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bioscienceexplained Vol 3 No 2 134567

www.bioscience-explained.org 1 COPYRIGHT © by the Authors, 2007

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’.

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,

CORRESPONDENCE TO Dean Madden

National Centre for Biotechnology Education,

The University of Reading, Earley Gate, Reading,

RG6 6BZ

The United Kingdom [email protected]

bioscienceexplained 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.

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

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.

DNA evidence shows that the tropical papaya (Carica papaya) is a close relative of the humble cabbage

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.



The internal structure of a chloroplast, showing the DNA.

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 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.

DNA within the chloroplast

Stroma

Outer membrane

Inner membrane

Starch granule

Granum a stack of thyalkoids

Stroma lamellae

Lipid globule



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 differently-sized 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

Related plants give

DNA fragments of similar sizes



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 Ready-To-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)



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.



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 co-factors 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.

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.

OPTIONAL STOPPING POINT

Figure 2.

Close the cover and squash the leaf

Figure 1. Place the leaf

tissue on the card

Ensure that it fits within a box

Write the plant’s name

in pencil on the cover

Allow the card to dry for 1 hour

Figure 3



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,

Figure 5. Push the disc from the

punch using some plastic ‘wire’

Wash the disc twice in Purification Reagent

Wash the disc twice in

Purification Reagent Add 100 μL of Purification

Reagent Flick to mix

Remove the liquid REPEAT

Steps 3-6

100 µL

Figure 6a Figure 6b

Figure 4. Cut discs using a punch



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.

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.


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



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)

PCR ‘bead’, containing:

– Taq polymerase – Buffer – dNTPs – MgCl2

Primer 1 10 µL

Primer 2

10 µL

Water

4 or 6 µL

Primer 1: 5'–CGAAATCGGTAGACGCTACG–3' Primer 2: 5'–GGGGATAGAGGGACTTGAAC–3'

Figure 8



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.

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)...

Figure 9. Use forceps to

add a disc with plant DNA

Figure 10. Label the tube

Figure 11a. A PCR thermal

cycler



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

the water bath when it is not in use.

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 single-

stranded)

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.


30 seconds 30 seconds

30 seconds

Repeat this

3-step cycle 30 times

Figure 11b

W

rite the plant’s name in pencil on the cover



D DNA gel electrophoresis

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

Caution!

Hot agarose can scald

Figure 12

Figure 13

Carbon

fibre tissue

Cut two electrodes

42 mm

22 mm



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.

5. 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.

Once the gel has set, pour on TBE buffer, then lift the comb

out gently

TBE buffer

solution

Label the end of

the tank to show

the contents of each well

Black card under the

tank reveals the wells for loading

2-3 mm depth of buffer over the gel

Figure 14

Figure 15



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 DNA-degrading enzymes. The loading dye does not affect

the DNA, but moves through the gel slightly ahead of all but the smallest of DNA fragments.

Voltage Time to run gel

27 3½ hours 30 2 hours

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.

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.

Electrodes

Direction of DNA movement

APPLY NO MORE THAN 36

VOLTS!

Serious or lethal electrical shock may occur if you connect the equipment directly to a mains

electricity supply

Figure 16

Figure 17



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.

1 kb ’ruler’ Sizes in base

pairs

Smaller fragments may run off the gel

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.)

5. 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.

Positively-charged Azure A

binds to the negatively-charged phosphate groups of the DNA

Stain for only 4 minutes

Wells

Region with DNA bands

Loading dye

Figure 18

Figure 19



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.

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.

CAUTION! Molten agarose can scald and

it must be handled with care, especially just after it has

been heated in a microwave oven.



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.

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.

CAUTION!

Serious or lethal electrical shock may occur if you connect

the equipment directly to a

mains electricity supply.



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.



Azure A

Stain for nucleic acids

Makes 50 mL of 2 concentrate


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.



Agarose gel

For separating small fragments of nucleic acids

Makes 100 mL of 1.5% agarose


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


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.



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.



Notes

Agarose, dissolved in diluted buffer, not water,

must be used for the ‘real’ gels with DNA.

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.

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



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.



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, on-

line 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 closely-

related 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)

Watercress consistently gives a larger PCR product ٭



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).

Specimen results

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.

Wells

Region with DNA bands

Loading dye

Figure 20



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 1,5-

bisphosphate. 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.



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.

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.

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.



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 single-stranded 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 base-

pairing.

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.



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.



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 Second cycle Third cycle Fourth cycle

The sequential steps in the PCR



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

Two types of syringes — conventional 1 mL syringes and narrower ‘microsyringes’ — can be fitted with

disposable graduated tips and used to measure small

volumes. The white microsyringe tips are marked at 2 and 10 microlitres (μL). The larger yellow tips can be

fitted to a 1 mL syringe and used to dispense volumes between 10 and 100 μL. To make an effective seal

between the tip and the syringe body, it is necessary to 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

syringe barrel.

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.

Hold here

Do not touch the point

10 μL

2 μL

White

graduated tip

Hold here Do not touch the point

Yellow graduated

tip

Soft rubber tubing

100 μL

50 μL

20 μL

10 μL

Measure to the

top of each band



Fixed volume micropipettes

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;

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.

These pipettes can take white or

yellow tips: use the right tip for the volume you require

Volume A microlitre is one millionth of a litre

1 000 microlitres (μL) = 1 millilitre (mL) 1 000 millilitres = 1 litre (L)

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