Hi Everyone,
(Warning: A bit of a long post due to the necessary background information, sorry in advance.)
So I am in the process of creating a phage display library for my dissertation research, and I have been having major difficulties in producing a significant number of E. coli colonies after electroporating my final ligated product (a 5kb phagemid backbone with a ~800bp scFv insert) into electrocompetent E. coli cells (strain SS320) that I made and tested with proper controls myself. I have been doing months of optimization, and believe that I am close, but recently realized where a potential issue may be that has been hiding right under my nose. During the initial digestion of the phagemid backbone, for which I use Not1 and Spe1, I found that I also needed to treat with calf intestinal phosphatase (CIP) in order to remove the leftover phosphate group located on the resulting sticky ends. The reason for this being that there was still a significant amount of background (colonies containing undigested, supercoiled phagemid) still present when a portion of the digestion reaction was transformed into the previously mentioned E. coli strain. As a quick side note, I understand that Not1 and Spe1 have different recognition sequences that when cut shouldn't pair with one another and re-ligate, but for whatever reason the addition of CIP helped eliminate this background. Moving on, because of the CIP treatment, I know that there will still be nicks in the DNA backbone even after successful ligation of my insert (one on each end), which was also digested by Not1 and Spe1. Could these nicks be preventing my ligated product from supercoiling and, thus reducing its ability to be transformed into cells. As a another side note, I have looked into and tested a nearly exhaustive amount of other possible variables all along the entire cloning process that could be resulting in these poor transformations, and I believe I am narrowing in on the potential cause and this particular issue is one of them. In short, if anyone could give me any insight as to whether or not this nicked DNA backbone could be the reason for awfully low transformation efficiency I would very much like to know so that way I can look into using a different restriction site within the MCS of the phagemid that wont require CIP treatment. Thank you all so much in advance for any and all help :).
Patrick,
I think that nicked circles and covalently closed circles should transform with similar efficiency. I can't say that it's 100% identical but I don't think that it is significantly different. Also remember that if you are creating covalently closed circles with ligation they are not necessarily supercoiled. You are just closing a relaxed circle. You would need the action of a topoisomerase to add supercoils.
However your results make me wonder whether the explanation of your results is what you think. As you know, NdeI and SpeI should not ligate but since Cip treatment is reducing the background you have concluded that the dephosphorylation is stopping this side reaction. But there might be other answers. For example, you might have a small percent of DNA that is singly digested with either NdeI or SpeI which upon ligation will recircularize at a high frequency. This background might be being reduced by CIP. Or there may be a side reaction taking place (maybe your CIP has tiny amount of Exo contamination) which is reducing the background.
So an experiment you could try: take some of your double digested DNA and ligate it (without CIP treatment), heat kill the ligase and split it into 3, digest one aliquot with NdeI, one with SpeI and leave one. If you have improper ligation products then all 3 should be equal since ligating the sites together does not recreate either site. However if these are ligation events of incomplete digestions then the transformant numbers will go way down upon digestion compared to the control since native sites for Nde and Spe would still be present.
Michael J. Benedik
Hello Mr. Benedik,
Thank you so much for taking the time to help me with this issue, and for your in-depth answers and explanations. I forgot to mention that the version of the phagemid (vector) that I am currently using already has a 800 bp "place holder" scFv present within the MCS that contains restriction sites for Not1 and Spe1 on its 5' and 3' end, respectively. When I run the digested product on an agarose gel, I can clearly see two bands corresponding to the linearized vector (~5 kb) and the scFv insert (~800 bp), thus confirming that proper digestion took place. I do not see any higher molecular weight bands above the 5 kb mark, which could possibly indicate the presence of linearized vector with the "place holder" scFv still attached at either restriction site due to being singly digested. Further more, the band containing the proper linearized vector at 5 kb is gel extracted and purified before being added to the final ligation reaction with new scFvs. With all that said, I don't believe its a digestion issue, but please correct me if I'm wrong or what other thoughts you have.
I am sure your plasmid was double digested (at least mostly). But if say 1% was only digested by a single enzyme and not both, you would never see it on a gel. However the efficiency of plasmid re-circularization is very high so even that 1% could give you lots of colonies. And depending on your gel, you may not be separating the 5kb from 5.8kb when you gel purify.
I am just skeptical that the NdeI and SpeI sites are ligating with any real efficiency so I'm trying to find other alternative explanations.
- Badges
- Science topic
- Similar topics
- Computer Science
- Research Topics