Ligation theory is complicated and confusing. Elaborate calculations can be
made to optimize the degree of circle formation versus concatamers (see Maniatis), but
in practice we seem to get what we want without going to this
trouble.
It is worth making
the calculations in special cases, e.g. in ligations for making libraries, etc. where
efficiency of the desired vs side reactions is extremely important. For the simple
constructions we usually do, the following protocols work well:
Ligations can be done at room temperature for anywhere from 30 minutes to overnight depending on what is most convenient.
We have different protocols for ligating blunt and sticky ends
Sometimes dephosphorylation is useful to prevent a DNA from closing on itself in
a ligation reaction. There are two methods for doing this BAP and CIP.
There are two situations where we generally use blunt ended ligations. It can be
fairly easily accomplished when you can select for something, e.g. antibiotic resistance
conferred by the presence of the fragment you want to blunt in. The other situation in
which blunt ligation is used is for the addition of linkers. Here the reaction takes place
in the presence of 20 to 100 fold molar excess of linker over vector or fragment, in a
small volume, at high enzyme concentration and at room temperature (25oC). We add
PEG 8000 to a final concentration of approx 10% as it has been demonstrated that this
form of macromolecular crowding greatly enhances blunt ended ligation efficiency.
Typically the reaction with linkers goes for 3 to 6 hrs.
A common problem is ensuring that the pH of the ligation reaction is approx 7.5
to 8.0 before addition of T4 DNA ligase. Check pH on 6 to 8 pH paper using approx .25
ul droplet of ligation mix, and bring the pH up (or down) as necessary by adding 2 M
Tris base (or HCl) in the minutest possible quantities. When checking pH this
way you have to check it immediately on touching the pH paper as it will fade in
seconds, given these small volumes.
A typical linker ligation reaction: 30 ul reaction, assembled on ice.
Don't forget to check the tips
Plasmid DNA (linear gel purified) this volume assumes you started with
10 ug, digested it, gel purified it and it is now in 40 ul of TE.
2uL
CB 10X T4 DNA ligase
3uL
EDTA 2mM (chelates heavy metals)
3uL
PEG 8000 (40% weight/vol)
8uL
linker
1uL
*water
12uL
T4 DNA ligase (high concentration)
1uL
*now check pH using 0.25uL increments of ligation mix and adding the tiniest droplets of Tris base 2M to adjust.
For a pBR322 sized plasmid linearized, 4-5ug is one picomole, therefore, approx.
0.1 pmoles of DNA to receive linkers is present in the reaction, if the vector has simply been linearized. If more fragments are present, use this ratio to increase the amount of linker and ligation volume correspondingly.
Vortex, spin down and incubate at 25ºC (room temp) for 3 to 6 hours or overnight.
Add 3uL of 1M MES pH 6 to your entire ligation reaction (to buffer the pH down
the competent cells seem more sensitive to pH when the ligation contains PEG.
You can transform up to 10uL into the appropriate E.coli strain (SURE cells are best for standard cloning).
For sticky-end ligations (eg.: a vector cut with NcoI and PstI and insert with the same ends), the best ligation conditions are those in which there is a 3 to 4 fold molar excess of insert over vector.
For a vector of approx. 3Kb and an insert of approx. 0.5Kb, if you start with 2.5ug of vector DNA and 5ug of
the plasmid you are going to cut the insert DNA out of and you will have approx. equimolar recoveries of your vector and insert DNA after
gel purification.
If you have multiple steps in your cloning, such as phosphatasing or Klenow reactions, you may lose some DNA and may need to compensate for this by adding more insert or vector to the ligation reaction (depending on whether it is the vector or insert which has undergone the multiple steps). In this case it is sometimes helpful to start out with a little more DNA (eg. 10ug).
You can then add a 3 to 4 fold molar excess of insert in your ligation reaction.
A sample protocol follows - don' forget to read the tips:
Ligation reaction (10 uL)
Vector DNA (gel
purified)
0.5uL
Insert DNA
2.0uL
CB 10XT4 DNA ligase
1.0uL
EDTA 2mM
1.0uL
*water
4.5uL
T4 DNA ligase (0.5units/uL)
1uL
*check pH with 0.5 , add small amount 2 M Tris to adjust to 7.5
an equivalent molar amount of
unligated vector to determine the amount of residual (presumably uncut plasmids)
contaminating the supposedly totally linearized vector DNA.
If you get very few
colonies in the third case, very few in the second case and greater than three fold
number of colonies in the ligation reaction with insert, you will probably get what you
want. Remember to screen for orientation if the two sticky ends were the same.
It is common practice in the lab to gel purify both the insert and the
vector. However, it is usually not necessary to gel purify the
vector. In fact I rarely did. I just heat killed the enzyme and
added 0.5 ul of the restriction digest to the ligation reaction.
If the fragment that you cut out of the vector contains an unique
restriction site (not in your desired product anywhere) then I usually add
0.2 units of that restriction enzyme to the ligation reaction for 15 minutes
before I transform bacteria.
If you don't get very many colonies on any of the plates, you can try transforming all of
the rest of your ligation mix. To do this, you will have to add 0.75 ul of 1M MES pH 6 to
the ligation first to lower the pH so as to not kill the bacteria.
