Usage of Phosphorylated Linkers

Protocol

1. Dissolve the linker in 100 μl of distilled water to give a concentration of 60 pmol/μl.
2. 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.
3. 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).
4. 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.
   Centrifuge.
   Dilute concentrate to 2 ml and centrifuge.Repeat.
   Recover concentrate by inverting concentrator and centrifuging at 300-1000 x g for 2 minutes.