Usage of Phosphorylated Linkers
- Dissolve the linker in 100 μl of distilled water to give a concentration of 60 pmol/μl.
- Ligation Reaction:
Ethanol precipitate DNA fragment(s).
Resuspend in 10 μl TE Buffer.
Add the proper amount of linker to give a 50-100 fold molar excess of linker to DNA end(s).
Add 2.5 μl of 10X Ligase Buffer (500 mM Tris-HCl pH 7.5, 100 mM MgCl2, 100 mM DTT, 10 mM ATP).
Adjust to 24 μl with water.
Add 400 units T4 DNA Ligase (NEB# M0202).
Incubate at 16°C overnight.
Heat inactivate reaction at 75°C for 5 minutes.
- Digestion Reaction:
Cool on ice after heat inactivation.
If the enzyme of choice requires high salt, adjust the salt concentration to the appropriate level with 5 mM NaCl.
Add 1-2 units of restriction enzyme/pmol of linker.
Adjust volume to 30 μl with distilled water.
Incubate at the appropriate temperature for the restriction enzyme for 4 hours.
Exctract the digest once with phenol:chloroform:isomaylalcohol (24:23:1) and once more with isoamylalcohol:chloroform (1:24).
- Removal of excess linker:
Excess linker may be removed by LMT agarose gel electrophoresis, a G-25 spin column, ultrafiltration or by PEG precipitation.
example using a Centricon 30 or 50:
Add 1 ml deionized water to the Centricon 30 or 50 sample reservoir.
Pipette reaction mixture containing DNA fragment - linker and non-ligated linkers into concentrator.
Dilute up to 2ml with deionized water.
Dilute concentrate to 2 ml and centrifuge.Repeat.
Recover concentrate by inverting concentrator and centrifuging at 300-1000 x g for 2 minutes.