bioscienceexplained
Vol 3  No 2
134567
Andy Harrison, John Schollar
and Dean Madden
NCBE, The University of Reading, Earley Gate, Reading,
RG6 6BZ, UK
Investigating plant evolution
Amplification of chloroplast DNA using
the polymerase chain reaction
Aim
Students amplify an approximately 400 base pair
fragment of DNA from the chloroplast genome of
different plant species. Electrophoresis of the PCR
product may suggest evolutionary relationships.
Introduction and background information
Chloroplast DNA
Gene regions provide a molecular guide to evolution
A revolution in the garden
In 1998, an international group of nearly a hundred
plant scientists published a revolutionary proposal.
After more than two centuries of classifying plants on
the basis of their appearance, botanists began grouping
flowering plants according to similarities in their genetic
material, DNA.
The new system proved controversial and threw up
some surprises. For instance, botanists used to think
that the tropical papaya was a close relative of the
passionflower. According to the DNA evidence,
however, the papaya turns out to be a ‘cousin’ of the
cabbage, and roses are more closely related to nettles
than had previously been thought. A result of this
research is the reordering of the plants within botanic
gardens worldwide — a botanical ‘Year Zero’.
CORRESPONDENCE TO
Dean Madden
National Centre for
Biotechnology Education,
The University of Reading,
Earley Gate, Reading,
RG6 6BZ
The United Kingdom
[email protected]
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The reclassification project was led by Mark Chase at
the Royal Botanic Gardens, Kew (in London) and two
colleagues: Kåre Bremer of the University
of Uppsala in Sweden and Peter Stevens,
then at Harvard University in the USA.
The work took several years to complete.
The Angiosperm Phylogeny Group (APG),
as the team became known, reclassified the world’s
flowering plants into 462 families. Five years later,
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Vol 3  No 2
using improved molecular data and computer analysis,
the number of flowering plant families was reduced to
457.
Molecular methods
At the end of the eighteenth century the Swedish
naturalist Carl von Linné, known as Linnaeus, attempted
a systematic classification of plants. Since then,
botanists have grouped plants into the different families
according to similarities in their shapes and structures
— the number of petals, the shape and organisation of
a plant’s leaves, vascular system, embryo, seeds and so
on.
DNA evidence shows that the
tropical papaya (Carica papaya) is a
close relative of the humble cabbage
Traditional classification tended to
reflect evolutionary relationships, but
because natural selection produces
similar-looking solutions to life’s
challenges, identifying the true
underlying relationships was little more than
guesswork. Now, by applying the techniques of
molecular biology, a more accurate picture has begun
to emerge. How has this been done?
An essential task is to identify DNA sequences that vary
between species. Gene regions that are too variable will
not, however, reveal the genetic similarities that are
needed to create a family tree. Many gene regions have
relatively slow rates of change and are therefore
suitable for studying plant evolution, but these regions
are often rare and hard to work with. Fortunately
chloroplasts contain their own plentiful supply of DNA
that is ideal for evolutionary studies.
Chloroplasts have DNA
The Angiosperm Phylogeny Group
compared genes encoding a large
subunit of the enzyme ribulose
bisphosphate carboxylase/
oxygenase (Rubisco). This model
shows the eight large and eight
small subunits from which Rubisco is
made. The enzyme is needed for the
carbon-trapping process of
photosynthesis, and the gene for the
large subunit, codenamed rbcL, is
found in the DNA of chloroplasts.
Variations in the DNA sequence of
the gene enabled the APG scientists
to propose a new ‘family tree’ for
flowering plants.
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Soon after the rediscovery of Mendel’s Laws in 1900,
unexpected patterns of inheritance were noticed in
plants. These observations hinted that plants might
have extra genes that were not part of their
chromosomes. It was not until 1962, however, that
clear proof that chloroplasts contain their own DNA was
provided.
Between 10–100 chloroplasts are found in each green
plant cell, and each chloroplast contains 50–100 copies
of the chloroplast DNA (cpDNA). In total, 10–20% of a
plant’s DNA is found in the chloroplasts. Like that of
plasmids and mitochondria, the DNA of chloroplasts is
circular: typically it forms a ring about 120–150
thousand base-pairs (120–150 kilobases, or kb) in
length, encoding about 80 proteins. The cpDNA of
many plant species has now been sequenced.
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Vol 3  No 2
Outer membrane
Inner
membrane
Stroma
DNA within the
chloroplast
Lipid globule
Starch
granule
Stroma
lamellae
Granum
a stack of thyalkoids
In most species, cpDNA is
inherited through the egg cells
and does not undergo
recombination, which limits the
amount of variation found.
cpDNA does, however, show
nucleotide insertion, deletion
and substitutions.
High-resolution techniques are
required to detect small
variations within a plant
species, but it is possible, even
with simple equipment, to
detect larger differences in the
cpDNA between major plant
groups.
The internal structure of a chloroplast, showing the DNA.
The right genes
Several different genes within cpDNA have been used
by botanists to trace genetic relationships between
plants. The APG team originally used one of the Rubisco
genes, rbcL. cpDNA also contains genes that encode a
complete set of transfer RNAs (tRNAs). tRNA genes are
highly conserved — they have not changed much
during the course of evolution. Identical nucleotide
sequences can therefore be found in the cpDNA of most
higher plants. While there are common regions in the
genes of chloroplasts, the stretches of DNA between
them, especially in a non-coding stretch of cpDNA, can
vary greatly. Such regions have a higher frequency of
mutation and demonstrate relatively high rates of
evolutionary change, revealing genetic differences
between populations of plants that have been
genetically isolated for some time. It is these genetic
‘signatures’ that are examined in this practical
investigation.
Comparison of cpDNAs
This investigation uses the polymerase chain reaction
(PCR) to make copies of the highly-variable DNA that
lies between the conserved tRNA genes in chloroplasts.
The PCR produces millions of copies of DNA which can
be seen as bands on an electrophoresis gel.
Electrophoresis also allows the lengths of the variable
cpDNA from different species to be compared. The
distance moved on the gel by the DNA will vary
according to the lengths of the DNA fragments: shorter
fragments will travel further than longer DNA fragments
on the same gel.
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If the DNA from two different plants moves the same
distance on the gel, the sizes of DNA fragments made
by the PCR are the same for those plants. This may
indicate that they are genetically similar and are
therefore closely related. Species that yield differentlysized fragments may be more distantly related. From
these results it is possible to infer evolutionary
relationships.
The results obtained here are relatively crude, and they
are derived from a single variable region of DNA alone.
Many such regions must be used along with other
techniques, such as DNA sequencing, to build a more
accurate picture of evolutionary relationships.
Summary of the protocol
A Collect DNA from
fresh green plant
tissue
B Clean the sample
C Amplify the variable regions of the chloroplast DNA
by PCR, using water baths or a PCR machine
D Separate the amplified DNA
fragments by gel electrophoresis
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4
Related plants give
DNA fragments of
similar sizes
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bioscienceexplained
Vol 3  No 2
Equipment and materials
In the following protocol references are made to
NCBE´s equipment. However other and similar
equipment may be used.
Required by each student or working group
Equipment
 Pestle
 Punch for cutting 2 mm discs
 Micropipettes in 0–100 µL range or alternative
measuring method
 Pair of forceps
 DNA electrophoresis equipment and power supply
 Protective gloves (to protect your hands from the
DNA stain)
 Floating microcentrifuge tube holder, e.g., small
rectangle of plastic foam with four small holes cut
out of it with a cork borer — this is useful for holding
the microtubes on the bench or floating them in a
water bath, if you not use a PCR-machine
 Pencil
 Permanent marker pen for marking the tubes and
the lanes on the gel tank
 Access to three water baths maintained at 55, 72
and 94 °C, or a thermal cycler (PCR machine)
Materials
 Microcentrifuge tubes (1.5 mL)
 Micropipette tips
 Whatman® FTA® Plant Card
 FTA® Purification Reagent (~ 350 µL per plant
sample)
 TE-1 buffer (~ 350 µL per plant sample)
 PCR bead e.g., GE Healthcare ‘PuReTaq PCR ReadyTo-Go Beads’ (1 per plant sample)
 PCR primers and sterile distilled water, for each
sample as follows:
 10 µL of Primer 1:
5'–CGAAATCGGTAGACGCTACG–3'
(containing 5 pmol of DNA per µL)
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 10 µL of Primer 2:
5'–GGGGATAGAGGGACTTGAAC–3'
(containing 5 pmol of DNA per µL)
 ~6 µL of distilled water
 1 kb DNA ladder or ruler (or similar — only one is
required per gel))
 Bromophenol blue loading dye (~2 µL per sample)
 Electrophoresis (TBE) buffer solution
 Agarose gel (1.5%), made by dissolving the agarose
powder in diluted electrophoresis buffer
 DNA stain suitable for school use (e.g., 0.04% Azure
A in 20% ethanol)
Choosing plant species
Research and select your test
species carefully to obtain the most
interesting results
Using the techniques described in this text, we have
successfully amplified chloroplast DNA from a wide
variety of plant species. These include many food plants
that can be found on the shelves of supermarkets or
greengrocers. Although the PCR primers are known to
work for algae, liverworts, ferns, non-flowering and
flowering plants, the FTA® card extraction method
described here is not generally suitable for plants such
as pine or cabbage that have tough or fibrous leaves
that are difficult to crush.
Plants that are distantly-related will tend to give DNA
fragments that differ most in size. It would be
interesting to compare, for example, orchids (which are
recently-evolved) with Ginkgo (which is similar to
plants found in the fossil record 225 million years ago).
Guidance for selecting plant species is provided by the
web-based Tree of Life project: http://tolweb.org/ and
by the work of the Angiosperm Phylogeny Group
http://www.mobot.org/MOBOT/research/APweb/
In our (limited) tests, none of the following plants gave
reliable amplification with the FTA® card method:
 ‘Cress’ (Brassica napus or Lepidium sativum
seedlings);
 Cabbage and broccoli (both plants are the same
species, Brassica oleracea);
 Thyme (Thymus vulgaris) or basil (Ocimum
basilicum);
 Cucumber (Cucumis sativus) or courgette (Cucurbita
pepo) skin — but we did not try the leaves.
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Figure 1.
Place the leaf
tissue on the card
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The practical protocol
A
Collecting DNA from plants
1.
Fold back the cover of the FTA® Plant Card. Take
a fresh leaf or piece of plant material and place it
on the card. Ensure that the plant material does
not extend outside one of the four boxes printed
on the card (Figure 1).
2.
Close the cover over the plant material and, using
a pestle, squash the leaf onto the card until
moisture has soaked through to the back of the
card (Figure 2). Try not to allow the sample to
spread outside the box printed on the card as this
can contaminate adjacent samples.
3.
Throw away the plant material.
4.
Write the plant’s name in the appropriate ‘Sample
ID’ box on the cover of the FTA® Plant Card
(Figure 3). The cover should be labelled after the
plant material has been squashed onto the card,
as the pestle can rub the writing off the front
cover.
5.
With the cover folded back, allow the card to dry
at room temperature for a minimum of one hour
(Figure 3). Important: Do not heat the card as
this may fix inhibitors of the PCR onto the
card.
6.
Once dry, the card can be stored indefinitely,
sealed in a foil bag with a desiccant sachet at
room temperature.
Ensure that it fits
within a box
Figure 2.
Close the cover and
squash the leaf
OPTIONAL STOPPING POINT
Write the
plant’s name
in pencil on
the cover
Figure 3
Allow the card to dry for 1 hour
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Notes
FTA® card is filter paper impregnated with SDS,
Tris/EDTA (TE) buffer, and uric acid. SDS (a
detergent) breaks open the plant cells. EDTA
binds to metal ions that are required as cofactors by the enzymes that degrade DNA, so
that the enzymes cannot work. Tris (a weak
base) is needed to ensure the chelating action of
the EDTA. The uric acid protects the DNA from
degradation by free radicals.
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Figure 4.
Cut discs using a punch
Figure 5.
Push the disc
from the
punch using
some plastic
‘wire’
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B
Cleaning the DNA sample
1.
Place a cutting board between the dry FTA® Plant
Card and the backing. Place the tip of the punch
over the area to be sampled i.e., an area coloured
green by chlorophyll. Press down firmly on the
punch and rotate to remove a paper disc (Figure
4). To avoid cross-contamination, do not take a
disc close to the edge of the box if the extracted
sample has overlapped with another sample in an
adjacent box.
2.
Transfer the small disc directly into a clear,
colourless 1.5 mL microcentrifuge tube by pushing
it out of the punch with the plastic ‘wire’ provided.
Label the tube to indicate the species used (Figure
5). Important: If the punch is to be used for
a second sample, first clean the metal cutter
by removing a disc of paper from an area of
the card that does not have plant material
applied to it. Discard this disc. This process
should prevent contamination of subsequent
samples.
3.
Using a 1 mL syringe fitted with a graduated
yellow tip, add 100 µL of Purification Reagent to
the tube containing the paper disc (Figure 6a). Do
not draw liquid into the syringe body.
4.
Close the tube and flick it several times to wash
the disc. Flick every 30 seconds for 2 minutes
(Figure 6b). Purification Reagent contains a
detergent, which helps to wash cell debris,
including PCR inhibitors, from the disc.
Important: If the disc is not washed
thoroughly the PCR may fail.
5.
Using the same yellow tip as in Step 3, remove
the Purification reagent from the tube. Try to
remove as much of the froth from the tube as you
can, but do not draw liquid or froth into the body
of the syringe.
6.
Wash the disc again by repeating Steps 3 to 5,
Wash the disc twice in
Purification Reagent
100 µL
Add 100 μL of Purification
Reagent
Flick to mix
Remove the liquid
REPEAT
Figure 6a
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Figure 6b
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Vol 3  No 2
using the same yellow tip for all operations.
7.
Now, use a new yellow tip attached to the 1 mL
syringe. Add 100 µL TE-1 buffer to the tube
(Figure 7a). TE-1 is very dilute Tris-EDTA buffer.
8.
Close the tube and flick it to wash the disc. Flick
every 30 seconds for 2 minutes (Figure 7b). This
removes detergent that was in the purification
reagent from the FTA® disc.
9.
Using the same yellow tip, remove and discard
the TE-1 buffer from the tube.
10.
Wash the disc again by repeating Steps 7 to 9,
using the same yellow tip.
100 µL
Wash the disc twice in
TE-1 buffer
Add 100 μL of TE-1 buffer
Flick to mix
Remove the liquid
REPEAT
Figure 7a
Figure 7b
11.
Go on to conduct the PCR, or …
12.
… alternatively, the disc can be dried in the tube
by warming to ~ 50 °C. You can use a hair-drier,
radiator or incubator to speed up this process, but
don’t ‘cook’ the DNA excessively. Once the disc is
dry, place the closed tube in a sealed foil bag with
a desiccant sachet and store it at room
temperature until it is needed.
OPTIONAL STOPPING POINT
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Testing for contamination
Because the PCR is so sensitive, it is
important to check whether contamination
with unwanted plant DNA has occurred. In
addition to the disc for the plant species,
take a disc from an area of the card that does not
deliberately have plant material applied to it. Treat
this disc in the same way as the ‘plant’ disc, washing
it and so on as though it were a disc impregnated
with plant material. This test will tell you whether
unwanted DNA has contaminated the punch and
samples taken with it. Ideally, each person who sets
up a PCR should do this test, but because the
reagents are costly, only one test is usually run per
class or group.
In addition, in ideal circumstances, you should also
test whether any of the PCR reagents have become
contaminated with DNA. This can be done by setting
up a reaction with the two primers and water but with
no paper disc. It can be assumed, however, that all
reagents supplied are free from contamination, so
this test should be unnecessary.
C
Amplifying the chloroplast DNA
1.
Using a microsyringe fitted with an unused white,
graduated tip for each reagent, add the following
to a 0.5 mL microcentrifuge tube containing a PCR
bead (Figure 8):

10 µL of Primer 1 (containing 5 pmol of DNA
per µL) 5'–CGAAATCGGTAGACGCTACG–3'

10 µL of Primer 2 (containing 5 pmol of DNA
per µL) 5'–GGGGATAGAGGGACTTGAAC–3'

4 µL of distilled water (add 6 µL if a dry FTA®
disc is used)
Figure 8
Primer 1 Primer 2 Water
10 µL
10 µL 4 or 6 µL
PCR ‘bead’, containing:
– Taq polymerase
– Buffer
– dNTPs
– MgCl2
Primer 1: 5'–CGAAATCGGTAGACGCTACG–3'
Primer 2: 5'–GGGGATAGAGGGACTTGAAC–3'
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Ensure that the tip does not touch the PCR bead.
If it does, the bead may stick to the tip. Instead,
place the tip against the side of the tube above
the bead. This will leave a drop of liquid on the
side of the tube. The liquid will then run down to
the bottom of the tube, causing the PCR bead to
dissolve.
Figure 9.
Use forceps to
add a disc with
plant DNA
Figure 10.
Label the tube
Figure 11a.
A PCR thermal
cycler
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2.
Close the tube. Flick the bottom of the tube gently
to mix the contents and to help the PCR bead
dissolve.
3.
Spin the tube briefly in a microcentrifuge or tap
the tube firmly on the bench to collect the liquid
at the bottom of the tube.
4.
Using a pair of forceps, place the FTA® disc with
the plant DNA into the tube containing the PCR
reagents (Figure 9). Important: Ensure that
the disc is submerged in the liquid in the
tube.
5.
Close the tube and label the top with the name of
the plant used and your initials (Figure 10).
6.
Place your tube in a thermal cycler, or if using
water baths, into a foam floater, along with any
tubes from others in your group or class.
To conduct the PCR, use the thermal cycler (Figure
11a)...
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Vol 3  No 2
Figure 11b
30 seconds
30 seconds
Repeat this
3-step cycle
30 times
30 seconds
If using water baths
Take care not to scald yourself when carrying out the PCR.
To maintain 94 °C it may be necessary to place a lid on
W the water bath when it is not in use.
rite the plant’s
name in pencil
on the cover
7.
(cont) ... or transfer the floater between three
water baths to provide temperatures as follows
(Figure 11b):
a. 94 °C for 2 minutes
(ensures that the template DNA is singlestranded)
b. 94 °C for 30 seconds
c. 55 °C for 30 seconds
d. 72 °C for 30 seconds
Repeat b-d 30 times
e. 72 °C for 2 minutes
(ensures that the DNA fragments are fully
extended).
8.
Go on to conduct the gel electrophoresis or …
9.
… alternatively, store the amplified DNA solutions
in a freezer at –20 °C until they are required.
OPTIONAL STOPPING POINT
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Caution!
Hot agarose
can scald
Figure 12
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D
The volumes and figures in the following description is
for NCBE´s electrophoresis tank, but other
electophoresis equipment can of course also be used.
I
Preparing the agarose gel
1.
Use a microwave oven or a boiling water bath to
melt some agarose gel (1.5% made up in TBE
buffer). Ensure that no lumps or fibres remain in
the molten agarose — you can see these by
holding the bottle up to the light. Once it has
melted, stand the molten agarose in a sealed
container in water bath at 55–60 °C until it is
needed.
2.
Place the electrophoresis tank on a level surface,
where you can leave it undisturbed for the next
20–30 minutes. This is necessary because if the
gel sets at an angle, the DNA fragments will not
run evenly through the gel.
3.
Slot a 6-toothed comb in place at one end of the
tank (Figure 12). Ensure that the frosted panel
faces the far end of the tank (see Figure 12).
4.
Pour ~12 mL of molten agarose into the tank so
that it fills the central cavity and flows under and
between the teeth of the comb. Add just enough
agarose so that the liquid is level with the top of
the two ridges in the tank, and does not bow
upwards (Figure 12). If you accidentally spill
liquid into the areas outside the two ridges, do
not worry — you can remove the agarose later,
when it has set.
5.
Leave the tank undisturbed until the liquid has
set. Agarose is opaque and looks slightly milky
when the gel is set.
6.
While the gel is setting, cut two pieces of carbon
fibre tissue, each about 42 mm  22 mm (Figure
13). These will be the electrodes at either end of
the electrophoresis tank. Put the electrodes to
one side until you have loaded the gel.
Frosted panel
on this side
Moulten agarose
55-60 °C
Carbon
fibre tissue
42 mm
22 mm
Cut two
electrodes
Figure 13
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DNA gel electrophoresis
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Figure 14
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II
Loading the gel
1.
Pour slightly more than 10 mL of TBE buffer
solution into the electrophoresis tank. The liquid
should cover the surface of the gel to a depth of
2–3 mm and flood into the areas that will hold the
electrodes.
2.
Very gently ease the comb from the gel. As you
do so, the buffer solution will fill the wells that
formed (Figure 14). Take care not to tear the
wells as you remove the comb.
3.
Put the tank where you are going to ‘run’ it,
where it will remain undisturbed.
4.
It is easier to see what you are doing next if the
tank is placed on a dark surface, such as a piece
of black card. Alternatively, a strip of black tape
can be stuck onto the bottom of the tank beneath
the wells. Put a clean white tip on the
microsyringe. Add 2 µL of bromophenol blue
loading dye to the tube containing the DNA you
wish to load. Mix the dye into the DNA sample
thoroughly, by drawing the mixture up and down
in the microsyringe. Pipette about 10 µL of the
loading dye and DNA mixture into one of the
wells, holding the tip above the well but under the
buffer solution (see Figure 15). Take great care
not to puncture the bottom of the well with the
microsyringe tip.
Once the gel has set, pour on
TBE buffer, then lift the comb
out gently
Figure 15
TBE buffer
solution
Label the end of
the tank to show
the contents of
each well
5.
2-3 mm depth of
buffer over the gel
Black card under the
tank reveals the wells for
loading
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Make a note of which DNA
you have put into the well
(e.g., write on the side of
the tank with a marker pen).
6.
Repeat Steps 4 and 5 with each DNA sample.
Remember to use a new tip for each sample, to
avoid cross-contamination.
7.
Load 10 µL of diluted DNA ‘ruler’ into one of the
wells.
8.
On one gel, also load 10 µL of the test sample
that had no plant DNA added to the reaction
mixture.
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Figure 16
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III
‘Running’ the gel
1.
Fit one electrode at each end of the tank as
shown in Figure 16. Join the electrodes to
batteries using wires with crocodile clips (Figure
17). Sufficient batteries (in series) should be used
to give a total voltage of no more than 36 volts.
Ensure that the positive terminal of the battery is
connected to the electrode furthest from the
wells. (With safely covered electrophoresis tanks
higher voltage can be used.)
2.
Check that contact is made between the buffer
solution and the electrodes (with care, you can
add more buffer if necessary). Place the comb
over the tank to reduce evaporation during the
electrophoresis. Leave the gel to ‘run’,
undisturbed, for several hours. The time required
will vary according to the voltage.
Electrodes
Figure 17
Voltage
27
30
Direction of
DNA movement
Time to run gel
3½ hours
2 hours
APPLY NO MORE THAN 36
VOLTS!
Serious or lethal electrical shock
may occur if you connect the
equipment directly to a mains
electricity supply
3.
Disconnect the batteries once the blue loading
dye is within 10 mm of the end of the gel. The
DNA fragments (400 base pairs in length) will be
just behind the loading dye. Important: If you
leave the batteries connected, the DNA will
run off the end of the gel and be lost!
4.
Rinse the crocodile clips in tap water and dry
them thoroughly to prevent corrosion.
Notes
Ions in the TBE buffer used in and above the gel
conduct electricity. This buffer is alkaline. Under
alkaline conditions the phosphate groups of the DNA
are negatively charged and therefore the DNA moves
towards the positive electrode (anode) when a current
is applied. EDTA in the buffer ‘mops up’ or chelates
divalent cations. This helps to prevent damage to the
DNA as such ions are required as co-factors by DNAdegrading enzymes. The loading dye does not affect
the DNA, but moves through the gel slightly ahead of
all but the smallest of DNA fragments.
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Positively-charged Azure A
binds to the negatively-charged
phosphate groups of the DNA
Figure 18
Stain for only
4 minutes
Vol 3  No 2
IV
Staining the DNA
1.
Remove and dispose of the electrodes. Pour off
the buffer solution. The buffer may be retained
and re-used several times, but it will not last
indefinitely.
2.
Put on some plastic gloves to prevent the stain
from touching your skin.
3.
Pour about 10 mL of staining solution (0.04%
Azure A in 20% ethanol) onto the surface of the
gel (Figure 18).
4.
Leave it for exactly 4 minutes, then return the
stain to a bottle for re-use. (Like the buffer, the
DNA stain can be re-used several times before
you need to replace it. However, you may find
that older stain needs to be left on the gel for a
little longer than 4 minutes.)
Figure 19
1 kb ’ruler’
Wells
Sizes in base
pairs
Region with
DNA bands
Loading dye
5.
Smaller
fragments may
run off the gel
Rinse the surface of the gel with water 3 or 4
times, but do not leave any water on the gel, as
this will remove stain from the top of 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
(Figure 19). The best results will be seen if the gel
is left to ‘develop’ overnight. Put the tank in a
plastic bag. This prevents the gel from drying out.
Notes
It is not necessary to destain the gel if you follow this
procedure.
Stained gels may be stored indefinitely in a sealed
plastic bag, kept in a dark, cool place such as a
refrigerator.
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Safety
FTA® cards and PCR reagents
FTA® paper is non-toxic to humans and hypo-allergenic.
The PCR reagents are also non-toxic. Cleanliness is
important to prevent cross-contamination and ensure
success, so dirty tubes and tips should not be re-used.
Used plastic tubes and tips, which are made of
polypropylene, can be disposed of in the normal waste.
Hot water baths
If you use water baths for the PCR (rather than an
automated thermal cycler), take care not to scald
yourself when transferring tubes between them. Use
forceps or a clothes peg to hold the foam floater and/or
wear heat-resistant gloves to protect your hands from
the hot water and steam.
Agarose gel
If a microwave oven is used to melt the agarose gel,
ensure that the gel is placed in an unsealed container.
A boiling water bath or hotplate may be used instead,
but the gel must be swirled as it melts to prevent
charring. The use of a Bunsen burner to melt agarose is
not recommended.
CAUTION!
Molten agarose can scald and
it must be handled with care,
especially just after it has
been heated in a microwave
oven.
TBE buffer (Tris-Borate-EDTA)
When used as directed, this buffer presents no serious
safety hazards. Spent buffer can be washed down the
drain.
Loading dye (Bromophenol blue)
When used as directed, this loading dye presents no
hazard. Used loading dye can be washed down the
drain.
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Electrode tissue
The carbon fibre electrode tissue may release small
fibres, which can cause skin irritation if you handle the
tissue a lot. Wear protective gloves if you find the
tissue unpleasant to handle. The fibres are too large to
enter the lungs however, so it is not necessary to wear
a face mask. The fibres are soluble in body fluids and
are completely biodegradable.
Electrical supply
The gel electrophoresis equipment was designed to be
used with direct current at low voltages (≤36 volts)
with dry cell batteries. Under no circumstances
should this voltage be exceeded, as the live
electrical components are not isolated from the
user.
CAUTION!
Serious or lethal electrical
shock may occur if you connect
the equipment directly to a
mains electricity supply.
If other electrophoresis equipment than NCBE´s is used
then instructions for that equipment should be followed.
DNA stain (Azure A)
The concentrated DNA stain solution is flammable and
it must be kept away from naked flames. The stain is
Azure A, which when diluted as directed, forms a
0.04% solution in 20% ethanol. At this concentration it
presents no serious safety hazard, although care should
be taken to prevent splashes on the skin or eyes e.g.,
wear protective gloves and safety glasses. Used
solution can be diluted with water and washed down
the drain.
Ethical and other concerns
This practical exercise raises no ethical considerations.
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Recipes
TBE (Tris-Borate-EDTA) buffer
Used for making up agarose gel and as the
electrophoresis buffer
Makes 1 litre of 10 concentrate
May be stored indefinitely at room temperature
Ingredients
1 g sodium hydroxide (MW 40,00)
108 g TRIS-base (MW 121,10)
55 g boric acid (MW 61,83)
7,4 g ethylene diamine tetraacetic acid (EDTA,
disodium salt, MW 372,24)
Directions
Add the ingredients to 700 mL deionized or distilled
water.
Stir to dissolve. Make up to 1 litre with deionised or
distilled water.
CAUTION!
Take care not to inhale any Tris base or EDTA powder.
Wear a protective mask over your mouth and nose.
Both of these chemicals are classified as irritants and
can cause a reaction upon skin contact or inhalation.
Once made up, 1 volume of this TBE concentrate
should be diluted with 9 volumes of distilled water
before use.
Important!
Do not use TRIS-HCl to prepare this buffer.
TRIS-base must be used.
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Azure A
Stain for nucleic acids
Makes 50 mL of 2 concentrate
May be stored indefinitely at room temperature
Ingredients
0.08 g Azure A (MW 291.8)
50 mL ethanol (40% aqueous solution)
Directions
Add the Azure A powder to 50 mL of 40% ethanol. Stir
to dissolve.
CAUTION!
The concentrated DNA stain is flammable and must not
be used or stored near naked flames or other sources
of ignition. The stain bottle must be kept closed to
prevent evaporation of the solvent.
When diluted to its working concentration (0.04% in
20% ethanol), the stain presents no serious safety
hazard, although care should be taken to avoid
splashes on the skin or eyes e.g. wear protective gloves
and glasses. Used stain may be diluted with water and
washed down the drain.
FIRST AID (Azure A powder)
EYE CONTACT
Flush eyes with copious amounts of water for at least
15 minutes. Seek medical attention.
SKIN CONTACT
Wash the skin with soap and plenty of water. If
irritation occurs seek medical attention.
INGESTION
Provided the person is conscious, wash out the mouth
with water. Induce vomiting. Seek medical attention.
NOTE
Once made up, this Azure A concentrate should be
diluted with an equal volume of distilled water before
use.
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Agarose gel
For separating small fragments of nucleic acids
Makes 100 mL of 1.5% agarose
May be stored indefinitely at room temperature
Ingredients
1.5 g electrophoresis grade agarose
100 mL TBE buffer
Directions
Add the agarose powder to the TBE buffer. Heat in a
boiling water bath or microwave oven to melt the
agarose. Less than a minute at full power in a 940 watt
oven is sufficient to melt 100 mL of gel. The container
used to hold the molten agarose must not be sealed,
but lightly covered with plastic film that has been
punctured with one or two small holes. Swirl the gel
halfway through the heating cycle to ensure that it is
thoroughly mixed.
Once molten, the agarose solution can be kept in this
state at 55–60 °C in a water bath. Ensure that the
agarose solution is mixed thoroughly before casting
gels.
CAUTION!
Hot, molten agarose can scald and so it must be
handled with care. It is advisable to let the molten
agarose cool until it is comfortable to handle before
pouring the gel.
Bromophenol blue loading dye
Used for loading DNA fragments into the
electrophoresis gel
Makes 100 mL
May be stored indefinitely at room temperature
Ingredients
0.25 g Bromophenol blue (MW 669.96)
50 g Sucrose (MW 342.30)
1 mL 1 M TRIS (pH 8, MW 121.10)
Directions
Add the ingredients listed above to 60 mL of deionised
or distilled water.
Stir to dissolve. Make up to 100 mL with deionised or
distilled water.
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Preparation and timing
This investigation is a complex multi-step process.
Careful planning is therefore necessary to fit it into a
normal series of school lessons or a single practical
session. The whole procedure, including drying the
FTA® cards and manual PCR but excluding the
electrophoresis, takes about 3 hours. The duration of
the electrophoresis depends upon the equipment and
voltage used — this can be as little as 30 minutes (at
90 volts) or as slow as three and a half hours (at 27
volts), using the NCBE equipment. Larger gels, using
other equipment, will take longer to run.
As a rough guide, the timing of this protocol is as
follows:
Extracting the DNA
onto FTA® cards:
10 minutes
Drying time:
60 minutes
STOP POINT
Cleaning the DNA samples:
20 minutes
STOP POINT
Preparing the PCR reagents:
10 minutes
PCR manual cycling:
50 minutes
PCR thermal cycler:
~ 90 minutes
STOP POINT
Preparing and loading the gel:
30 minutes
Electrophoresis:
30 minutes
- 3.5 hours
Staining the gel:
10 minutes
To save time, gels may be cast in advance. Cast gels
may be stored in a refrigerator for several days before
use, if they are covered in TBE buffer and placed in a
sealed plastic bag.
It is also possible to stop the procedure after running
the gel, although this is not recommended. The gel may
be stained up to 24 hours after running it, but if it is
stored for longer periods, the DNA will diffuse into the
gel and the bands will be less discrete. Unstained gels
should be stored covered in TBE buffer in a sealed
plastic bag in a refrigerator.
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Troubleshooting
Micropipette practise
Students’ ability to dispense precise volumes using
micropipettes will improve with experience. It is
therefore a good idea to practise transferring liquids
(e.g. food colouring) before starting work with
expensive DNA, enzymes and other reagents.
Loading the gel
A steady hand is required to load gels. Practice gels
may be cast from cheaper agar (use a 1% solution)
rather than agarose. When practising, use water rather
than buffer over the gel. Practice gels can be washed
out and re-used several times.
Notes
Agarose, dissolved in diluted buffer, not water,
must be used for the ‘real’ gels with DNA.
Melting agarose
Ensure that the container used to prepare the agarose
gel is clean. Tiny flecks of dust will not affect the way
the gel runs, but they can prove a nuisance when you
are trying to see faint bands in the gel.
For convenience, dissolve and melt the agarose using a
microwave oven. Less than a minute at full power in a
940 watt oven is sufficient for 100 mL of gel. The
container (flask or beaker) used to hold the molten
agarose must not be sealed, but covered lightly with
plastic film that has been punctured with one or two
small holes. Swirl the gel half way through the heating
cycle to ensure that it is mixed thoroughly.
Agarose gel can be prepared using a hotplate. If this is
done, the gel must be swirled as it melts, to prevent
charring. Better still, stand the container of agarose in
a saucepan of boiling water. The use of a Bunsen
burner to melt agarose is not recommended.
Once melted, the gel may be kept in a molten state by
standing the container in a water bath at 60 °C until
the gel is needed. Take care when handling the molten
gel; it will be very hot, and can scald. The gel should be
allowed to cool before it is poured.
Gel takes too long to run
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If, after the first 20 minutes, the loading dye does not
seem to have moved and bubbles are not visible at the
cathode, check the electrical contacts between the
batteries and the electrodes. Ensure that there is
enough buffer above the gel, but not so much that
most of the current passes through the buffer solution
rather than the gel. Remember than during long runs
(e.g., overnight) or in a warm environment liquid may
evaporate from the buffer. The tank should be covered
to reduce such evaporation.
There’s no DNA on the gel
Assuming the PCR has worked correctly, there are two
or three common causes of this problem.
Have you loaded sufficient DNA into the well?
Has the well been punctured by accident so that the
DNA sample has been lost?
Has the DNA sample been mixed thoroughly with the
loading dye?
The DNA bands are very faint
There are several potential causes of this problem.
Assuming that the PCR has worked, a common cause is
the failure to mix the DNA with the loading dye. To
ensure that this is done, always draw the DNA solution
up and down in the microsyringe tip a few times.
Has sufficient DNA sample been loaded into the well?
Are you using the stain correctly?
Once you have applied the Azure A to the gel, poured
off the stain, then rinsed off the excess, take care not
to leave any water on the gel. If necessary, wipe over
the gel gently with a finger tip to disperse beads of
water. The blue stain that starts in the upper layer of
the gel can re-dissolve even in small drops of water left
on the surface. ‘Soaking’ away the stain into water
drops on the surface prevents it from passing down
through the gel to stain the nucleic acid.
Under no circumstances should you attempt to de-stain
the gel. Although this is necessary with methylene blue
(to remove ‘background’ stain), the method using
Azure A has been specifically devised to avoid the need
for de-staining. Unlike the bands on gels stained with
methylene blue, those revealed by Azure A should not
fade for several months.
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The bands on the gel are very close together and
are difficult to tell apart
There are several possible causes of this. Provided the
buffer has not evaporated excessively while the gel was
running (slowing the electrophoresis down), it may be
that the concentration of agarose you are using is
wrong. Always make up more gel than you need to
ensure that its concentration is accurate. There is a
very significant difference between say, a 0.8% and a
1.5% agarose gel — only a slight error in weighing a
small amount of agarose powder can produce
inaccuracies of this scale. The concentration of agarose
you should use with this PCR protocol is 1.5% (mass to
volume).
Scope for open-ended investigations
Students may select any plants they wish for this
investigation, e.g., plants which are thought to have
evolved recently compared with those that have
relatives in the distant fossil record. Note that tough or
fibrous tissue such as pine needles and plant stems
may not be easy to process using the FTA® card
method.
If sequence data is available for the plant species, online bioinformatics tools such as Clustal-W may be used
to construct phylogenetic trees:
http://www.ebi.ac.uk/clustalw/
It is sometimes possible to distinguish between closelyrelated species (such as barley and wheat) by
restricting the amplified DNA. See, for example,
Streicher, H. and Brodte, A. (2002).
NCBE has tested and obtained good results with:
Nasturtium
Chive
Spinach
Tatsoi
Rocket
Corriander
Spring onion
Mange tout
Watercress‫٭‬
Parsley
Scenedesmus
quadricauda
Brussels sprout
Chlorella
vulgaris
Selenastrum
gracile
Red chard
Mizuna
Sweet and chilli
peppers (skin of
green varieties)
‫٭‬
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Watercress consistently gives a larger PCR product
25
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Vol 3  No 2
Suppliers
All of the materials required for this investigation may
be purchased from laboratory supply companies. FTA®
cards may be purchased from distributors of Whatman®
products worldwide (www.whatman.com). PCR beads
are available from GE Healthcare (formerly Amersham
Biosciences), and other suppliers.
The National Centre for Biotechnology Education (UK)
supplies a kit to schools for carrying out this protocol
(www.ncbe.reading.ac.uk).
Disposal of waste and recycling of
materials
All of the waste from this procedure can be disposed of
without special precautions. Although tips,
microcentrifuge tubes etc. can be cleaned, autoclaved
and re-used, this is not recommended due to the
sensitive nature of the PCR reagents, which may be
adversely affected by contaminants.
Storage of materials
Most of the reagents should be stored dry, at room
temperature. PCR primers may need to be refrigerated
or stored at -20 °C (refer to the supplier’s instructions).
Figure 20
Wells
Specimen results
Region with
DNA bands
Loading dye
Figure 20 shows a typical result.
Other sources of information
Mullis, K.B. (1990) The unusual origin of the
polymerase chain reaction. Scientific American 262,
36–43.
Making PCR. A story of biotechnology by Paul Rabinow
(1996) The University of Chicago Press. ISBN: 0 226
70147 6.
Willmott, C. (1998) An introduction to the polymerase
chain reaction. School Science Review 80, 49–54.
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Vol 3  No 2
References
This practical protocol is based on:
Taberlet P., et al. (1991) Universal primers for the
amplification of three non-coding regions of chloroplast
DNA. Plant Molecular Biology 17, 1105–1109.
An extension of this procedure, using the restriction
enzyme Dde1 to restrict the PCR product, allowing
identification of species is described in:
Streicher, H. and Brodte, A. (2002) Introducing
students to DNA: Identifying nutritional plants in a
simple tried and tested laboratory experiment.
Biochemical Education 30, 104–105.
Acknowledgements
Molecular structure data
The molecular images were created using data from the
Protein Data Bank: www.rcsb.org/pdb
The data for Taq polymerase was published in: Eom,
S.H., et al. (1996) Structure of Taq polymerase with
DNA at the polymerase active site. Nature 382, 278–
281. [Protein Data Bank ID: 1TAU].
The Rubisco molecule is from: Taylor, T.C., and
Andersson, I. (1997) The structure of the complex
between rubisco and its natural substrate ribulose 1,5bisphosphate. Journal of Molecular Biology 265, 432–
444. [PDB ID: 1RCX]
The software used to produce the molecular model
images was VMD (Visual Molecular Dynamics) which
can be downloaded free-of-charge from:
www.ks.uiuc.edu/Research/vmd/
PCR protocol
The use of universal primers for amplification of
chloroplast DNA was originally brought to our attention
by Kenny Hamilton from Breadalbane Academy in
Aberfeldy, Scotland. Kenny developed a preliminary
protocol while on a Royal Society of Edinburgh Teacher
Fellowship at the University of Edinburgh. Craig
Simpson of the Gene Expression Programme at the
Scottish Crop Research Institute provided invaluable
advice on the design of the original protocol as well as
laboratory facilities.
Andy Harrison at the National Centre for Biotechnology
Education at The University of Reading refined and
simplified the original protocol, and introduced the use
of FTA® cards for the extraction of plant DNA. He also
optimised the PCR process so that it could be done
more easily and quickly in the school laboratory.
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Appendix 1: Making a haystack from a needle
The polymerase chain reaction
A very useful process
The polymerase chain reaction (PCR) has been likened
to a ‘genetic photocopier’. From a small amount of
biological material millions of copies of a chosen section
of its DNA can be made quickly, accurately and
automatically.
An active site
here strips
the primers
from the DNA
DNA is made at
this active site
The breakthrough that made the PCR
into a convenient laboratory technique
was the isolation of DNA polymerase
from Thermus aquaticus, a bacterium
found in hot water springs. This
enzyme (known as Taq polymerase) is
stable at high temperatures and can
therefore withstand the rigours of the
reaction. In the computer-generated
structure shown here, -pleated sheets
are coloured yellow, while the -helices
are magenta. A small fragment of the
DNA that is being made is shown as a
space-filling model.
Although it was first devised for carrying out medical
diagnostic tests, today the PCR has a wide range of
uses ranging from forensic genetic fingerprinting and
archaeological investigations to tracing the mating and
feeding habits of animal species.
A method called ‘sexual PCR’ or ‘DNA shuffling’ enables
protein engineers to develop better enzymes for
industrial use by mimicking the processes of evolution.
Without a clever variant of the PCR called ‘cycle
sequencing’ the Human Genome Project would have
been almost impossible — as would much of the
molecular biology that is carried out in laboratories
around the world. It is hardly surprising therefore that
the PCR won for its inventor, Kary Mullis, a Nobel Prize
for Chemistry in 1993. What is more surprising is the
PCR’s unusual origin.
A drive through the woods
The road to the PCR began on a moonlit evening in
April 1983. In Kary Mullis’s account of his invention, he
describes how its two key features occurred to him
while driving to his weekend retreat in California. Twice
Mullis had to pull over and stop the car: once to
calculate the number of copies of DNA that the process
might generate; and a second time to sketch a
diagram, showing how it might duplicate a specific
stretch of DNA.
The idea was so simple that at first people refused to
believe that it would work. No one could believe that it
hadn’t been tried before and found to be impractical for
some reason. After Mullis’s initial work, his colleagues
at the Cetus Corporation, one of the first modern
biotechnology companies, carried out extensive
research to refine the method and eventually the
process was patented.
Mullis was paid a bonus of US$ 10,000 for his
invention. The PCR has become so important however,
that Cetus was later able to sell the rights associated
with the process to Hoffman-La Roche for US$ 300
million.
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How the PCR works
The modern method for PCR (a slightly modified version
of Mullis’s original idea) operates as follows:
The components
 Double-stranded DNA (the ‘template’) is extracted
from the biological sample under study.
 Two single-stranded ‘primers’ are made that flank
the specific stretch of DNA to be copied. These are
generally 15–30 bases long, and are complementary
to (in other words, they pair up with) the 5' end of
each target DNA strand.
 The template DNA is mixed with the primers.
 Equal proportions of deoxyribonucleoside
triphosphates (dNTPs) are added. dNTPs are the
units from which DNA is made.
 Heat-stable Taq DNA polymerase (obtained from the
hot-water bacterium Thermus aquaticus) is added to
the reaction mix, together with Mg2+ ions that are
required as a co-factor by the enzyme. This enzyme
strings together dNTPs to make new DNA.
All of the components needed for the PCR (apart from
the primers and the template DNA) can be dried into a
small pellet in the correct proportions. Then, all that
needs to be done is to add the primers and template
DNA to a tube containing the pellet. This removes the
need to dispense very small volumes of numerous
reagents and improves the reliability of the process.
The process
Once the chemicals necessary for making DNA have
been mixed, the PCR has three steps which are
repeated 20–40 times:
Denaturation
The double-stranded DNA is split into two singlestranded templates by heating it to 94–98 °C.
Annealing
The mixture is cooled to 50–65 °C. The primers bind
to the complementary strands of DNA by basepairing.
Extension
Heating to 72 °C (the optimum temperature for Taq
polymerase) encourages the synthesis of new DNA
strands alongside the templates. This step doubles
the number of DNA molecules present.
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The cycle of heating and cooling is now repeated. With
each cycle (lasting about 2 minutes) the number of
copies of the DNA is doubled. After 25 cycles more than
a million copies of the target DNA will have been made.
The first attempts at PCR were made using water baths
maintained at three different temperatures. It was not
long however, before the process was automated — the
first PCR machine was known affectionately as ‘Mr
Cycler’. To reduce evaporation of the small volume of
liquid as it is repeatedly heated and cooled, a modern
PCR machine has a heated plate over the tubes.
The PCR is a very sensitive method and in theory it can
amplify a single DNA molecule. Scrupulous precautions
are therefore often necessary to ensure that stray DNA
molecules stay out of the reaction mixture.
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The sequential steps in the PCR
50–65 °C
Primers anneal to complementary
sequences of bases
72 °C
Taq DNA polymerase extends
the DNA strands
94 °C
Double-stranded DNA
is denatured
Taq polymerase
Non-target DNA
Target DNA
Primers
Start
First cycle
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Second cycle
31
Third cycle
Fourth cycle
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Vol 3  No 2
Appendix 2: Measuring small volumes
Three different devices are provided in this kit so that
you can dispense microlitre volumes of liquid accurately
Syringes with graduated tips
White
graduated
tip
Soft rubber
tubing
Two types of syringes — conventional 1 mL syringes
and narrower ‘microsyringes’ — can be fitted with
Hold here
disposable graduated tips and used to measure small
Do not touch
volumes. The white microsyringe tips are marked at 2
the point
and 10 microlitres (μL). The larger yellow tips can be
fitted to a 1 mL syringe and used to dispense volumes
10 μL
between 10 and 100 μL. To make an effective seal
between the tip and the syringe body, it is necessary to
2 μL
use a short length of soft silicone rubber tubing that
extends 1–2 mm beyond the nozzle of the syringe (if
the tubing is cut too short, you will not be able to fix a
tip onto the syringe).
When you use syringes like this, it is very important to
draw liquid into the tip only, and to use the graduations
on the tip as a guide. Ignore any markings on the
syringe itself and take care not to draw liquid into the
Hold here
Do not touch syringe barrel.
the point
Yellow
graduated
tip
100 μL
50 μL
20 μL
10 μL
Measure
to the
top of
each
band
With both types of syringe, the best results will be
obtained if you observe the following precautions:
 Never pull the plunger right out of the syringe —
when the plunger is re-inserted the seal may be
damaged;
 Before you load the tip, pull the plunger out a little
(1–2 mm). This will give you some extra air with
which to expel the last drop of liquid from the tip;
 When you dispense liquids, hold the syringe as near
to vertical as possible, and at eye level so that you
can see what you are doing;
 Remove the liquid from the point of the tip by
holding it against the inner wall of the tube into
which you are transferring the liquid;
 Do not touch the point of the tip with your fingers.
There are proteases and DNases in your sweat which
may contaminate and degrade the reactants.
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Fixed volume micropipettes
These pipettes can take white or
yellow tips: use the right tip for the
volume you require
Fixed volume micropipettes can be used to measure
small volumes with less than 1% error, but to achieve
this accuracy you need to use them correctly. Firstly,
the tip must be pushed firmly onto the pipette, so that
it forms an airtight seal. Secondly, note that the
pipettes have a double action. Use them as follows:
 Press the plunger down to the first stop;
 Place the tip into the liquid then slowly release the
plunger, drawing liquid into the tip;
Volume
A microlitre is one millionth of a litre
1 000 microlitres (μL) = 1 millilitre (mL)
1 000 millilitres = 1 litre (L)
 To expel every last drop of liquid from the tip, press
the plunger down hard to the second stop;
 After dispensing the liquid, lift the tip clear of the
liquid before letting go of the plunger (otherwise the
pipette will draw the liquid up again).
Acknowledgment
The Volvox project is funded by the Sixth Framework Program of the
European Commission.
www.bioscience-explained.org
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