I want to anneal to oligonucleotides and then ligate them in a plasmid vector. The annealed dsDNA will have monoclonal ends, so that it will be ready for ligation into a plasmid vector, which will be cut by two restriction enzymes. The monoclonal ends of the insert will be complementary with the ends of the cut vector. I will order the two oligos and then anneal them, but the question is: do I have to order them with 5-phosphorylation modification? If not, the efficiency of ligation will be reduced?
Thank you in advance.
I have cloned dozen of annealed oligos into various vectors and none of them was phosphorylated. In ligation reaction I use 100 pm of the annealed double stranded oligo. Typically, I obtain 30 - 50% of plasmids with the inserts. The crucial point in the cloning of the oligos is the annealing step. Since oligos may form complex structrures that impede their annealing, the actual concentration of the double stranded oligo may be very low. For good annealing I use 1x buffer for Taq polymerase without Mg2+ and add 0.5 mM Mg2+ and 1 M betain/ The latter disrupts low-energy intra- and inter- nucleotide structures to ensure more complete formation of double stranded oligos.
Hi there,
You can run the experiment with non phosphorylated oligos which will generate a non phoshorylated insert. As your linearized vector will be 5' phosphate on both strands it will be OK : the situation is similar to the one where you ligate dephosphorylated vector with 5'phosphate insert (for avoiding self ligation of the vector). Actually, you need 5'phosphate for efficient ligation with T4 ligase. So what you will get from your ligation is a circular plasmid with a nick at each 5' OH end of the insert. This nicked plasmid will be then efficiently repared into the cell after transformation.
Hope it helps.
Thank you all so much. So I will order the oligos without the 5-phosphorylation. The problem was that the 5-P oligos are very very expensive! On the other hand, the molecular ratio 100 insert : 1 vector looks too much for me.. But I will try it. The typical 3:1 ratio will not work? Thank you again!
Hi there again,
Whatever the phosphorylation state of vector/insert, the 1:3 to 1:5 ratio rule is always valid! The dephosphorylation is just avoiding self ligation event...
Just a quick note to ask a question to Andrei:
Why should ligation work when vector is dephosphorylated and insert phosphorylated at 1:3 ratio and not work the other way around in the same condition (the limiting step for the ligation being the meeting of compatible ends, the reactions should occur at the same rate in the 2 experiments, shouldn't they?)
I would be very interested in reading some of the publication you mentioned. As I couldn't find any myself, could you send the reference please? Thanks!
Thanks very much for the pdf. The purpose developed in it is, as far as I understand, very different from the one here: the paper is trying to optimize the generation of a library plasmid from random oligonucleotides insertion therefore the very high ratio used is such to generate maximal diversity in the sequence inserted and avoid concatenation of the vector (which is only a concern when using single digestion)... Here it's "only" about cloning one unique sequence in compatible double digested vector where concatenation will not occur. So the diversity of event is much less important...
I have cloned dozen of annealed oligos into various vectors and none of them was phosphorylated. In ligation reaction I use 100 pm of the annealed double stranded oligo. Typically, I obtain 30 - 50% of plasmids with the inserts. The crucial point in the cloning of the oligos is the annealing step. Since oligos may form complex structrures that impede their annealing, the actual concentration of the double stranded oligo may be very low. For good annealing I use 1x buffer for Taq polymerase without Mg2+ and add 0.5 mM Mg2+ and 1 M betain/ The latter disrupts low-energy intra- and inter- nucleotide structures to ensure more complete formation of double stranded oligos.
I'm still not convinced by Andrei, sorry... You are talking about the cheaper experiment but what is actually the cheapest: the one with 1000 molar ratio excess or the one with 3 to 10 molar ration excess... And as Dmitry said annealing condition is essential. I also had a few cloning experiments with hybridized non phosphorylated oligos without any particular problem compared to the usual protocol...
Provided the vector is not dephosphorylated then you can ligate in normal (dephosphorylated) double stranded oligos. If the plasmid is dephosphorylated however (eg phosphatase treatment) then the oligos would need to have a 5'phosphate
I fully agree with Robin and Dmitry.
BTW, even if you need a 5' phosphate (e.g. dephosphorylated vector), ordering a modified oligo is an unnecessary expense. You can always phosphorylate it with PNK - one of the cheapest standard enzymes. It works better on double stranded, so anneal then PNK is most efficient.
The question is interesting from an academic standpoint, but have you considered just using a gene synthesis approach instead of ordering expensive single stranded oligos? It should be much less expensive. Just have a company synthesize double stranded DNA out past the next unique restriction site and then ask them to clone it into the vector for you.
Hi all,
Whether we should phosphorylate both vector and a PCR product for a ligation reaction or whether there is any need for phosphorylating both for improved ligation efficiency?
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