How to: Agarose Gel Electrophoresis
Preparation of the Gel
1. Find a gel apparatus to use: box, tray and comb.
Make sure you have a comb that has enough wells for your sample. If you have a comb
with many wells, invite another group to go with you.
2. Figure out how much buffer you will need to fill the box, covering the gel.
Put the gel tray in the box, then fill the box with water until the level is over the gel tray by
a bit more than a half a centimeter. How much did you use? Round up to the next 100 mL.
3. Prepare 1X TBE or 1X TAE to fill the box (as determined above)
For a discussion of TBE vs TAE, see below. Be sure to record the information from the
stock bottle of 10X TBE or 50X TAE, and the volumes you use to prepare the gel.
4. Measure the gel tray; figure out the volume of agarose you will need
Plan to fill the tray to a depth of about 0.5 cm. (Recall that cm3 = mL). Round to some
convenient number.
5. Prepare the gel tray
Close off the ends (with device or tape) and setting an appropriate comb in place (note that
the comb should NOT touch the bottom, or you will be making holes instead of wells!)
6. Find an Erlenmeyer flask for prep of the agarose.
It should be for a volume quite a bit larger than the volume you want to actually make – so
that it doesn’t boil over. For 30 mL of agarose, a 75 mL flask is good. The Erlenmeyer is
good for mixing by swirling, and minimizing loss of fluid volume due to evaporation
7. Decide what concentration of agarose you need, and how much you need to weigh out
1% is good for resolving pieces of DNA 0.5 KB to about 3 KB. If your fragments are
smaller, say in the 100s of bp, increase to 1.2 or 1.5% (using regular agarose); if larger, use
a lower concentration (down to about 0.6%).
Make sure you do not use a low melting agarose. These are for resolving small fragments.
They are expensive and a bit difficult to use, and they require higher concentrations to firm
up. There are lots of agarose products – if you are in doubt, ask. Record the manufacturer,
product number, lot number of the agarose.
8. Weigh out the agarose, put it into the flask.
Don’t kill yourself to be precise here. If you are aiming for 0.35g, it is OK to be 0.34-0.35.
Pat it out of the bottle, so that you don’t contaminate agarose with nucleases. Record the
9. Add the appropriate amount of 1X buffer, and swirl it around a bit to mix.
MAKE SURE YOU DON’T ACCIDENTALLY USE WATER HERE!
Try to not get agarose high on the walls of the Erlenmeyer – it won’t mix in well
10. Microwave until no visible flecks remain.
The time will depend on the volume and the microwave. You must generally go past the
boiling stage, but not too long, or too much water will evaporate. That would make the
concentration of the agarose and the buffer components too high.
Watch the flask – when it starts to boil, take it out, swirl it (use an autoclave glove, and use
caution so it does not burn you), and put it back if you still see flecks of undisclosed
agarose. You can put a few flasks in at once, but then, as with frozen dinners, the batch
will take longer.
NOTE: At this point, some people put Ethidium Bromide directly into the gel to stain the
DNA. The appropriate amount is (hm… need to look this up!) _____________ .The
Ethidium intercalates between the bases of the DNA while it runs, and this saves time because
you don’t need to stain and destain the gel following the run. You also generate smaller
volumes of Eth Bromide waste this way, but there are some caveats:
Do not do this if the pieces you are looking for are quite small (400 bp or less). The
Ethidium travels toward the cathode, passing the DNA on its way to the anode.
Sometimes they pass too quickly, and smaller pieces do not get adequately labeled.
Do not do this if you are quantitating. The reasoning is the same as above. You want to
give all pieces the same opportunity to be stained to the same degree.
11. Pour the hot agarose into the gel tray (w/ the comb in it), and allow it to cool/harden.
Immediately put water in the flask (so that the residual agarose does not solidify in the
flask) It will take about 45-60 minutes to harden, which gives you time to take a break,
prepare samples (see Sample Prep section), find standards, or whatever you need to do.
12. Remove the gel from the casting setup and place it in the gel box. Make sure that the
wells are at the end where the black electrodes are. You will run toward the red
13. Pour 1X Running buffer into the box, covering the gel completely.
14. Load samples, noting any problems as you go (a missed well, one that flows over etc)
Set your sample up in the order for loading (see sample prep, “load plan”)
15. Attach the top, and connect the electrodes to the power supply.Turn the power supply
on, and adjust the Voltage.
In a TBE mini gel, we generally run at 120 V, and the samples will typically run nicely in
45 minutes. If you are in TAE, especially if you have low melting agarose, you may want
to run more slowly.
The real “rule of thumb” here is 10 V per cm of gel length. That (supposedly) gives optimal
separation. The minigels are typically 10 cm long, which would make it 100 volts. 120 V
is for getting it done quicker. More than that and it gets too hot.
If there are two gels on the same power supply, the voltage should still be set at 120V. The
total current will increase so that both gels run at 120V.
16. The gel can be stopped at any time
The pieces separate from each other as they go, so the longer the migration, the better the
separation. You may or may not need optimal separation, and so in some cases, you may
choose to stop the gel sooner or later. For most gels, it is good to let the tracker dye run to
the bottom half or third of the gel. If you look at the gel and your bands have not separated
to your satisfaction, you can put it back in the box and run it longer. (the same is NOT true
if you run the pieces you want off the end!!)
17. Staining:
If Ethidium Bromide has already been added, the gel can be examined and photographed
on the transilluminator without delay. (If you destain it (by soaking in some water), the
faint pink background will go away)
If Ethidium was not already added:
Stain: slide the gel into a glass pan containing 100 mL of water (or running buffer) and
10 uL of Ethidium Bromide (what is that final concetnration? _____). Put on the shaker
for 5-30 minutes, then discard the Ethidium Bromide [into a marked waste container].
Destain: replace the solution with water or buffer and shaking gently for 5 – 30
minutes.
The amount of time you stain or destain depends on what you need to do.
If you are quantitating, stain for the longer time to ensure equal access to the stain, and
destain the longer time to reduce the background as much as possible. If you only need to
know where in the gel the DNA is, and/or have plento of it you can use a shorter time. You
can shorten the destain time too, if you don’t really care so much about contrast
(sensitivity).
Use of water (di) is fine here, but if you think you might want to put the gel back into the
box to run longer, it would be wise to use the running buffer. This is because as you stain
and destain, the buffer in the gel will diffuse out, and water in, diluting the buffer in the gel.
18. Photograph and record the results.
If you are cutting out a band, make a photograph before and after the cut. (some labs have
video gel documentation systems)
Unless you are doing a transfer (Southern Blot), discard the gel in the dry Ethidium
Bromide waste.
The photograph is your record of the results. Tape it securely into your notebook
IMMEDIATLEY. Do not let it hang around loose, waiting for you to “catch up” in your
notebook. Its much worse to lose the data than to have it taped in “too soon” Label it with
the date, the samples, and any other information.
Sharpie works well on the photograph, but avoid making marks in the data field –keep the
marks to the periphery.
Sample Prep for Agarose Gels
This is hard to make a “protocol” for, because it is so dependent on what you are doing.
VOLUME:
The size of well defines the volume you can load. The widths are more or less standard, but the
volume is also a function of how deep the wells are, and that can vary depending on the
position of the comb, or the thickness of the gel. In a 0.5 cm deep gel, with 0.5 mm wide
combs, 6-15 uL is usually good.
SAMPLE BUFFER
The samples do all need to have “DNA Sample Buffer” , or “Loading Buffer” added to them.
This buffer has two critical ingredients:

Glycerol, sucrose, ficoll or some other component to increase the density of the
solution. This helps you get the sample into the well.

Tracking dye, usually bromophenol blue – a small colored dye that will run toward the
anode, ahead of almost all of the DNA pieces you might have. This allows you to see
how far the smallest pieces have run, so you don’t accidentally let them run off the gel.

Other components that may be included (but are not strictly necessary) are
Buffer (usually Tris, at pH 8, which DNA “likes”, also makes the BB blue – it
is a yukky green color at a more acid pH)
EDTA (which can stop enzyme reactions by chelating Mg, a common DNA
enzyme cofactor)

DNA Sample buffer is also sometimes called “6X” because it is made up at a
concentration that is 6 times higher than it needs to be when it is used (final, or working
concentration).
The volume of 6X that you add to a DNA sample is 1/5 of the starting sample
volume (so that it will be 1/6 of the final volume, which dilutes 6X down to 1X).
If you have 20 uL of an enzyme reaction, you would add 4 uL of 6X to it.
If you then load half of that, 12 uL, it would be the same as loading 10 uL (half)
of the orignal reaction.
HOW MUCH DNA?
The volume of sample that you add depends on the DNA concentration, and is really driven by
the amount of DNA you want to get into the well. This depends on the situation – the size of
the wells, and your purpose.
When you stain with Ethidium Bromide, you can see as little as 1 ng of DNA in a “standard”
size well (about 0.4 mm wide). In a wider well, that 1 ng will be spread out more and may not
be visible. In a narrower well, it will be all scrunched together, and be more visible. A good,
easily visible band will have 20 – 100 ng (standard well). More than 200 ng (0.2 ug) and the
band starts to get too fat – DNA smears above the band.
If the DNA is cut into many bands, we need to consider the amount of DNA in each band is a
fraction of the total amount of the added, and proportional to the size of the DNA.
Example:
A plasmid 4 KB in size is cut into two pieces, 1 Kb and 3 Kb
If I load 200 ng of this, ¼ of the DNA (50 ng) will be in the 1 Kb piece, and ¾ o(or 150
ng) will be in the 3 Kb piece.
If I want to purify a fragment, I may try to load a lot of DNA per well, because I will need to
remove the agarose, and more DNA I have per mg of agarose, the better. I have to be careful,
though, not to load so much that the DNA runs badly.
Sometimes I don’t have any idea what my DNA concentration really is, and I may work from
past experience, or make and educated guess.
For example Experience tells me that if I do a plasmid mini-prep from 3 mL of
bacterial culture, and resuspend the final DNA in 50 uL, that if I digest 5 uL of that in a
20 uL volume, loading half will generally give me good bands. By the same token, for
PCR reactions, 10 uL of reaction (+2 uL of 6X) should give me a good band.
STANDARDS
It is important to include a DNA standard on every gel. Without this, it is impossible to figure
out how big your fragments are. It also tells you that your gel is OK, even if your samples look
icky. Standards can also be used to help you determine your DNA concentration
There are two main kinds of standards: DNA cut into fragments, and Ladders
DNA cut into recognizable fragments.
The most common standard is DNA from bacteriophage Lambda that has been cut with the
Restriction Enzyme Hind III. The fragments are (in order of largest to smallest) 23 kb, 9.4 kb,
6.6 kb, 4.4 kb, 2.3 kb, 2.0 kb and 0.56 kb (there’s also one that is .125 kb, but you seldom see
it). The pattern made by H3 is instantly recognized by molecular biologists, and useful for
sizing anything 0.5 kb and larger.
For smaller pieces, the marker of choice is fx174 HaeIII (phage fx174, cut up with HaeIII). Its
fragments range from 1.3 kb down to 72 bp.
Both of these DNAs can be used as markers for quantitation as well as for size
l is about 44 Kb in size. If you load 500 ng of total H3, when you look at the cut up version,
about half of it will be in the 23 kb fragment (so 250 ng), and about 1/20th will be in the 2.3 kb
fragment (25 ng) – you can estimate the amount (in ng) of DNA in each band.
You can do the same with fx164HaeIII.
When using H3 a size standard, we usually load 500 ng, in a 12 uL volume, in a normal well.
The standard comes at 500 ug/mL (which is 500 ng/uL) – lets say we got 50 uL
We dilute it 1/10, to 50 ng/uL (= 500 ng/10 uL).
To do this, we add 450 uL of water, now its 500 uL.
We 1/5 that volume of 6X (to 500 uL we add 100 uL of 6X, so now there are 600 uL)
Now the amount to add for 500 ng is 12 uL. (but of course, we can add more or less)
The rest keeps nicely at 4 C until we need it.
One note about H3: Although phage  is linear DNA, the two ends are cohesive, and tend to
hang together by hydrogen bonds, so that the DNA is circular. The 23 kb piece is at one “end”
of linear , the 4.4 is at the other. When the fragments are all in solution, these two will tend to
hook up, form those H-bonds in solution, and the two will run as 27 kb. Because you cannot
see the difference between 24 and 27 kb, it will just look like the 4.4 kb piece has disappeared.
To prevent this, heat the sample for 2 min at 65 C before loading
Ladders
These are made by making strings of pieces attached end to end (concatamers), in all possible
sizes. A 100 bp ladder will have fragments 100 bp, 200 bp, 300 bp, 400 bp, 500 bp etc, so it is
much like a ruler. (The “rungs” are not all evenly spaced, though, because the migration
through the gel is not linear). You can get ladders in other size increments (50 bp, 200 bp, 1
kb). Because there is no pattern, the manufacturers of ladders often spike the mix with extra of
one size, so that you can pick out the one that is, say 500 bp. These provide an easy ruler for
size, but do not work as well for quantitation.
RECORDING SAMPLE PREP AND LOAD PLAN
When recording sample prep, it is important to note the initials/date of the 6X, and the volumes
used. For standards already prepared you should record the identity, concentration, initials and
date; if you make new standards, you will need manufacturer, lot number, original
concentration, calculations, volumes used, diluant added (is it water or buffer?) and 6X
(identified by initials and date of preparation) and final concentration.
Recording the load plan is important. If you don’t remember which samples are in which
wells, you may not be able to interpret your results. Even if you think it is obvious now, you
may not think so when you look back in a year or so.
Assign a lane number for each well, and indicate the sample that will be loaded into it. Keep in
mind that by convention, we almost always show gels running from the top of the page to the
bottom. If you number wells consecutively down the page, you run the risk of getting
confused about which one is on the right and with on the left. For that reason, I always write
my lane assignments across the page, the way I am going to show the gel.
TBE vs. TAE
The common electrophoresis buffers for DNA are TBE (Tris Borate EDTA) and Tris Acetate
EDTA. (Sometimes Tris Phosphate EDTA is used). Since borate, acetate and phosphate are all
buffers by themselves (weak acids), and Tris is a buffer (weak base), you can see that the buffer
control here is relatively complex. There are probably also electrochemical differences between
them that I do not begin to understand (DNAs do run at slightly different rates in the different
buffers, but still all run in proportion to their size). I am not sure what theory is behind the
use of each one, but practical considerations usually drive the choice. Some labs use TBE most
of the time and TAE when needed, some labs use TAE all the time. Here are the issues.

TAE can be made as a 50X stock, which saves time and space. TBE is made as a 10X
stock, and even at that concentration, the borate can crystallize out (which changes the
composition and pH of the buffer). To avoid this problem, people often autoclave their
TBE in order to more thoroughly dissolve the borate so that crystallization is less likely.

TBE can be reused whereas TAE cannot unless the tank is re-circulated during the run.
This is a pain; most people don’t do it. Not a huge issue in labs that are only running a
few mini-gels a day, but can be an issue if there are lots of gels or big gels (with big
tanks). Then one thinks about time and money.

TAE has a bit more resistance that TBE, and the gels get hotter during the run. This
usually isn’t a problem when using regular agarose, but can be a huge issue with lowmelting agarose. (It means you need to run at a lower voltage, so as not to let the temp
get high, because if it does, the low-melting agarose will melt during the run!).

The borate in TBE can inhibit some enzymatic reactions, so if you plan to use the DNA
you separate (cut out the piece and digest or ligate or otherwise use enzymes to treat it,)
we often run the gel in TAE instead. (If you are going to take steps to remove the
agarose, generally you also get rid of the TBE, but maybe not completely, and so its
better to just avoid that problem altogether by using TAE.