Hello everyone
I've been trying to clone a 1640 pb insert into a 4300pb vector. Transformation works great, after O/N incubation at 37ºC, lots of colonies are found both in the control and in the ligation petri dish. However, the efficiency rate is extremelly low with only 1 or 2 colonies (sometimes none) positive out of 20. Most of them have the empty vector with no insert inside, so they got the ampR but not my insert.
How can we improve this efficiency? What is the ideal vector:insert ratio? I've been trying 1:3 with 50ng of vector, but I was told that too much insert for the vector will inhibit the reaction so to not perform 1:6 or 1:10 ratios. Is that really true?
Also what about tips like heating up the vector-insert mix 5min at 65ºC before performing ligation in order to avoid vector/vector and insert/insert constructs? Does that work?
By the way, we check that insert is fully digested in both extremes by performing an insert-insert ligation, in which we will observe several bands multiples of its own size (1600, 3200, 4800...etc) so I don´t believe the insert is the problem. We of course also check plasmid is fully digested/linearized by running a gel and checking that its band is higher than the undigested plasmid and all that.
Thank you very much
Bea
Try the following ;
I think a 1:3 ratio sounds about right, but don't see a significant problem with going higher. Is the insert flanked by the same restriction sites or by different sites, in other words is the vector prone to self ligation if a single site and did you dephosphorylate to reduce that?
You might also try a transformation with digested but unligated vector to see what the background is. If you even have a few percent undigested vector molecules that can swamp your transformation.
I would always start with 1:1 molar ratio then move from there. There are many things you can do to improve ligation. It's always recommended, if possible, to digest your vector with two enzymes that generate incompatible ends and I would recommend doing the digestion sequentially not double digest i.e digest with first enzyme, clean up then digest with second enzyme. This will prevent self-ligation of plasmid that normally occurs when doing a single digest.
The choice of restriction enzymes can have very strong impact on your results. Some restriction enzymes are a lot more efficient than others which will result in complete digestion of your plasmid which what you really want. Some enzymes are extremely unstable and cannot be used for extended digestion which may result in incomplete digestion which obviously will result in colonies that have no inserts.
Always make sure that you clean up the insert very well especially if it is a PCR product as primer dimers can and will ligate to the vector much faster and more efficiently than the desired insert due to their small sizes. Using PEG6000 can improve ligation but bear in mind that PEG reduces transformation efficiency quite drastically.
Best of luck
Sometimes problems with self-ligation arise even in case of using two enzymes with incompatible ends. The reason is more effective digestion by one of restriction enzymes and this situation may take place when restriction sites are very close to each other in plasmid . You can check required number of bases at the free end here https://international.neb.com/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments
Another possible explanation is that one of using enzymes works better in such conditions and a small part of linearized plasmid is cut only once. Self-ligation is more effective that's why even a small percentage of such plasmid can allow competition with your target product. Dephosphorylation of linearized plasmid before ligation could really help in such case.
Alexandra Dolgikh
I read the table about cleavage close to the end of DNA fragments provided on the NEB website. According to that table efficiency of NotI cleavage is 0% of short DNA fragments. I'm asking 'cause I have problems with my SacI/NotI cloning procedure. Digesting the vector with both enzymes simultaneously is 100% efficient as I'm cutting out an unwanted DNA fragment and purify the linearized vector from the agarose gel. I also simultaneously digest and gel purify my PCR product. After ligation and transformation I get no or just a few colonies on the selective LB plates. Do you suggest to add further nucleotides at the 5' end of my primer (upstream of the RE site)? Or what might be the problem? I also tried various vector:insert ratios, different ligation temperatures and durations, freshly made competent cells...
Thanks in advance!
Sylvia Kohler
NotI doesn't require addition of nucleotide before the restriction site. If the cutting efficiency of NotI and SacI is fine and you can clearly see the linearized vector on the gel. Then here are a few suggestions:
1. When you purify bands from gels, try to avoid using TBE gels as boric acid can be a potent inhibitor of T4 DNA liagse. Use TAE instead or even better if the band is not too large use polyacrylamide gel.
2. Purify the DNA with spin column from residual agarose.
3. Optional: to get the maximum recovery of DNA, upscale the restriction digestion reaction. Setup a 100 ul reaction and cut 4 ug of DNA or more. Clean up the reaction with plasmid miniprep spin column then elute in 50 ul. You can then load the entire 50 ul on TAE gel and proceed with gel extraction.
Best of luck
Waleed Albihlal, thank you very much for your comments. What I meant was that some restriction enzymes need additional bases to work properly on PCR products. The primers I use do not have additional bases at the 5' ends which may be crucial at least for SacI and/or NotI. I'm currently sequencing the positive clones which I mentioned before but I am sure to have false positives.
(1) I didn't know that boric acid can inhibit T4 DNA Ligase. In fact I used TBE buffer during horizontal electrophoresis. I will try TAE buffer.
(2) I use spin columns for purification (Macherey-Nagel).
(3) I will try that as well!
Best,
Sylvia
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