I'm trying to clone a ~500 bp gene into the pET28a plasmid.  I first cloned this gene into the pCMV6-AC-DDK plasmid (using AscI and XhoI) to incorporate the DDK tag.  This was successful verified by sequencing.

I am following the same basic procedure for amplifying the gene + DDK using primers that add NcoI (N-term) and NotI (C-term) sites but I am not having any luck. I have double checked my primers and they appear to match up with the regions I am interested in, and the Tm's appear to be ok.  I purify after PCR and then do a double digest for 4h (at least 6 bases are included so the restriction site isn't at the very end).  After the digestion I gel purify and I see a nice band around 500 bp.  The pET28a was digested similarly and again yields a nice band on a gel. 

To ligate (T7) I have been using an excess of the gene insert (approximately 4:1).  I have tried 25 C for about 20 minutes followed by overnight at 4C.  I have also tried ligating using a custom PCR routine that is closer to the Tm of the sticky end regions (10C 1 min, 14C 1 min, 25 C 1 min; x60, followed by a 10C hold).  I transformed 4 uL of both of these ligations into 50 uL of XL10 Gold cells (fresh) and plated the entire transformation reaction.  I used the uncut pET28a as a control and saw many colonies with it.  

I am fairly confident that I have amplified the correct PCR product because I see a solid band on the gel.  Should I try to change the ratio of insert to plasmid in the ligation?  I am transforming a larger amount of the ligation (4uL) which might effect the efficiency of the transformation but would it completely inhibit it?     

Any suggestions on where to trouble shoot or what the most probable cause of failure might be? 

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