and describe
growth.
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ENZYMES & FRUIT JUICE PRODUCTION
Source: Adapted from NCBE Practical Biotechnology
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PRACBIOTECH/PDF/
juice.pdf
Extracting Fruit Juice
The enzyme pectinase can break down the pectin found in cell walls. Fruit
juice companies use pectinase to improve the juice extraction and to produce
a clearer juice.
Read the following procedure and think carefully about what apparatus you
will need and what information you will need to record.
Remember to label your beakers and measuring cylinders
Experiment 1
Practical details

Collect a piece of apple and grate it.
Place an equal weight of grated apple into two beakers

To one beaker add 2 ml of pectinase and to the other add 2 ml of distilled
water.

Stir each beaker and leave for 5 minutes.

Filter the juice from the apples.

Record the volume of juice every 2 minutes.
Apparatus
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Results
Use the following grid and design your results table
Conclusion
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ENZYMES
Extracting Fruit Juice
Questions
1 What effect did the enzyme pectinase have on the volume of juice
produced?
___________________________________________________________
2 What effect did the enzyme have on the clarity of the juice?
___________________________________________________________
3 Why did you set up a beaker which had the apples and distilled water in it?
___________________________________________________________
4 What other factors might affect the volume of juice produced?
___________________________________________________________
___________________________________________________________
You will investigate some of these factors when you know a little more
about juice extraction.
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ENZYMES
Clarifying Fruit Juice
You have already discovered some effects of the enzyme pectinase.
Pectinase was first used by industry to clarify juice. In this experiment you
are going to look at the effects of pectinase and amylase on the clarity of
juice.
What does the enzyme amylase do?
______________________________________________________________
Experiment 2
Practical Details
Set up the following test tubes
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ENZYMES
Clarifying Fruit Juice
Add 10 ml of cloudy apple juice to each tube and stir
Incubate for 30 minutes at 40C
Observe the tubes every 5 minutes and record their appearance
Results: Make a results table to record your results
Questions
1 What effect did each enzyme have on the clarity of the juice?
AMYLASE
___________________________________________________________
PECTINASE
___________________________________________________________
2 What was the effect of using both enzymes?
___________________________________________________________
___________________________________________________________
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DEHYDROGENASE ACTIVITY IN YEAST
Source: NCBE Practical Fermentation
http://www.ncbe.reading.ac.uk/ncbe/protocols/fermentation.html
Yeasts are living organisms. They belong to the group of microbes known as
FUNGI.
Yeasts are single celled organisms which reproduce by splitting into two.
This process in yeast is known as budding.
Yeast has been used for thousands of years to make beer, bread and wine.
These technologies have developed because yeast cells can produce carbondioxide and alcohol when they grow. The process is known as fermentation.
Nowadays yeast is still used in the brewing and baking industries but it is also
used to produce new products such as flavourings for foods, fizzy drinks and
the enzymes needed to make cheese.
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Dehydrogenase Enzymes in Yeast
During aerobic respiration glucose is gradually broken down and energy is
released.
glucose + oxygen  energy + carbon dioxide + water
In a metabolic pathway such as this, it is usually the removal of hydrogen
from the glucose which allows the energy to be released. The removal of
hydrogen is called oxidation. As the glucose is oxidised, hydrogen is
released at various stages along the pathway. This hydrogen binds to a
chemical called a co-enzyme and each reaction is catalysed by an enzyme
known as a dehydrogenase.
Therefore dehydrogenase enzymes catalyse the oxidation of substrates by
transferring hydrogen to co-enzymes such as NAD. These co-enzymes carry
the hydrogen as they become reduced.
AH2
substrate
e.g. glucose
+
2 NAD
co-enzyme
A
oxidised
substrate
+
2 NADH
reduced co-enzyme
Although it would not be possible to detect this reaction in a test-tube some
chemicals such as resazurin dye change colour when they gain hydrogen.
We say the chemical has become reduced.
It changes colour in the following ways:
blue
(unreduced)
lilac
mauve
(partially reduced)
pink
colourless
(reduced)
This reaction enables you to test to see if respiration is taking place. You can
also use it to investigate respiration rates as you alter variables. Yeast is a
suitable organism to use for these investigations.
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Experiment – Testing respiration rate in yeast
Apparatus
3 test tubes
3 labels
5 ml or 10 ml syringe
resazurin dye
fresh yeast solution
boiled and cooled yeast solution
glucose solution
water bath
timer
Method
1 Label 3 test-tubes A, B & C
2 Add 3 ml of resazurin dye to each tube
3 Add 3 ml of glucose solution to A & C
Add 3 ml of water to B
4 Add 3 ml of fresh yeast suspension to A & B
Add 3 ml of boiled and cooled yeast suspension to C
5 Shake each tube and place in a water bath at 35C
Result
Record the colour in each tube in the table below
Time in
minutes
A
B
C
0
Blue
3
Lilac
6
Mauve
9
Pink
12
Colourless
15
18
21
24
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QUESTIONS
1 Give an example of a metabolic pathway __________________________
___________________________________________________________
2 What happens to the hydrogen when it is first released from the glucose?
___________________________________________________________
___________________________________________________________
3 What do dehydrogenase enzymes do?
___________________________________________________________
___________________________________________________________
4 What is a reduced coenzyme?
___________________________________________________________
5 What happens to resazurin dye when hydrogen is added to it?
___________________________________________________________
6 In which test-tube did the yeast respire most rapidly?
___________________________________________________________
7 What was the purpose of test-tube C?
___________________________________________________________
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DNA FROM KIWI FRUIT
Source: Adapted from NCBE Illuminating DNA
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PRACBIOTECH/PDF/
onion.pdf
In this practical you are going to isolate DNA from plant cells (i.e. not from
micro-organisms).
Detergent is used to degrade the cell and nuclear membranes. Cell
fragments are separated by filtration leaving the DNA and the soluble proteins
in the filtrate. A protease enzyme called neutrase breaks down the protein
and then the DNA is precipitated using ice cold ethanol.
QUESTIONS
1 Which part of the cell and nuclear membrane is the detergent degrading?
___________________________________________________________
2 What will the neutrase enzyme break its substrate into?
___________________________________________________________
APPARATUS
A balance
Weigh boat
Sodium chloride
Washing up liquid
5 ml syringe
2 x 250 ml beaker
50 g of kiwi fruit
stirring rod
coffee filter paper
measuring cylinder
boiling tube
ice cold ethanol
A liquidizer, an ice bath and a water bath at 60C must be available in the lab.
METHOD
Part 1 Preparing the Fruit extract
1 Add 1.5 g of sodium chloride and 5 ml of
washing up liquid to a 250 ml beaker.
2 Make up to 50 ml with distilled water.
3 Now, collect 50 g of fruit and chop it up, then add it to the salt/detergent
solution.
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4 Stir the mixture and incubate for 13 minutes at 60C
5 Cool the mixture in an ice bath for 4 minutes.
6 Pour the mixture into a liquidizer and blend for 5 seconds
at high speed.
7 Filter this mixture into a second beaker.
Separating the DNA
1 Add 4 ml of fruit extract to a boiling tube
2 Add 2 drops of neutrase enzyme to the extract
3 Slowly trickle 6 ml of ice-cold ethanol very slowly
down the side of the test-tube.
Two layers of liquid should form.
4 Leave the tube for 2-3 minutes without disturbing it.
5 Gently rock the liquids taking care not to let the two
layers mix too much.
RESULT
Describe the DNA which you have precipitated
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DNA FROM CAVIAR
Source: online Bioscience journal
http://www.bioscience-explained.org/EN1.1/pdf/CaviarEN.pdf
AIM
This simple practical procedure allows the isolation of impure DNA from
‘caviar’ or fish eggs. The result is a pellet of thread-like material, which
includes DNA but will still be contaminated with lipids, carbohydrates and
proteins.
EQUIPMENT AND MATERIALS
Needed by each person or group

20-30 g caviar (about 2 heaped teaspoonsful) e.g. roe from capelin
(Mallotus villosus) or lumpsucker (Cyclopteris lumpus).
Note: such roe is sold under the Abba® brand name. The yellow or
‘natural’ variety works best.

15 ml washing-up liquid e.g. Fairy Liquid, diluted 1:10 with distilled water

1 teaspoon (about 6 g) of table salt

2 ml ethanol. This must be ice-cold and at least 80% pure.

3-4 drops of protease, e.g. Novozymes Neutrase ®

Glass rod

Coffee filter paper

Funnel

Small test tube

Dropper or pipette for dispensing the enzyme

Pasteur pipette, the tip of which has been melted and curved to form a
small hook
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PROCEDURE
1 Add the caviar and salt to a mortar, then crush the eggs using a pestle.
The shells of the eggs have to be broken. Proteins are precipitated by the
salt.
2 Add the washing-up liquid solution to the mortar. The liquid should cover
the caviar completely. The detergent dissolves lipids from the membranes
of the roe.
3 Add 3-4 drops of protease to the mixture and stir vigorously. The enzyme
will partially degrade any soluble proteins.
4 Filter the mixture through the coffee filter and collect the filtrate in a clean
test tube.
5 Add the ice-cold ethanol by carefully pouring it along the wall of the tube or
use a pipette and add it at the bottom of the test tube. DNA precipitates as
long threads in cold ethanol and can be found at the interface between the
detergent solution and the ethanol.
6 Collect the DNA with the help of a Pasteur pipette with a hooked tip. The
DNA may be transferred to a microcentrifuge tube and stored, frozen, for
later use e.g. for gel electrophoresis or staining of the DNA.
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RESTRICTION ENZYME DIGESTS & DNA GEL ELECTROPHORESIS
Source: Adapted from NCBE ‘The Lambda DNA Protocol’ at
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PDF/LambdaSG.pdf
A technical/teachers guide to this activity can be found at
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PDF/LambdaTG.pdf
1 CUTTING THE LAMBDA DNA WITH RESTRICTION ENZYMES
The restriction enzymes are in the following tubes:
Yellow – empty
Green – HindIII
Blue – Bam H1
Pink – EcoR1
Wear gloves (to protect the DNA from digestion by enzymes in your sweat).
Add a fresh tip to the Gilson pipette.
Practice aspirating and ejecting 10l of blue dye. [Volume errors in pipetting
when using such small volumes can make big errors in your results].
Add a fresh tip to the pipette. Put 20 l of your chosen DNA solution into an
enzyme tube of your choice.
Mix the liquid with the dried enzyme by carefully drawing the liquid up and
down in the tip a few times. The liquid in the enzyme tube should have a
distinct blue colour and there should be no concentration of dye at the bottom
of the tube.
Cap the tube containing the enzyme and DNA, then incubate it at 37C in a
waterbath for 30-45 minutes.
Repeat with the other enzyme tubes using a fresh tip each time. It helps to
flick the tubes occasionally during incubation, to ensure that their contents are
thoroughly mixed.
2 LOADING THE GELS
After the incubation, add 2 l of loading dye to each enzyme tube using a
fresh tip. [The loading dye contains sucrose so the sample will sink to the
bottom of the well in the gel and not float away in the buffer in the tank. It also
is blue so you can see your samples].
Mix the loading dye and the DNA sample very thoroughly by drawing the
mixture up and down the pipette tip.
Put the gel and tank over a piece of black paper or card to make the wells
more visible. Pour 10 ml of TBE buffer onto the gel. Load 10l of the mixture
of loading dye and DNA into one of the wells, holding the tip above the well
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but under the buffer solution. Take great care not to puncture the bottom of
the well with the pipette tip. Depress the pipette plunger gently and the DNA
and dye mixture should sink into the well as it leaves the pipette tip.
Make a note of which DNA you have put in each well.
Repeat these steps with the other DNA samples using a fresh tip for each
sample.
Load dye markers (5 l) in one well.
3 RUNNING THE GEL
Fit a piece of carbon fibre tissue to each end of the tank so one end of it is in
the buffer.
Check the power pack is turned OFF.
Put a lid on your tank.
Using the red and black leads, connect the tissue to the power source. The
red lead should go to the positive and be at the end of the tank furthest from
the wells. The black lead should go to the negative and be at the end nearest
to the wells. [The reason it is this way round is that DNA in this running buffer,
has a negative charge and will be attracted to the positive terminal (anode)
when a current is applied].
After a few minutes you should see bubbles forming at the cathode (negative
end of the tank). After a few more minutes, the DNA and dye mixture should
start migrating through the gel. If it is going the wrong way, or not moving at
all, you have connected your leads up incorrectly.
Try again.
The gel should run until the blue dye (which runs in front of any DNA
fragments) reaches the end of the gel. This will probably take a few hours.
Switch off the power and remove the leads. If you let it run too long, the DNA
will come off the end of the gel and be lost in the buffer!!!
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4 STAINING THE GEL
Wear plastic gloves.
Pour off the buffer solution and pour the stain onto the surface of the gel.
Leave it for exactly 4 minutes, then return stain to the beaker/stain bottle.
Very carefully wash the gel surface with cold distilled water to remove excess
stain. Repeat 3-4 times, finally pouring off all the water from the gel.
The remaining stain will gradually move down through the gel, staining the
DNA as it does so. Faint bands should start to appear after 10 minutes.
Better results are seen if the gel is left to develop over night. Leave the gel in
a plastic bag in the fridge, to prevent the gel from drying out.
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-GALACTOSIDASE INDUCTION
Source: Adapted from NCBE Illuminating DNA
http://www.ncbe.reading.ac.uk/ncbe/PROTOCOLS/DNA/PDF/DNA06.pdf
The lac operon is the classic example of gene regulation, in which the
production of -galactosidase (lactase) is induced by the presence of lactose
in the growth medium. In this practical task, ONPG, rather than lactose, is
used as a substrate for the enzyme.
Aim
To induce and measure the production of the enzyme -galactosidase
(lactase) by E.coli.
Day 1: Preparation
You will need cultures of E.coli from a strain that possesses the lacZ (galactosidase) gene. These can be grown on solid agar 24-48 hours in
advance. To induce the production of -galactosidase, lactose must be
present in the growth medium.
From stock E.coli plate prepare the following two streak plates:
E.coli on nutrient agar
E.coli on nutrient agar and lactose.
Incubate at 30C for 24-48 hours.
Day 2: Timing
This activity takes about 60 minutes, including an incubation period of 10
minutes.
Materials and equipment needed by each person or group










Cultures of E.coli
ONPG (ortho-nitrophenyl--D-galactoside) solution (2 cm3 per test sample)
Methylbenzene; 1 drop per test sample
Test tubes, caps, rack and marker pen
Inoculation loop
Pasteur pipettes
5 cm3 syringe, for transferring ONPG solution
Waste container with disinfectant
Stopclock
Safety spectacles
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Procedure
Quick qualitative method
1 Using a syringe, transfer 2 cm3 of ONPG solution into each of two test
tubes. One of these tubes will be a ‘control’; the other will be for the test
culture.
2 Label the tubes appropriately.
3 Use a flamed wire loop to aseptically transfer a large colony of E.coli from
the nutrient agar plate into the test solution. Suspend the microorganisms
by agitating the loop, then sterilise it by flaming. Take care to introduce
the loop into the flame slowly, to avoid sputtering! Repeat the process,
this time transferring an E.coli colony from the plate where nutrient agar
contains lactose, into the test solution.
4 Add a drop of methylbenzene to each tube, cap the tubes and shake well
to mix. Methylbenzene kills the cells and partially disrupts the cell
membranes, allowing the ONPG to diffuse into the cells.
5 Let the test tubes stand on the bench, until a strong yellow colour
develops. This generally takes 5-20 minutes. The reaction can be
speeded up by incubating the tubes at 37C. The colourless ONPG is
broken down by -galactosidase to produce galactose and orthonitrophenyl (ONP). ONP is bright yellow in alkaline conditions.
6 Compare the colour of the two tubes.
Safety
Handling microorganisms and methylbenzene

Good microbiological practice must be observed when handling
microorganisms.

Methylbenzene is flammable and produces harmful
vapour. Large volumes should therefore be handled
in a fume cupboard, although the small amounts used
here can safely be handled at the bench (but keep
away from flames). Skin and eye contact should be
avoided.

Eye protection must be worn.
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Qualitative method
STEPS
1&2
Transfer 2cm3 of ONPG solution into each of two test tubes. Label the
tubes appropriately.
3.
Aseptically transfer a colony from the plate to one of the tubes of
ONPG solution. Twiddle to disperse the cells, then flame the loop to
kill any remaining cells.
4-6.
Add a drop of methylbenzene to each tube, cap the tubes and shake
well to mix. Stand the tubes for 5-20 minutes until a yellow colour
develops.
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GENETIC CONTROL
Source: Adapted from NCBE Practical Biotechnology
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PRACBIOTECH/PDF/cat
milk.pdf
The genes found in living cells control the production of proteins such as
enzymes. Any one cell can contain many thousands of genes all coding for
different proteins. In a multicellular organism only a small proportion of the
genes available within the cell will be required. Scientists think that genes
become switched on or off as they are required. We are going to look at the
gene which codes for an enzyme which breaks down the sugar LACTOSE.
The, so called, Jacob-Monod hypothesis suggests that the gene which
produces the enzyme required to break down lactose becomes switched on
only when the substrate is present. When there is no lactose present the
gene would be switched off.
Firstly you will set up an experiment to investigate the effect that the enzyme
has on its substrate.
Production of Lactose – free Milk
The naturally occurring sugar found in milk is lactose. This disaccharide
sugar is broken down to glucose and galactose by the enzyme 
galactosidase. Many people (75% of the world’s adult population) cannot
digest lactose because they cannot produce this enzyme. They are known as
lactose intolerant. These people cannot consume milk or milk products.
In this experiment you are going to attempt to produce lactose free milk by
immobilising the  galactosidase enzyme and setting up a continuous flow
system.
APPARATUS
 galactosidase enzyme
calcium chloride
sodium alginate
milk
2 x 10 ml syringe
250 ml beaker
2 x 100 ml beaker
circle of gauze
stirring rod
tea strainer
3 way tap
2 clinistix
paper towel
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METHOD
Step 1 Immobilising the  galactosidase
1 Use a 10 ml syringe to draw up 2 ml of  galactosidase enzyme
2 Collect a piece of paper towel and then add 8 ml of sodium alginate to the
same syringe (use the paper towel to catch any alginate drips)
3 Rock the syringe until the two liquids mix completely (this may take about
5 minutes)
4 Collect a 250 ml beaker and add 100 ml of calcium chloride
5 Add the alginate/enzyme mixture to the calcium
chloride one drop at a time. Leave the beads in
the calcium chloride for 3 minutes to allow them
to set.
6 Filter the immobilised enzyme beads from the
calcium chloride solution and then rinse the beads
in distilled water
Step 2 Setting up a Continuous Flow System
7 Collect a 3 way tap and check that you know how
to use it. Practise with water.
8 Place a piece of nylon gauze at the base of a
10 ml syringe and attach a 3 way tap to the end of
the syringe.
9 Add the beads to the syringe.
10 Collect 25 ml of milk in a beaker.
11 Test the milk for the presence of glucose using a
CLINISTIX (Clinistix changes from pink to purple
if glucose is present).
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12 Slowly add the milk to the syringe and alter the 3 way tap to ensure that
the milk passes through at a slow but steady rate. Collect the treated milk
in a small beaker.
13 Test the milk produced for the presence of glucose
RESULT:
Clinistix colour
Glucose presence
(+ or -)
Milk before enzyme
treatment
Milk after enzyme
treatment
CONCLUSION:
QUESTIONS:
1 Name the enzyme which breaks down lactose ______________________
2 What induces the production of this enzyme? ______________________
___________________________________________________________
3 Give 2 advantages of using a continuous flow system, such as this, rather
than a batch process to produce lactose free milk.
___________________________________________________________
___________________________________________________________
4 Cheese making produces the waste product whey which is rich in lactose.
The whey is usually dumped at sea. Describe a way in which this waste
product might be upgraded into a useful product.
___________________________________________________________
___________________________________________________________
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Lactose Metabolism in E. Coli
The bacterium E. coli produces  galactosidase only when lactose is present
in it’s growing medium. The Jacob-Monod hypothesis suggests that the
genes controlling the production of this enzyme work in the following way.
There are 3 genes involved:
The structural gene – codes for the enzyme
The operator gene – switches on the structural gene
The regulator gene – codes for a repressor molecule
DNA
chain
Regulator
gene
Operator
gene
Structural gene
DNA
chain
Repressor
molecule
The presence of lactose induces the production of the enzyme. Lactose is
therefore known as the inducer.
LACTOSE ABSENT
When lactose is absent the repressor molecule combines with the operator,
the structural gene is switched OFF and no  galacosidase is produced.
DNA
chain
Regulator
gene
Operator
gene
Structural gene
OFF
DNA
chain
repressor
LACTOSE PRESENT
When lactose is present some of it binds with the repressor and the operator
gene is able to switch ON the structural gene and  galactosidase is
produced.
DNA
chain
Regulator
gene
repressor
Operator
gene
Structural gene
ON
DNA
chain
lactose
When all the lactose has been digested the repressor molecule will bind again
with the operator and the gene will be switched back off.
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QUESTIONS
1 Name the 3 genes involved with this hypothesis ____________________
___________________________________________________________
2 Which gene codes for the enzyme? ______________________________
___________________________________________________________
3 What does the repressor molecule do? ___________________________
___________________________________________________________
4 Why is lactose known as the inducer? ____________________________
___________________________________________________________
5 Why might this system of gene control be described as energy-efficient?
___________________________________________________________
___________________________________________________________
6 Complete the blanks in the following passage.
In E. coli a _________________________ gene codes for the production
of an enzyme called ____________________. When lactose is absent a
___________________ molecule binds with the ___________________
gene and the structural gene is switched ____________________ .
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ANTIBODY/ANTIGEN REACTIONS
Source: James Watt College
Suppliers of antibodies (antiserum) and antigens include Alba Bioscience
(formerly the Scottish Antibody Production Unit)
http://www.show.scot.nhs.uk/sapu/index.html and Sigma-Aldrich
http://www.sigmaaldrich.com/Area_of_Interest/Europe_Home/UK.html
This simple experiment utilizes (purified) agar plates with wells cut into them
as depicted here:
Antibody
Ab1
Distance between
central well & test
wells must be equal
Ab4
Ab2
Ab3
Antibody (Antiserum) is dropped in the central well and antigen (serum) from
a variety of animals is placed in surrounding wells.
If the antibodies meet recognizable antigens, the proteins react and
precipitate, giving a visible line.
Whether a reaction occurs depends on the evolutionary relatedness of the
animals, so that sheep antiserum reacts with goat serum but not with mouse
serum. This point illustrates the specificity of the antibody/antigen reaction.
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Set up TWO plates as shown here: note which antibody you put in the central
well
1
4
2
3
Note which Antigens you put in wells 1  8:
Antigen 1 =
Antigen 2 =
Antigen 3 =
Antigen 4 =
5
6
8
7
Antigen 5 =
Antigen 6 =
Antigen 7 =
Antigen 8 =
* Remember to include a control
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THE EFFECT OF DISINFECTANTS AND ANTISEPTICS ON MICROBIAL
GROWTH
Source: Based on material from biology4all website
http://www.biology4all.com/resources_library/source/2.doc
AIMS
Determine the effect of chemical agents on bacterial growth.
1 Describe the physical effect on the growth of bacteria on solid nutrient
agar.
2 Evaluate the effectiveness of commonly available antiseptics and
disinfectants.
3 Discuss the inaccuracies that are inherent in the filter paper disk method.
INTRODUCTION
You will study how certain commonly available disinfectants and antiseptics
affect the growth of two common bacterial species.
Disinfectants are described as antimicrobial agents used on inanimate
objects such as instruments (medical or dental), plastics or surfaces such as
kitchen worktops, toilets, washbasins etc). Disinfectants should remove
pathogenic organisms.
Antiseptics are antimicrobial agents that are used on living tissue such as
skin.
Disinfectants and antiseptics do not always sterilise because these
compounds usually do not kill all fungal and bacterial spores and vegetative
bacteria and fungi. Organic compounds e.g. dirt, grease … etc. can interfere
with their action reducing the efficiency of the antimicrobial agent.
This practical enables you to study some commonly available antiseptics and
disinfectants and assess their efficiency against two common bacteria that
can be isolated from kitchens, hospitals and from humans. You can use
household products such as Dettol or you can vary the experiment to
compare antibacterial soaps, skincare products such as facewashes etc,
toothpastes, mouthwashes…….
Useful references on the internet
Lister and antiseptics – http://web.ukonline.co.uk/b.gardner/Lister.html
E. coli – http://vm.cfsan.fda.gov/~mow/chap14.html
Staphylococcus – http://www.niaid.nih.gov/dmid/staph.htm
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MATERIALS
four disinfectants or antiseptics (can bring from home)
overnight broth cultures of:
Escherichia coli
Staphylococcus epidermis albus
sterile nutrient agar plates (x2)
sterile filter paper disks
forceps
glass spreader
METHOD
1 Using a sterile pipette remove 0.1 ml from the broth culture of Escherichia
coli, and inoculate one labelled nutrient agar plate. Spread evenly to
obtain confluent growth after incubation.
2 Do the same with the other agar plate, using the broth culture of
Staphylococcus epidermis albus.
3 Label the four antiseptics or disinfectants that you will be testing A, B, C
and D.
4 On the bottom of the agar plates, mark four sectors using a marker pen.
Label the sectors A, B, C and D.
 These sectors correspond to the letters you put on the antiseptic
or disinfectant containers.
5 Sterilise the tip of your forceps by passing it through the flame of the
Bunsen burner two or three times.
6 Aseptically pick up a sterile filter paper disk with your sterile forceps and
dip the disk into the disinfectant or antiseptic labelled A.
 Be sure that the excess disinfectant has drained off.
7 Place the disk in centre of Sector A of the Staphylococcus epidermis albus
inoculated plate.
8 Using the same disinfectant, place another disk in Sector A of the E. coliinoculated plate.
 You are comparing the effectiveness of each disinfectant or
antiseptic on both organisms.
9 Repeat steps 8 and 9, placing the other disinfectants in the other sectors.
10 Gently press the disks down with the tip of your flamed forceps to ensure
contact with the nutrient agar.
11 When all four disinfectant-soaked disks have been placed in all four
sectors of both plates, diametrically seal them with tabs of tape, invert the
plates and incubate them at 30C for 48 hours.
12 Observe, measure, and compare the zone of no growth (inhibition), if any,
around the disk for each disinfectant or antiseptic for both organisms.
13 Evaluate the effectiveness of each of the antiseptics/disinfectants at
controlling the growth of the microbes used here.
14 Describe the usefulness of this experiment in terms of controls, reliability, &
possible inaccuracies.
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TRANSFORMING BACTERIA
Source: Bio-Rad ‘pGLO Bacterial Transformation Quick Guide’
Available online at http://www.caam.rice.edu/~cox/lab1.pdf
Transformation Procedure
The aim of this experiment is to transform E.coli bacteria to express the green
fluorescent protein. The pGLO plasmid is used as a vector in this experiment.
1 Label one closed micro test tube +pGLO and another -pGLO. Label both
tubes with your group’s name. Place them in the foam tube rack.
2 Open the tubes and, using a sterile transfer pipette, transfer
250 l of transformation solution (CaCl2) into each tube.
3
Place the tubes on ice.
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4 Use a sterile loop to pick up a single colony of bacteria from your starter
plate. Pick up the +pGLO tube and immerse the loop into the
transformation solution at the bottom of the tube. Spin the loop between
your index finger and thumb until the entire colony is dispersed in the
transformation solution (with no floating chunks). Place the tube back in
the tube rack in the ice. Using a new sterile loop, repeat for the –pGLO
tube.
5 Examine the pGLO DNA solution with the UV lamp. Note your
observations. Immerse a new sterile loop into the pGLO plasmid DNA
stock tube. Withdraw a loopful. There should be a film of plasmid solution
across the ring. This is similar to seeing a soapy film across a ring for
blowing soap bubbles. Mix the loopful into the cell suspension of the
+pGLO tube. Close the tube and return it to the rack on ice. Also close the
–pGLO tube. Do not add plasmid DNA to the –pGLO tube. Why not?
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6 Incubate the tubes on ice for 10 minutes. Make sure to push the tubes all
the way down in the rack so the bottom of the tubes stick out and make
contact with the ice.
7 While the tubes are sitting on ice, label your four LB nutrient agar plates on
the bottom (not the lid) as follows:

Label one LB/amp plate:
+pGLO

Label the LB/amp/ara plate:
+pGLO

Label the other LB/amp plate:
-pGLO

Label the LB plate:
-pGLO
8 Heat shock. Using the foam rack as a holder, transfer both the (+) pGLO
and (-) pGLO tubes into the water bath, set at 42C, for exactly 50
seconds. Make sure to push the tubes all the way down in the rack so the
bottom of the tubes stick out and make contact with the warm water.
When the 50 seconds are done, place both tubes back on ice. For the
best transformation results, the transfer from the ice (0C) to 42C and
then back to the ice must be rapid.
Incubate tubes on ice for 2 minutes.
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9
Remove the rack containing the tubes from the ice and place on the
bench top. Open a tube and, using a new sterile pipette, add 250 l of LB
nutrient broth to the tube and reclose it. Repeat with a new sterile pipette
for the other tube. Incubate the tubes for 10 minutes at room
temperature.
10 Tap the closed tubes with your finger to mix. Using a new sterile pipette
for each tube, pipette 100 l of the transformation and control suspensions
onto the appropriate nutrient agar plates.
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Transformation
plates
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11 Use a new sterile loop for each plate. Spread the suspensions evenly
around the surface of the LB nutrient agar by quickly skating the flat
surface of a new sterile loop back and forth across the plate surface. DO
NOT PRESS TOO DEEP INTO THE AGAR.
+pGLO
LB/amp
+pGLO
-pGLO
-pGLO
LB/amp/ara LB/amp
LB
12 Stack up your plates and tape them together. Put your group name on
the bottom of the stack and place the stack of plates upside down in the
37C incubator until the next day.
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MAKING PROTOPLASTS
Source: Adapted from NCBE Practical Biotechnology
http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/PRACBIOTECH/PDF/pro
to.pdf
A similar protocol using lettuce leaves is available at
http://www-saps.plantsci.cam.ac.uk/docs/protofusion.pdf
Protoplasts are plant, fungal or bacterial cells which have had their cell walls
removed; usually by enzymic digestion. The resultant ‘naked’ cells can be
used in techniques such as the creation of transgenic plants.
Materials
1 lettuce leaf (from a round lettuce)
0.1 cm3 21% sorbitol solution
20cm3 13% sorbitol solution
0.5cm3 Viscozyme (mixture of carbohydrase enzymes)
Paper tissue
10ml syringe
2 x 1ml syringe
Test tube
Centrifuge tube
Stirring rod
Filter funnel
Nylon gauze to plug filter funnel
Microscope slide
Cover slip
Microscope with x40 objective
Water bath set at 37oC
Bench centrifuge
Method
Preparing the lettuce
1. Cut the lettuce leaf into pieces roughly 5mm x 5mm
2. Add 15-20 lettuce pieces to 9.5cm3 13% sorbitol solution in a test tube
3. Incubate the tube at 37oC for 5 minutes.
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Digesting the cell walls
4. Gently stir 0.5cm3 of viscozyme into the sorbitol and lettuce preparation
5. Return the tube to the water bath for another 20 minutes. Gently stir the
contents from time to time.
Recovery of the protoplasts
6. Tightly pack the spout of the filter funnel with nylon gauze
7. Pour the digested lettuce suspension into the filter funnel
8. Wash any trapped protoplasts through the filter using 10cm3 of 13%
sorbitol solution. Collect all the filtrates in a centrifuge tube.
9. Centrifuge the filtrate for approximately 5 minutes at 2000 rpm
10. Carefully pour off the supernatant, leaving a pellet of protoplasts at the
bottom of the tube.
11. Resuspend the pellet in approximately 0.1cm3 of 21% sorbitol solution.
Examining the protoplasts
12. Put some resuspended protoplasts on a slide and gently lower a coverslip
into position. Protoplasts can easily be seen without staining using a x40
objective lense.
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PLAQUE ASSAY
Source: James Watt College
This technique is used to estimate the number of phage virus particles in a
stock suspension. Plaques are clear areas in a lawn of bacteria caused by
the lysis of a phage-infected bacterial cell.
Materials
24 hour broth culture of E.coli B
Suspension of phage virus (eg Philip Harris Bacteriophage T4B)
8 sterile nutrient broths (4.5 ml each)
Gilson pipette
Sterile blue tips
10 tubes sterile sloppy (0.7%) nutrient agar: 6mls each
(NOTE: these must be kept at 45oC).
10 nutrient agar plates
Water bath at 45oC
Serial diluting the phage
1. Use sterile nutrient broth to prepare serial dilutions of the stock phage
ranging from 10-1 to 10-8 (use 0.5ml phage and 4.5ml broth each time).
Sterile nutrient broth (4.5ml per tube)
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Combining the bacteria and virus
NOTE: The following steps use sloppy agar. This will start to set if the
temperature is allowed to drop below 450C. These tubes must be
kept warm at all times.
2. Label each of the tubes of sloppy agar with a concentration ranging from
10-1 to 10-8.
Also label a tube as ‘stock’ and another tube as ‘control’.
3. Add 0.4ml E.coli B to every tube.
4. To each tube add 0.2ml appropriate phage dilution: remember to add
undiluted phage to the tube labelled ‘stock’ and to add nutrient broth to the
tube labelled ‘control’.
5. Rapidly mix by rotating the tubes between the palms of your hands.
Pouring the plates
6. Label the nutrient agar plates from 10-1 to 10-8, plus ‘stock’ and ‘control’
plates.
7. Pour the contents of the appropriate tube (see above) over the surface of a
nutrient agar plate so it forms an even layer.
8. Allow to set and then invert & incubate for 28-24 hours at 30oC.
Estimating the number of phage
Some plates will have so many plaques they will overlap each other, whilst
other plates (at higher dilutions) will have very few plaques. At least one plate
will have plaques in countable numbers.
9. Find the plate or plates with plaques in countable numbers.
10.Count the number of plaques & multiply this figure by the dilution factor to
obtain the concentration of the phage.
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