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!