I have been trying to clone a ~900 bp Drosophila
gene into pUASTattB
vector. It is not so hard, as it seems, but it's just not working.
Here's what I have done: Insert:
PCR product (purified) of the gene with 5' NotI and 3' XbaI sites, incorporated while PCRing (with those sites on the 5' and 3' primers respectively). Vector: pUASTattB Double digestion:
Insert and vector doubly digested with NotI and XbaI (I know that XbaI is sensitive to dam
site overlapping, and I have checked and found that there is no dam
site overlapping in the vector.) I have gel purified and checked the products on a gel and found there is sufficient DNA in the purified products. I used0.5- 2 microlitres of each enzyme in a 50 microlitre of total digest volume. I have tried all sorts of incubation periods from 2 to 16 hours at 37C. I have also tried with and without phosphatase treatment of the digested vector. Ligation:
I have set up a ligation of 20-60 microlitre volumes with 3:1 insert:vector ratio and used 0.2-1 microlitre T4 ligase. I have also set up reactions of empty vector with or without ligase for negative controls. Incubations were at 14C overnight. Electroporation:
I have used 2 microlitres of the ligation mixture to electroporate 20 microlitres of DH5alpha electro-competent cells. Sometimes I have done salt-precipitation of ligation product and sometimes without salt-precipitation. But, never did I got colonies on my plates!! My negative control (mock transformed) and positive control (transformed with the uncut pUASTattB
vector) worked efficiently. But, no success with my experimental plates.
I have done these steps so many times by altering the protocols, incubation periods etc. I have used enzymes from NEB and a few times used ligase from Fermentas.
Please give me your suggestions.