DNA lab 2 (temporary): Agarose Gel Electrophoresis
How to pour, load, and run an agarose gel.
MATERIALS
Buffers and Solutions
Agarose solutions (please see Step 3)
DNA staining solution
Electrophoresis buffer
6x Gel-loading buffer
Nucleic Acids and Oligonucleotides
DNA samples
DNA size standards
Samples of DNAs of known size are typically generated by restriction enzyme digestion of a plasmid or
bacteriophage DNA of known sequence. Alternatively, they are produced by ligating a monomer DNA
fragment of known size into a ladder of polymeric forms.
METHOD
1. Seal the edges of a clean, dry glass plate (or the open ends of the plastic tray supplied with the
electrophoresis apparatus) with tape to form a mold. Set the mold on a horizontal section of the
bench.
2. Prepare sufficient electrophoresis buffer (usually 1x TAE or 0.5x TBE) to fill the electrophoresis
tank and to cast the gel.
It is important to use the same batch of electrophoresis buffer in both the electrophoresis tank
and the gel.
3. Prepare a solution of agarose in electrophoresis buffer at a concentration appropriate for
separating the particular size fragments expected in the DNA sample(s): Add the correct amount
of powdered agarose (please see table below) to a measured quantity of electrophoresis buffer in
an Erlenmeyer flask or a glass bottle.
Range of Separation in Cells Containing Different Amounts of Standard Low-EEO Agarose
Agarose Concentration Range of Separation of
in Gel (% [w/v])
Linear DNA Molecules (kb)
0.3
5-60
0.6
1-20
0.7
0.8-10
0.9
0.5-7
1.2
0.4-6
1.5
0.2-3
2.0
0.1-2
4. Agarose gels are cast by melting the agarose in the presence of the desired buffer until a clear,
transparent solution is achieved. The melted solution is then poured into a mold and allowed to
harden. Upon hardening, the agarose forms a matrix, the density of which is determined by the
concentration of the agarose.
5. Loosely plug the neck of the Erlenmeyer flask with Kimwipes. If using a glass bottle, make certain
the cap is loose. Heat the slurry in a boiling-water bath or a microwave oven until the agarose
dissolves.
Heat the slurry for the minimum time required to allow all of the grains of agarose to dissolve.
6. Use insulated gloves or tongs to transfer the flask/bottle into a water bath at 55°C. When the
molten gel has cooled, add ethidium bromide to a final concentration of 0.5 μg/ml. Mix the gel
solution thoroughly by gentle swirling.
IMPORTANT SYBR Gold should not be added to the molten gel solution.
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7. While the agarose solution is cooling, choose an appropriate comb for forming the sample slots in
the gel. Position the comb 0.5-1.0 mm above the plate so that a complete well is formed when the
agarose is added to the mold.
8. Pour the warm agarose solution into the mold.
The gel should be between 3 mm and 5 mm thick. Check that no air bubbles are under or
between the teeth of the comb. Air bubbles present in the molten gel can be removed easily by
poking them with the corner of a Kimwipe.
9. Allow the gel to set completely (30-45 minutes at room temperature), then pour a small amount of
electrophoresis buffer on the top of the gel, and carefully remove the comb. Pour off the
electrophoresis buffer and carefully remove the tape. Mount the gel in the electrophoresis tank.
10. Add just enough electrophoresis buffer to cover the gel to a depth of approx. 1 mm.
11. Mix the samples of DNA with 0.20 volume of the desired 6x gel-loading buffer.
The maximum amount of DNA that can be applied to a slot depends on the number of fragments
in the sample and their sizes. The minimum amount of DNA that can be detected by photography
of ethidium-bromide-stained gels is approximately 2 ng in a 0.5-cm-wide band (the usual width of
a slot). More sensitive dyes such as SYBR Gold can detect as little as 20 pg of DNA in a band.
12. Slowly load the sample mixture into the slots of the submerged gel using a disposable
micropipette, an automatic micropipettor, or a drawn-out Pasteur pipette or glass capillary tube.
Load size standards into slots on both the right and left sides of the gel.
13. Close the lid of the gel tank and attach the electrical leads so that the DNA will migrate toward the
positive anode (red lead). Apply a voltage of 1-5 V/cm (measured as the distance between the
positive and negative electrodes). If the leads have been attached correctly, bubbles should be
generated at the anode and cathode (due to electrolysis), and within a few minutes, the
bromophenol blue should migrate from the wells into the body of the gel. Run the gel until the
bromophenol blue and xylene cyanol FF have migrated an appropriate distance through the gel.
The presence of ethidium bromide allows the gel to be examined by UV illumination at any stage
during electrophoresis. The gel tray may be removed and placed directly on a transilluminator.
Alternatively, the gel may be examined using a hand-held source of UV light. In either case, turn
off the power supply before examining the gel! (In a typical 1% agarose gel in TAE buffer or TBE
buffer, bromophenol blue migrates at the same rate as a DNA fragment of approximately 500bp,
Xylene cyanol typically migrates at about the same rate as a 4000bp DNA fragment.)
14. When the DNA samples or dyes have migrated a sufficient distance through the gel, turn off the
electric current and remove the leads and lid from the gel tank. If ethidium bromide is present in
the gel and electrophoresis buffer, examine the gel by UV light and photograph the gel.
Otherwise, stain the gel by immersing it in electrophoresis buffer or H 2O containing ethidium
bromide (0.5μg/ml) for 30-45 minutes at room temperature or by soaking in a 1:10,000-fold
dilution of SYBR Gold stock solution in electrophoresis buffer.
RECIPES
6x Gel-loading Buffer I
0.25% (w/v) bromophenol blue
0.25% (w/v) xylene cyanol FF
40% (w/v) sucrose in H2O
Store at 4°C.
6x Gel-loading Buffer II
0.25% (w/v) bromophenol blue
0.25% (w/v) xylene cyanol FF
15% (w/v) Ficoll (Type 400; Pharmacia) in H2O
Store at room temperature.
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6x Gel-loading Buffer III
0.25% (w/v) bromophenol blue
0.25% (w/v) xylene cyanol FF
30% (v/v) glycerol in H2O
Store at 4°C.
6x Gel-loading Buffer IV
0.25% (w/v) bromophenol blue
40% (w/v) sucrose in H2O
Store at 4°C.
Alkaline Gel-loading Buffer
300 mM NaOH
6 mM EDTA
18% (w/v) Ficoll (Type 400, Pharmacia)
0.15% (w/v) bromocresol green
0.25% (w/v) xylene cyanol
For a 6x buffer.
DNA Staining Solution
ethidium bromide (10 mg/ml)
SYBR Gold
EDTA
To prepare EDTA at 0.5 M (pH 8.0): Add 186.1 g of disodium EDTA·2H2O to 800 ml of H2O. Stir
vigorously on a magnetic stirrer. Adjust the pH to 8.0 with NaOH (approx. 20 g of NaOH pellets).
Dispense into aliquots and sterilize by autoclaving. The disodium salt of EDTA will not go into
solution until the pH of the solution is adjusted to approx. 8.0 by the addition of NaOH.
Ethidium Bromide
Add 1g of ethidium bromide to 100 ml of H2O. Stir on a magnetic stirrer for several hours to ensure
that the dye has dissolved. Wrap the container in aluminum foil or transfer the 10 mg/ml solution
to a dark bottle and store at room temperature.
Ficoll 400 (20% w/v)
Dissolve the Ficoll in sterile H2O and store the solution frozen in 100μl aliquots at -20°C.
Glycerol
To prepare a 10% (v/v) solution: Dilute 1 volume of molecular-biology-grade glycerol in 9 volumes
of sterile pure H2O. Sterilize the solution by passing it through a pre-rinsed 0.22-μm filter. Store in
200-ml aliquots at 4°C.
NaOH
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The preparation of 10 N NaOH involves a highly exothermic reaction, which can cause breakage
of glass containers. Prepare this solution with extreme care in plastic beakers. To 800 ml of H 2O,
slowly add 400g of NaOH pellets, stirring continuously. As an added precaution, place the beaker
on ice. When the pellets have dissolved completely, adjust the volume to 1 liter with H2O. Store
the solution in a plastic container at room temperature. Sterilization is not necessary.
SYBR Gold
SYBR Gold (Molecular Probes) is supplied as a stock solution of unknown concentration in
dimethylsulfoxide. Agarose gels are stained in a working solution of SYBR Gold, which is a
1:10,000 dilution of SYBR Gold nucleic acid stain in electrophoresis buffer. Prepare working
stocks of SYBR Gold daily and store in the dark at regulated room temperature.
TAE
Prepare a 50x stock solution in 1 liter of H2O:
242 g of Tris base
57.1 ml of glacial acetic acid
100 ml of 0.5 M EDTA (pH 8.0)
The 1x working solution is 40 mM Tris-acetate/1 mM EDTA.
TBE
Prepare a 5x stock solution in 1 liter of H2O:
54 g of Tris base
27.5 g of boric acid
20 ml of 0.5 M EDTA (pH 8.0)
The 0.5x working solution is 45 mM Tris-borate/1 mM EDTA.
TBE is usually made and stored as a 5x or 10x stock solution. The pH of the concentrated stock
buffer should be approx. 8.3. Dilute the concentrated stock buffer just before use and make the
gel solution and the electrophoresis buffer from the same concentrated stock solution. Some
investigators prefer to use more concentrated stock solutions of TBE (10x as opposed to 5x).
However, 5x stock solution is more stable because the solutes do not precipitate during storage.
Passing the 5x or 10x buffer stocks through a 0.22-μm filter can prevent or delay formation of
precipitates.
TPE
Prepare a 10x stock solution in 1 liter of H2O:
108 g Tris base
15.5 ml of 85% (1.679 g/ml) phosphoric acid
40 ml of 0.5 M EDTA (pH 8.0)
The 1x working solution is 90 mM Tris-phosphate/2 mM EDTA.
CAUTIONS
Ethidium bromide
Ethidium bromide is a powerful mutagen and is toxic. Consult the local institutional safety officer
for specific handling and disposal procedures. Avoid breathing the dust. Wear appropriate gloves
when working with solutions that contain this dye.
SYBR Gold
SYBR Gold is supplied by the manufacturer as a 10,000-fold concentrate in DMSO which
transports chemicals across the skin and other tissues. Wear appropriate gloves and safety
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glasses and decontaminate according to Safety Office guidelines. See DMSO.
PCR NOTES
1. PCR buffers are generally supplied by the manufacturer when you purchase a thermostable DNA
polymerase. Check the composition of the buffer and specifically whether it contains MgCl2. Magnesium
ions are critical for DNA synthesis. Some buffers will contain MgCl2, typically designed to give a final
concentration of 1.5 mM in the final PCR. Other buffers will not contain any MgCl2, but a stock solution
will usually be supplied by the manufacturer to allow you to determine the optimal MgCl2 concentration.
2. If you are setting up several reactions then prepare a premix of any common components to reduce
pipetting steps and potential contamination.
3. The denaturation temperature should be as low as reasonable to denature the template DNA and often
92°C will be efficient, although most protocols will recommend 94°C, and most people use this
temperature. For difficult templates, such as GC-rich sequences, a higher temperature may be necessary,
perhaps 96°C. Also this extended initial denaturation phase may not be necessary or could be
significantly reduced to 1 or 2 min in many applications. These measures will extend the functional life of
the DNA polymerase molecules.
4. The length of incubation times at each step will depend critically on the thermal cycler characteristics.
Often short times of 10–30 s are sufficient for the denaturation and annealing steps. In robust PCR
screening for thermal cyclers that monitor tube temperature the incubations can be as short as 1 s.
5. The time for the extension step is usually based on the rule of thumb of 1 kb/min. For shorter products
therefore the time can be reduced, while for longer templates it should be increased.
6. This annealing temperature of 55°C is a useful starting point for many PCRs, but can optimally be
between 40 and 72°C, depending upon the primer–template combination. The annealing temperature is
usually set to 3~5°C lower than Tm.
7. Optimal working temperature for Taq DNA polymerase is 72°C.
8. The number of cycles depends upon the complexity and amount of template added. Generally for
plasmid templates 25 cycles is sufficient whereas for genomic DNA between 30 and 35 cycles are usually
necessary. It is sometimes helpful during a genomic amplification to remove 5 μl aliquots at 30 and 35
cycles to compare with the 40-cycle sample to follow the accumulation of the specific band.
9. Several equations are available to calculate the melting temperature of hybrids formed between an
oligonucleotide primer and its complementary target sequence. An empirical and convenient equation,
known as "The Wallace rule" (Suggs et al. 1981; Thein and Wallace 1986), can be used to calculate the
melting temperature for perfect duplexes 15-20 nucleotides in length in solvents of high ionic strength
(e.g., 1M NaCl):
Tm (Celsius) = 2(A+T) + 4(G+C)
Where (A+T) is the sum of the A and T residues in the oligonucleotide and (G+C) is the sum of G and C
residues in the oligonucleotide.
10. The melting temperature (Tm) is the temperature at which one-half of a particular DNA duplex will
dissociate and become single strand DNA (the temperature at which half of the DNA strands are in the
double-helical state and half are in the "random-coil" states). The stability of a primer-template DNA
duplex can be measured by its Tm. Primers with melting temperatures in the range of 52-58°C generally
produce better results than primers with lower melting temperatures. While the annealing temperature
can go as high as 72°C, primers with melting temperatures above 65°C have a higher potential for
secondary annealing.
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11. One consequence of having too low an annealing temperature is that one or both primers will anneal
to sequences other than the true target, as internal single-base mismatches or partial annealing may be
tolerated. This can lead to nonspecific amplification and will consequently reduce the yield of the desired
product if the 3′ -most base is paired with a target. Conversely, too high an annealing temperature may
yield little product, as the likelihood of primer annealing is reduced.
12. Generally, many primer pairs producing longer amplification products worked better at lower salt
concentrations, whereas many primer pairs producing short amplification products worked better at higher
salt concentrations.
Troubleshooting for PCR and multiplex PCR
Troubleshooting discussion is based on the PCR protocol as described in the table below. All reactions
are run for 30 cycles.
COMPONENT
VOLUME FINAL CONCENTRATION
1.autoclaved ultra-filtered water (pH 20.7µL
7.0)
-
2.10x PCR Buffer*
2.5µL
1x
3.dNTPs mix (25 mM each
nucleotide)
0.2µL
200 µM (each nucleotide)
4.primer mix (25 pmoles/µL each
primer)
0.4µL
0.4 µM (each primer)
5.Taq DNA polymerase (native
enzyme)
0.2µL
1 Unit/25 µL
6.genomic DNA template (100
ng/µL)
1.0µL
100 ng/25 µL
* The 10x PCR buffer contains: 500 mM KCl; 100 mM Tris-HCl (pH 8.3); 15 mM MgCl2 (the final
concentrations of these ingredients in the PCR mix are: 50 mM KCl; 10 mM Tris-HCl; 1.5 mM MgCl2).
QUESTIONS
1. I get (many) longer unspecific
products. What can I do?
SOLUTIONS
Decrease annealing time
Increase annealing temperature
Decrease extension time
Decrease extension temperature to 62-68º C
Increase KCl (buffer) concentration to 1.2x-2x, but keep MgCl2
concentration at 1.5-2mM.
Increase MgCl2 concentration up to 3-4.5 mM but keep dNTP
concentration constant.
Take less primer
Take less DNA template
Take less Taq polymerase
If none of the above works: check the primer for repetitive
sequences (BLAST align the sequence with the databases) and
change the primer(s)
Combine some/all of the above
2. I get (many) shorter unspecific Increase annealing temperature
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products. What can I do?
Increase annealing time
Increase extension time
Increase extension temperature to 74-78º C
Decrease KCl (buffer) concentration to 0.7-0.8x, but keep MgCl2
concentration at 1.5-2mM
Increase MgCl2 concentration up to 3-4.5 mM but keep dNTP
concentration constant
Take less primer
Take less DNA template
Take less Taq polymerase
If none of the above works: check the primer for repetitive
sequences (BLAST align the sequence with the databases) and
change the primer(s)
Combine some/all of the above
3. Reaction was working before,
but now I can't get any product.
Make sure all PCR ingredients are taken in the reaction (buffer,
template, Taq, etc)
Change the dNTP solution (very sensitive to cycles of thawing
and freezing, especially in multiplex PCR)
If you just bought new primers, check for their reliability (bad
primer synthesis ?)
Increase primer amount
Increase template amount
Decrease annealing temperature by 6-10º C and check if you get
any product. If you don't, check all your PCR ingredients. If you
do get products (including unspecific ones) reaction conditions as
described above.
Combine some/all of the above
4. My PCR product is weak. Is
Gradually decrease the annealing temperature to the lowest
there a way to increase the yield? possible.
Increase the amount of PCR primer
Increase the amount of DNA template
Increase the amount of Taq polymerase
Change buffer (KCl) concentration (higher if product is lower than
1000bp or lower if product is higher than 1000bp)
Add adjuvants. Best, use BSA (0.1 to 0.8 µg/µL final
concentration). You can also try 5% (v/v, final concentration)
DMSO or glycerol.
Check primer sequences for mismatches and/or increase the
primer length by 5 nucleotides
Combine some/all of the above
5. My two primers have very
different melting temperatures
(Tm) but I cannot change their
locus. What can I do to improve
PCR amplification?
An easy solution is to increase the length of the primer with low
Tm. If you need to keep the size of the product constant, add a
few bases at the 3' end. If size is not a concern, add a few bases
at either the 3' or the 5' end of that primer.
6. I have a number of primer pairs
I would like to use together. Can I
run a multiplex PCR with them?.
How?
Very likely, yes.
Try amplify all loci seaprately using the same PCR program. If
one of the primer pairs yields unspecific products, keep the
cycling conditions constant and change other parameters as
mentioned above (#1 and #2).
Mix equimolar amounts of primers and run the multiplex reaction
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either in the same cycling conditions or by decreasing only the
annealing temperature by 4º C.
If some of the loci are weak or not amplified, read below !!
7. How many loci can I amplify in Difficult to say. The author has routinely amplified from 2 to 14
multiplex PCR at the same time? loci.
Literature describes up to 25 loci or so.
8. One or a few loci in my
multiplex reaction are very weak
or invisible. How can amplify
them?
The first choice should be increasing the amount of primer for the
"weak" loci at the same time with decreasing the amount of primer
for all loci that can be amplified. The balance between these
amounts is more important than the absolute values used !!.
Check primer sequences for primer-primer interactions
9. Short PCR products in my
Increase KCl (buffer) concentration to 1.2x-2x, but keep MgCl2
multiplex reaction are weak. How concentration at 1.5-2mM
can I improve their yield?
Decrease denaturing time
Decrease annealing time and temperature
Decrease extension time and temperature
Increase amount of primers for the "weak" loci while decreasing
the amount for the "strong" loci.
Add adjuvants. Best, use BSA (0.1 to 0.8 µg/µL final
concentration). You can also try 5% (v/v, final concentration)
DMSO or glycerol
Combine some/all of the above
10. Longer PCR products in my Decrease KCl (buffer) concentration to 0.7-0.8x, but keep MgCl2
multiplex reaction are weak. How concentration at 1.5-2mM
can I improve their yield?
Increase MgCl2 concentration up to 3-4.5 mM but keep dNTP
concentration constant.
Increase denaturing time
Increase annealing time
Decrease annealing temperature
Increase extension time and temperature
Increase amount of primers for the "weak" loci while decreasing
the amount for the "strong" loci
Add adjuvants. Best, use BSA (0.1 to 0.8 µg/µL final
concentration). You can also try 5% (v/v, final concentration)
DMSO or glycerol
Combine some/all of the above
11. All products in my multiplex
reaction are weak. How can I
improve the yield?
Decrease annealing temperature in small steps (2º C)
Decrease extension temperature to 62-68º C
Increase extension time
Increase template concentration
Increase overall primer concentration
Adjust Taq polymerase concentration
Change KCl (buffer) concentration, but keep MgCl2 concentration
at 1.5-2mM
Increase MgCl2 concentration up to 3-4.5 mM but keep dNTP
concentration constant.
Add adjuvants. Best, use BSA (0.1 to 0.8 µg/µL final
concentration). You can also try 5% (v/v, final concentration)
DMSO or glycerol
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Combine some/all of the above
12. Unspecific products appear in If long: increase buffer concentration to 1.2-2x, but keep MgCl2
my multiplex reaction. Can I get concentration at 1.5-2mM
rid of them somehow?
If short: decrease buffer concentration to 0.7-0.9x, but keep
MgCl2 concentration at 1.5-2mM
Gradually increase the annealing temperature
Decrease amount of template
Decrease amount of primer
Decrease amount of enzyme
Increase MgCl2 concentration up to 3-4.5 mM but keep dNTP
concentration constant
Add adjuvants. Best, use BSA (0.1 to 0.8 µg/µL final
concentration). You can also try 5% (v/v, final concentration)
DMSO or glycerol
If nothing works: run PCR reactions for each (multiplexed) locus
individually, using an annealing temperature lower than usual.
Compare the unspecific products for each locus tested with the
unspecific products seen when running the multiplex PCR. This
may indicate which primer pair yields the unspecific products in
the multiplex reaction.
Combine some/all of the above
(Note: primer-primer interactions in multiplex PCR are usually
translated into lack of some amplification products rather than the
appearance of unspecific products)
KCl (salt) concentration in PCR
People who develop PCR buffers for a living has probably figured out that it is easy to melt (denature)
DNA in the denaturation step by simply using a high temperature, usually 95C. But it is harder to get the
primers to stick in the annealing step. Adding some salt to thePCR buffer would stabilize all doublestranded DNA, and in this case, it's probably more important to stabilize the primer-template double
strands so the polymerase can have a nice starting point to work from. As you know both strand of
DNA are negatively charged, salt (especially the cations) can form a salt bridge between the two
negatively charged strand and bring the two strand close enough so that they can easily recognize and
bind to the complementary sequence using hydrogen bond with each other thus become more stable.
Generally, primer pairs producing longer amplification products(usually more than 1kb) worked better at
lower salt concentrations, whereas primer pairs producing short amplification products(usually less than
1kb) worked better at higher salt concentrations.
This is because at lower salt concentration, for short PCR product, the binding of the primer to template is
relatively not strong which will reduce the short PCR products, for long PCR product, in low salt
concentration salt bridge effect is minimal, the two strand of long PCR product can still be easily
separated, although binding of the primer to template is also relatively not strong, but good separation of
two strand here plays dominant role.
In higher salt concentration, short PCR product can still separate efficiently easily although there is salt
bridge effect, and the annealing of the primer to the template is enhanced due to the salt bridge, and this
will greatly increase the production of short PCR product, on the other hand, however, for the
long PCR product, although the primer can also bind more tightly to the template, but the high salt
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concentration here will decrease the efficient separation of the double strand of the long PCR product,
thus prevent the binding of the primer to the template and decrease the long PCR product.
In short, the basic rule is if you want to enhance short PCR products or reduce the long PCR products,
you increase the KCl concentration in the buffer by 1.2X-2X, if you want to do the opposite way which is
to enhance the long PCR products and reduce short PCR products, you reduce the KCl concentration by
1.2X-2X.
References:
Molecular Cloning, a Laboratory Manual, 3rd edition. Joseph Sambrook and David W. Russell. 2001.
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