OK, so I have been struggling to clone a 2 kb insert into a 9 kb vector.
Briefly, the steps that I did are as follows:
- insert: amplify using a Hi-Fi Taq from a vector, A-tailed using standard Taq, run PCR product on gel, extract from gel (promega SV wizard), T-A cloned into pCR2.1 TOPO, transform into TOP10, harvest plasmid, digest with EcoRI-HF, cut insert from gel, gel extraction, run on gel & Nanodrop, used for ligation
- vector: cut using EcoRI-HF (then deactivate the enzyme), Nanodrop & run few microlitre on gel to check (there was only one band, so digestion seems complete), Antarctic phosphatase treatment 15 min at 37oC deactivate 70oC 5 min, used for ligation
- ligation: I used a molar ratio of 3:1 for insert:vector. I used 3 femtomole (fmol) of insert and 1 fmol of vector in 20 microlitre reaction. Incubation 2 hrs at room temperature. Run on gel the following (lane order): pre-ligation mixture, post-ligation mixture, vector re-ligation (no insert, no phosphatase treatment), uncut vector. I attached the gel image.
Some possible culprits that I suspect:
1. the concentration of insert was actually very low after gel extraction, around 5 ng/microlitre, but if I run on gel around 10 microlitre the band was clear. However, in ligation mixture I put 15 microlitre of insert to achieve the molarity that I needed. This might be not ideal. Should I concentrate it first using ethanol precipitation or phenol-chloroform?
2. I did not see the ligation product in gel (see attached image). At least I should be able to see the insert band or the vector, but it seems that the DNA is disappearing? I did this gel couple of times from different batch of ligation reaction and always get similar results.
The competent cells are OK, it's still new from Invitrogen and tested using other vectors it's working.
What have I done wrong or have I missed something? I need help! Really appreciate your comments and inputs on this.