I need help with sorting through my current predicament. I have been trying to blunt-ligate my inserts for a week but nothing working, here are the specifics.
My inserts are about 400bp and using the pBluescript II KS+ vector, which is 3kb in size. I first linearized vector using blunt-ended RE digest (Hinc II) then gel purified, phenol/chlorof and Etoh/NaAcet. I run on gel and nanodrop quantification is about 43ng/ul.
My inserts have a T7 promoter (for in vitro transcription), are PCR amplified and were gel purified too. They range in quantity between 12ng/ul - 25ng/ul.
used a 3:1, Insert:vector ratio for the ligation rxns.
At the end of each extraction, i set up the following rxn for a one tube blunt-end ligation
Our protocol is as follows; combine cocktail/2 ligation rxns as follows
10X T4 DNA ligase buffer 3ul
linearized vector xul
T4 DNA polymerase 0.25ul
T4 DNA ligase 0.625ul
T4 DNA kinase 0.3ul
final volume 10ul
combine 5ul of cocktail and 10ul of insert + ddH2O. incubate overnight at 12-14oC.
Run 5ul of ligation on agarose.
no ligation worked. the no insert control had linearized vector fragment only and the rest had just insert and linear vector size fragments on the gel.
I have run out of ideas at this time, please help. What could i do differently with my protocol. I could use a TA cloning vector but boss wants me (grad student) to clone into pBluescript, with this we can then linearize for blunt ends again before in vitro transcription.