How to: Restriction Enzyme (RE) Digests
RE Digests are actually easy to set up, because the only things you need to combine are

The DNA

The enzyme(s) of choice

The buffer that the enzyme “likes”
However, if you have to be the one to figure out how much of each to use, it seems complicated,
especially at first. It is mainly driven by the amount of DNA you choose to digest. What
follows is a pretty complete (I think) description of the several different sorts of thing that
contribute to determining the appropriate amounts of the various components.
The DNA
You know what you want to cut – but you do need to decide how much to use. This depends on
what you are doing, and flows either from calculations or experience (or a little of each). Some
detail on how one decides these things are [far] below, but for now, lets just plan for each pair to
digest 5 ug of pCAT-P to obtain the insert. (By the way, this is a lot!)
The enzyme of choice – how much do we use?
Enzymes are quantitated in Units, which
For REs, A unit is defined as the amount (ug) of enzyme required to digest 1 ug of substrate
DNA in 1 hr at the recommended temperature in the recommended buffer. The substrate could
be any DNA that has a site for (i.e., sequence of bases recognized by) the enzyme – for the
purposes of defining the Unit it is often bacteriophage lambda, or one of about 4 other plasmids
or viruses.
You could determine exactly how much enzyme you need for, say 4 ug of your DNA by
calculating how many moles of this particular RE site there are in 1 ug of lambda, and then
comparing that with the number of moles of sites in 4 ug of your DNA. This is not hard, really,
but it is often more trouble than it’s actually worth. Why?
1) You can calculate the amount that should work, but if the enzyme has lost any
activity, you would need to add more anyway, and how much more is hard to guess.
2) If you are worried that you may not have enough enzyme, you can usually let the
reaction go longer – several hours, even overnight. (This is true because the
commercially prepared enzymes are “clean” -- they have extremely low amounts
contaminating nucleases, and thus are “safe” to leave on your DNA for long
incubations).
3) A handy rule of thumb usually works just as well anyway.
The rule of thumb many of us use is: 10 U per ug of DNA.
Since most enzymes are sold at 10 U/uL, this means 1 uL per ug of DNA. (some labs use even
less – it depends on how good they think their enzymes are, how expensive they were, how much
they care that digestion go to completion, how much of a hurry they are in .... etc.)
We will use the rule of thumb.
What temperature will be used?
The temperature an enzyme likes can be looked up. Most of the ones people use routinely like
37 C, but some, like Sma I like room temperature, and others, such as Taq I, prefer high
temperatures. Although you won’t really need to have the correct water bath or block ready until
you actually do the reaction, its good to know what you will need, and make sure you can get it.
It also may be something you need to think about if you are digesting with two enzymes at once
(if you digest with Sma I and EcoRI at 37 C, you’ll just kill the Sma and it won’t cut)
What buffer to use?
The enzyme also dictates the buffer.
Different enzymes have different requirements for pH, [NaCl], [Mg++] and other things.
Enzymes are sold with 10X solutions which, when diluted to 1X will make the conditions just
ducky for that enzyme.
If you are only digesting with one enzyme, it's very easy – just use the manufacturers
recommended buffer. However, if you are digesting with more than one, and they “like”
different buffers, you have to figure out how to make conditions nice for each to work. There are
several ways to do this
1) Look up buffer compatibility for the enzymes you are using. All the
manufacturers make these charts. They tell you if your enzyme will work in
another buffer. Some companies make a “multi-enzyme buffer” that most
enzymes will work pretty well in. If you can, its good to use enzymes form a
single company, because then you can trust the activity claims for each enzyme in
their buffers. If you have two different manufacturers for your enzymes, you
have to really look at the situation more carefully.
2) If you cannot get them in the same buffer, perhaps you can do the digest in a
buffer that one of the enzymes likes, and then change the conditions (say, increase
the salt concentration) to achieve conditions that the other one likes.
3) If you can’t do this, you may need to “desalt” the solution – which is generally
done by using EtOH to precipitate the DNA, and then resuspending it in water, so
that a different 10X buffer can be added (gets the ideal condition, but usually
results in the loss of 10-30% of your DNA.
We have to cut with EcoRI and HindIII. In the Invitrogen system, one works in the buffer called
REact 2, and the other in REact 3. One of these is actually OK for both enzymes. The new
enzymes I just bought are from Biolabs. Will one of their enzymes work?
How much buffer to use? Well its 10X and you want it to be 1X, so you need to use 1/10 your
total reaction volume. So ....
What volume should I set up my reaction in?
Generally we try to minimize volumes, so that collisions between molecules that need to interact
(enzymes and their DNA targets) happen more frequently. But we also need to make sure we
have an appropriate volume to work with. (Just because you could get away with doing the
reaction in 2 uL doesn’t mean you should – too small to measure accurately, and also loads of
evaporation in even a very small tube). A volume of 20 uL to 100 uL is typical.
What often determines the final reaction volume is the volume of enzyme you use.
Here’s why:
Enzymes are stored in the freezer because most tend to lose activity if stored at warmer
temperatures (hydrogen bonds get broken and such). They are stored in 50% glycerol to avoid
the formation of solid ice crystals, which can rip up proteins). Glycerol, however, can have
peculiar effects on the way enzymes recognize their targets. It can make them cut in unexpected
places, which is called “star activity” (for example, EcoRI generally recognizes GAATTC, but in
too much glycerol it also cuts at AATT sites, so it cuts too often, and may cut your desired piece
into smaller ones).
You need to set the volume such that final concentration of glycerol is only 5%, or less.
This means it must be at least 10 times more than the volume of the enzyme you use
Do the C1V1 if you need to: (50%)(2 uL) = (5%) X
OK so...... now you should be able to start filling in the blanks. Assuming we can do a
simultaneous double digest (i.e., both enzymes at the same time, in the same buffer) .....
For 5 ug of DNA we need ____ U of each enzyme (using the rule of thumb)
If the enzymes are at 10U/uL, that’s _____ uL each, or ____ uL total
(note: sometimes commonly used enzymes are sold at higher concentrations, such as 40 or 50
U/uL, and sometimes very expensive enzymes are sold at lower concentrations – pay attention to
that before you set up your reaction)
That means the total volume will need to be _____ uL (so that the glycerol is 5%).
That targeted total volume tells you how much of the 10X buffer to use: _______ uL.
What volume of DNA to use? Well, you need to know the concentration! What if the DNA is at
0.25 ug/uL, how many uL do I need for 5 ug? _______ uL
OK so having determined these, you write the components of the reaction down:
DNA
EcoRI
Hind III
10X Buffer
H2O
Final Volume:
_____ uL
_____ uL
_____ uL
_____ uL
_____ uL
20 uL
note: be sure to record concentration (# U/uL) and lot numbers, as well as
manuf, and expiration dates on enzymes. They are relatively “sensitive” reagents
Your actual calculation will clearly be a little different, depending on the concentration of DNA
and the enzymes, so find out, recalculate if necessary and then you can proceed to .....
Actually setting up the reactions:
The reactions are generally done in 1.5 mL microfuge tubes.
Label your tube(s) – initials, date and what it is. (“What it is” should be on the top)
When you actually set up the reaction, the order of addition is flexible with the exception of the
enzyme -- add the enzyme last.
I usually best to start with whatever is the largest volume, because its easer to add small volumes
of liquid to a larger volume – very tiny volumes are hard to get out of the tube.
When all reagents have been added, vortex or flick to mix
If droplets are on the sides or top of the tube, get them down by tapping gently on the table, or
use the small centrifuges for a second or so.
Set the tube in a water bath or heat block set to the appropriate temperature, and RECORD THE
TIME YOU DID SO as well as the TEMP.
Record the time you take it out too, so you know how long the reaction really was.
When the reaction is done, you can just transfer it to 4 C and all will be well.
If you want to “kill” the enzymes, usually heating to 65 C will do it. (Not all are sensitive to heat
though. There are charts in the product books – use them!)
Sometimes people add DNA sample buffer to it (see Electrophoresis handout). This, by itself,
won’t stop the reaction unless there is EDTA in the buffer (EDTA chelates Mg++ and Mg is
needed for most DNA modifying enzymes to work), but it does make things easier if your next
step is a gel.
If your next step is something other than a gel (such as another digest) you need to think about
whether EDTA or heating is a good idea. If you aren’t sure, just put it in the fridge.
*********************************************************************
How do you decide how much DNA to digest? It depends on what you are doing
If you are digesting plasmid to just see the bands (for example, to find out where the restriction
enzyme sites are on a new piece of DNA – mapping), you need to make sure you have between 5
and 100 ng in each band loaded in a subsequent gel (with a standard size well that is about .5-.6
cm in width, and the DNA stained with Ethidium Bromide). This way you will be able to see
them when they are stained, but they won’t be so overloaded that the bands become distorted. (If
your bands are skinnier, you need to use less, because it will be crammed into a smaller space;
with wider bands, you may not be able to see less than 5 ng, because it will be spread out over a
larger area.
So lets say you had a plasmid that is 4000 base pairs in length.
You are going to cut it into pieces that are 1000 bp and 3000 bp.
If you digest 1 ug of the plasmid DNA, the 1 kb piece will have 250 ng, and the 3 kb piece will
have 750 ng. So if you loaded the whole digested amount on the gel it would be too much! You
can either do your digest in a large volume and load only a fraction of it, or use less DNA to start
with.
a)
digest 1 kb in 100 uL; when the reaction is complete load only 10 uL of that; you will
have 25 and 75 ng in each band)
b)
use 200 ng of DNA, do the reaction in 20 uL and load half of it – you will again have
25 and 75 ng in each band (I suggest that you generally make a bit more digest than
you plan to load, so that if something bad happens to your gel, or when you load, you
have some sample in reserve)
If you just want to see bands, but you have no idea what the starting concentration of your
DNA is, you make an educated guess. For example, it is difficult to quantitate the DNA in
plasmid mini-preps (more on that later), but experience suggests the following:

if I am using a high copy number plasmid (something you need to find out about your
plasmid), and I take 1.5 mL of an overnight culture,

carry out one of several standard protocols for preparing plasmid,

resuspend the final plasmid DNA in 50 uL of water,

if I digest 5 uL of that in a volume of 20 uL,

and load half of that on a gel ....... I generally can see nice bands.
Some digests, however, are preparatory – meaning that the goal is to “prepare” quite a bit of
some DNA. In order to figure out how much to start with, we sort of have to look far
downstream, to see how much we need when we get to the end.
For a subcloning, we will be ligating insert and vector together.
 The ligation reaction requires that the concentration of DNA, and the ratio between insert
and vector, to be within certain parameters. Typically we want between 10 and 100
ng of DNA, and an insert:vector ratio of 3:1 (that's a molar ratio, indicating that there
are three molecules of insert fragment for every molecule of vector).
 If will be putting a small insert (say 500 bp) into a vector that is 2000 bp, for 50 ng of
vector to get a 1:1 molar ratio we need 12.5 ng of the small insert (the small piece is
one quarter the size of the vector, so one of it weighs a quarter what one of the other
one weighs);
 For a 3:1 molar ratio we would need 37.5 ng.
 That would be a total of 50 ng vector + 37.5 ng insert, for a total of about 88 ug of DNA
 If we want to do more than one reaction, we should make more insert – lets say 200 ng.
If the 500 bp piece is coming from a 4500 bp plasmid, how much DNA do we need to digest to
get 200 ng of the small piece?
See if you can figure this out, before you go on to the next page to get the answer:
OK the answer is 1.8 ug.
500bp is 1/9 of a 4500 bp plasmid. Therefore to get 200 ng you need 9 x 200 ng, or 1.8 ug.
That’s the logic. Of course it sets up as a ratio/proportion:
500 bp
=
4500 bp
200 ng
4500 (200) = 500 X; X = 4500 (200) = 9000/5 = 1800 ng
500
x
Be sure you double check the logic to make sure you did not turn the ratio upside down.
Remember the larger the molecule the more grams you need for a given number of moles or
molecules (or in DNA, fragments).
Now, if we cut 2 ug, and then run it on a gel, and cut out the band (200 ng worth) and then purify
it out of agarose, we stand to lose some – maybe a lot. We also need to make sure we have some
to “spend” on quantitating it after purification, so that we can in fact know its concentration and
set up a reaction with the right amount of DNA. So I usually make 2-5 times what I really need
– that way I can afford some losses. I do not want to go through this whole elaborate procedure
and end up with not enough DNA.