I used the following protocol with good results:
Dilute oligo to 1µg/µl. 1 µl top oligo + 1 µl bottom oligo + 2 µl annealing buffer (10 x buffer: 100 mM Tris pH8,5; 1 M NaCl; 10 mM EDTA) + 16 µl ddH2O. Heat for 5 min in PCR cycler to 95°C. switch of and let cool to room temperature for 3 hours. use 2 µl of the annealing mix and ligate with 100 ng digested vector in 20 µl reaction volume over night. Electroporate 1 µl of the mixture in DH10B cells and plate.
To check annealing I usually run 1 µl of the annealing mix on a 3 % gel. I also often get 2 bands in the unannealed oligo control but it still works if the upper band in annealed oligo is stronger than in the untreated oligo.
Thanks stardust. I'll try this protocol soon. Haha. But there is something that I would like to enquire first, when u turn off the PCR cycler, did u take out the oligo mixture to RT? Or you just keep it inside the PCR cycler? Then what about your ligation condition? 16C overnight incubation?