Bacterial transformation problems - (Oct/11/2013 )
I'm having trouble ligating my insert (Pyp1 promoter region, 1.6kb) into my plasmid (pRip42Pyp1C20SpkC, 8.1kb). I managed to make this plasmid using the same bacterial transformation procedure as I am using now (transforming Pyp1C20S into pRip42pkC), so I am unsure why it isn't working now.
I digested my insert and 5ug plasmid with Sph1 and Nde1. I initially digested my insert and plasmid with both restriction enzymes together in an optimal buffer, however when the bacterial transformations didn't produce any colonies, I decided to try digesting with each enzyme separately, with a clean up in between, and then phosphatase treating the plasmid.
I digested 5ug and 10ug plasmid with 1ul Sph1 and 1ul Nde1 separately (3 hours at 37C for each enzyme, with a clean up in between where I eluted in 40ul buffer EB), and then treated the plasmid with 1ul alkaline phosphatase for 30 mins at 37C. I wasn't sure how much DNA I would lose during the clean ups and digests, so this is why I tried digesting 5ug and 10ug plasmid. (Using the Nanodrop, I had roughly double the amount of DNA left after digesting 10ug plasmid as I did for 5ug plasmid).
As digestion with Sph1 and Nde1 was predicted to excise the nmt promoter from my plasmid, I ran the digested plasmid on a 0.7% agarose gel. As expected I got a band at 8.1kb to represent the linearised plasmid, and a band at 1.2kb to represent the nmt promoter. (It should be noted that when I digested the plasmid with Sph1 and Nde1 together and ran the digested plasmind on a 0.7% gel I still saw the excision of the nmt promoter, so this double digest was working but perhaps not to a great extent). I then gel extracted the 8.1kb band to use in the ligations.
I ligated 50ng plasmid with my insert at the ratios 1:1, 1:3 and 1:5 in 20ul (1ul T4 DNA ligase, 2ul 10xligase buffer). I calculated the ng of insert to add based on the size of the insert and of the plasmid, so for the 1:1 ratio I added (1.6/8.1) x 50ng = 10ng insert. I left the ligations overnight at 15C. The vector only ligation acted as a negative control as this is the linearised vector which shouldn't produce colonies.
For the bacterial transformation I added 100ul competent E. coli cells to the whole ligation mix (vector, and vector+insert 1:1 1:3 and 1:5). As a positive control, I added 20ul competent cells to 0.5ul undigested vector. These were left on ice for 30 mins, then heat shocked at 42C for 2 mins. I then added 1ml LB to the tubes and incubated at 37C for 1 hour. For the positive control I plated 50ul onto LB+Amp plates. For the negative control and vector and vector+insert, I plated 100ul onto LB+Amp plates. I spun these tubes (QuickPulse) to get a small pellet, then removed the supernatant to leave 100ul, which I then plated onto LB+Amp plates and called 10x.
I left these plates overnight at 37C and got no colonies on my vector or vector+insert, but did get colonies on my positive control, so I know the cells are competent enough. Getting no colonies on the vector plate was a good sign that my plasmid was linearised and correct, but getting no colonies on my vector+insert plate obviously wasn't great!
I then tried to repeat the ligation using 100ng plasmid with the insert at the ratios 1:3, 1:5 and 1:15 in 20ul. Following the same procedure as above, I still got no colonies for the vector+insert plates, but did get colonies on my positive control plate.
Any suggestions/tips/help would be much appreciated!
You do NOT know the cells are competent enough. There is a huge difference between transforming uncut vector and ligation product. The only way to know your cells are competent enough is to measure their competence by serially diluting plasmid to the 10 pg/ul level and calculating the CFU/ug of plasmid DNA number. It should be 10**8 to 10**9 CFU/ug.
Also, you are transforming far too much of the ligation mix -- I gather a 20 ul reaction in 100 ul of "competent" cells. You should be using more like 1-2 ul of the ligation mix in that volume of cells.
I would also strongly suggest you eliminate the CIP treatment, or at least try this reaction with and without it. You should not need it, and it often causes far more problems than it solves.