Hi shivani. If you're using a single restriction site, it would be helpful to dephosphorylate your vector. Religation of a vector linearized with a single cut is highly frequent. What you should NOT do inthis case is digest the ligation mix. The trick of digesting the mix can only be used when you're using two restriction sites for cloning and you're losing a site in the middle (which is also not present on the insert you introduce).
What do you mean by saying that you do not get the insert-vector to ligate? do you mean that you're not getting any colonies or that those colonies you get consist only of the vector re-ligated? 'cause the problems involved could be different in either case.

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hi, i certainly never had that problem with sap (at least not with sap from roche); for sure, the incubation times provided with the enzymes are not enough to get good de-phosphorylation; i usually dephosphorylate around 1µg of vector in a total vol of 50µl with 2U of SAP for 1h at 37°, add further 2U of enzyme and again 1h@37° before heat-inactivation at 65° for 15min. this is usually enough to get no or only a few re-ligations of the vector; ligations are then for 5' @ RT in a 2x ligation buffer, be it blunt-end or sticky ends, before transformation into e.coli. cheers.

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Well it happened to me precisely with SAP from Roche. When I asked to people in other labs to lent me an aliquot of their SAP and explained what had happened to me, two other people from different labs and using different brand of SAP told me that the same thing had happened to them occassionally. Since then, I've talked with three more people with similar experiences. But I agree that when using only one RE for cloning it is necessary to dephosphorylate. I've used SAP from Roche since then, and my samples survived, but now I prefer to use CIP.
Cheers!

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Hello,

I tried dephosphorylating my linearized, double-RE digested vector. I run it on agarose gel and I was surprized to see the band become heavier!

Before dephosphorylation, the size of my DNA is ~ 4 kb. However after dephosphorylation, it became so heavy that it remained on the top of the gel, very near the well! Can anyone explain what happened here.

Thanks!

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Sounds like a problem with your electrophoresis rather than your DNA. How did your marker migrate? What buffer did you use? How fast did you run the gel?

Also, if you load a lot of DNA sometimes you will get a smear that is at the top of the well.

Just things to think about-
slapolla

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My agarose gel elec. is totally fine. What I meant is that I ran them side by side: 1kb-marker, vector (phosphorylated), and vector (de phosphorylated). The vector (phosphorylated) migrated at the ~4 kb pace. However, the dephosphorylated vector becomes heavier (~ 20 - 25 kb) and stayed almost on top

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Hey guys i am not a Pro in Molecular Biology technics but ca some one help me out:
I am using EcoRI and bam HI to insert a vector 0.3kbp into pUC29 vector. It has a high copy number. EcoRI and BamHI sits were introduced using a primer in a PCR reaction. The PCR product
was purified, digested and the insert isolated from the gel. I did a Ligation 3:1 ratio and transformation with no control. i was sure the transformation worked well. I had very good plates with excellent colonies. I picked up the colonies and successfully analysed them using
EcoRI/ BamHI with a common buffer EcoRI buffer. Unfortunately i have bearly any inserts. Actually i am working with three inserts with same vector. Of all three inserts i pick 8 good clones. But in the end i have just 1...1 only one insert in 24 colonies. This is really ridiculous. This is supposed to be the simplest cloning, directional cloning....I have no clue. I have conducted this experiment twice.Please anybody, help a sinking person!!!!!

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Sorry my vector is pUC19 not pUC29

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Same thing here. By annealling 2 oligos, I got a 32bp short dsDNA with BssH II site at one end (The other end is blund end and end-labeled with HEX and FAM)to be ligated with DNA fragment with various sizes(from 171bp to 5386bp). All the fragments bear a overhang at one end generated by BssH II digestion and the other end is either blunt, 3' overhang or 5' overhang depending on the other restriction enzyme used together with BssH II. The 32bp oligo is 5' phosphorylated. I am currently running test ligation using 171 bp fragment from double digestion. If the 171 fragment is put together with 32bp oligo, I can see 3 prominent bands, corresponding to dimer of 32bp oligo, concatemers of the fragment (doulbe the size of original fragment), and the wanted ligation product(32bp+fragment). There are also some weak bands which fit to size of the concatemers of the fragment(tripled, etc.), but not a trace of the intact fragment. If the fragment is dephophorylated with antarctic phophotase from NEB, not concatemers can be seen, but the ligation efficiency is really low. In this case, still 3 prominent bands, dimer of 32bp oligo, intact fragment, ligated product. I use the ratio of 32bp to 171 bp fragment 5:1 and 3:1. Should I increase the ratio to 10:1? Any other suggestions to get high yield of ligation product? Can someone help me out?

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First of all, I am new to this forum. I guess you may have solved your problem. If so, you may let us know the reason for such low success rate in ligation.

If you have not solved the problem, I would suggest that you try producing a control (with the vector only). Ideally, there should not be any colonies. The other thing I would like to know is whether you have compared the undigested and digested plasmids. You need to know whether there is any cut in your plasmid. This may indicate whether your enzymes are working. Lastly, did you make sure that there are neither EcoRI nor BamHI sites in your inserts.

Regards,

Charles

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