Hi there,

The causes can be very diverse!

What do you mean by unsuccessful ligation : no clones at all or no positive clones? If no clones at all, the reason can be as trivial as a lack of competence of the recipient cells, an incompatibility between insert and vector ends or even as not using the right marker of selection. If you have clones but no positive, it may be that the linearized vector is not compatible with the insert (one of the two cloning sites being not properly digested) and that vector preferentially self ligates. 

Have you run the usual controls for transformation? Transformation of your recipient cells with a known amount of circular plasmid in order to measure the competence of cells.

Do you perform the ligation using the right molar ratio between vector and insert? In your case I would use in 20µL ligation mix, 100ng vector and 60 to 100ng of insert which represents a 3:1 to 5:1 molar ratio. Because of the blunt end, it might be worth forcing the dose of insert up to 10:1...

Are you sure of the quality of digest? Are you sure of the efficiency of the two enzymes in your digest? To check, just single digest the circular vector and spot the linearization on agarose gel.

Are you performing double or sequential digestion? How close are the two sites on the vector? If too close there might be interference between enzymes in a double digest mix... How close to the end are the sites of the PCR product? Some enzymes are sensitive to this but I don't remeber if BamHI and HpaI are (have a look in an old NEB catalog where this feature was indicated for restriction enzymes)

Have you gel purified the fragment? It's necessary to make sure you get rid of little bits of DNA susceptible to ligate with your digested products...

How do you quantify the DNA prior ligation? Nanodrop is the best...

How much of the ligation mix do you use for transformation? If too much it will negatively affect transformation efficiency. I normally use 5µL ligation mix for 50µL competent cells.

Hope you will find something helpful...

Have a look at this NEB page it will give you even more ideas:

https://www.neb.com/tools-and-resources/usage-guidelines/cloning-guide

Hi there,

The causes can be very diverse!

What do you mean by unsuccessful ligation : no clones at all or no positive clones? If no clones at all, the reason can be as trivial as a lack of competence of the recipient cells, an incompatibility between insert and vector ends or even as not using the right marker of selection. If you have clones but no positive, it may be that the linearized vector is not compatible with the insert (one of the two cloning sites being not properly digested) and that vector preferentially self ligates. 

Have you run the usual controls for transformation? Transformation of your recipient cells with a known amount of circular plasmid in order to measure the competence of cells.

Do you perform the ligation using the right molar ratio between vector and insert? In your case I would use in 20µL ligation mix, 100ng vector and 60 to 100ng of insert which represents a 3:1 to 5:1 molar ratio. Because of the blunt end, it might be worth forcing the dose of insert up to 10:1...

Are you sure of the quality of digest? Are you sure of the efficiency of the two enzymes in your digest? To check, just single digest the circular vector and spot the linearization on agarose gel.

Are you performing double or sequential digestion? How close are the two sites on the vector? If too close there might be interference between enzymes in a double digest mix... How close to the end are the sites of the PCR product? Some enzymes are sensitive to this but I don't remeber if BamHI and HpaI are (have a look in an old NEB catalog where this feature was indicated for restriction enzymes)

Have you gel purified the fragment? It's necessary to make sure you get rid of little bits of DNA susceptible to ligate with your digested products...

How do you quantify the DNA prior ligation? Nanodrop is the best...

How much of the ligation mix do you use for transformation? If too much it will negatively affect transformation efficiency. I normally use 5µL ligation mix for 50µL competent cells.

Hope you will find something helpful...

Have a look at this NEB page it will give you even more ideas:

https://www.neb.com/tools-and-resources/usage-guidelines/cloning-guide

Hi,

in case of complicated (or rather unsuccessful) cloning I like to do a TA-cloning of the PCR product first and subsequently cut my insert out of this vector and clone it in the final (or intermediate) one. You get rid of the digest-problems which are due to restriction sites to close to the product ends. Furthermore once you have a positive TA clone which has a mutation free insert sequence you can always use this vector for you cloning attempts without endless repetition of the PCR.

Best,

Andreas

Thanks everyone for your response!! Let me answer each of you separately with more information about this ligation we've tried so many times: 

Andrew J King thanks for you comments, let me answer you

a)  yes, we've added a few nucleotides for that reason

b) No, we didnt dephosphorilate the vector. We didnt think it was necessary given the ciscumstances of the blunt + sticky end... We didnt have any religate at all though!

Andreas Finke that is good idea, thanks for sharing it! I will take a look at some protocols and I will take that into account if we are not succsselful with using the enzimes

Dominique Liger, thanks for taking your time to help us with this problem, let me clarify this situation.

By unsuccesful ligation I mean that most of the time we dont get any colonies at all, not even religate. We got a few colonies once (with ratio 10:1) but when we checked digesting the mini prep, they were not what we expected. The competent cells were ok, because we had our positive control perfectly and negative as well. They were ultracompetent cells that we made a few time ago, but they are still ok. 

The molar ratio that we used was 3:1, 5:1 and we even tried 10:1. In the last ratio we obtained those colonies that were not our clon.

The digestion is Ok, because we did the corresponding controls. For this ligation we had to do a double digestion. We performed this digestion on the vector, and on the other side on the purified-from-gel (using kit) pcr product insert. Then, the vector was purified from gel and insert double digested were purified using phenol-cloroform protocol, because neither of the two enzimes could be heat inactivated. We quantified by Nanodrop and was ok. We used 20 uL mix and transformed 100uL ultra competent cells.

We will change the primers and try to use other enzimes so that we get two non complimentary sticky ends, but we are limited in the enzimes that we can use becuase in the MCS of the vector we have other sites for enzimes that we need to use to clon another fragment. (and then clon a 3rd fragment into another region of the vector).

I really appreciate that you have taken your time on this case! Thanks! 

HI Marcos,

Why don't you try SLIC (sequence-ligation independent cloning). Just in case you keep failing with your current approach. For the SLIC approach you need to do the following: 1) amplify your insert with primers which at their flanks contain 30 nucleotides complementarity to your vector including whole restriction site blunt or sticky (to restore it), gel purify the fragment, 2) cut only your vector with the restrictases and gel purify it, 3) mix your vector (say 100ng) with your insert (100ng) and T4 DNA polymerase (just minute amounts f.e. 1unit of NEB T4 DNA pol - M0203) in respective buffer (I am using buffer G from Thermo) without any dNTP in final 30ul, incubate 30 mins at RT, at this step polymerase eats out 3' ends, 4) place in ice add one only one dNTP to final 1mM incubate for further 30mins at 37C 5) transform your bacteria. Prepare also a control without the insert. In my current lab IBB PAS Warsaw and in my hands the approach works well also for more difficult cloning of longer insert than vector and somtimes it works for 3 fragment cloning.

original methods paper: Li, Elledge 2007 Nature Methods Harnessing homologous recombination in vitro to generate recombinant DNA via SLIC

Best and good luck, Zbig

Hi Marcos, many correct things have been said and you have exhaustively answered to the good points posed by Dominique. Thanks to all this I still see one procedural flaw, and I would give you two additional suggestions.

The flaw concerns your purification of the insert after digestion. It is not sufficient to extract with phenol/choloroform and Ethanol precipitate the digested insert, since you will also precipitate the tiny ends that you have digested away from the insert and they will efficiently compete to religate with insert and vector when you set up the ligation. While it is not necessary to gel purify your PCR fragment prior to digestion (ethanol precipitation or a spin column will be sufficient to desalt and remove the polymerase), it is essential that you gel purify AFTER digestion to remove the small endings (and possibly sort out any unspecific you had in the PCR reaction).

The suggestions: first one concerns the ligation. A sticky end and a blunt end ligation have very different kinetic requirements. The sticky end needs slow kinetics to favour the annealing of the ends themselves, while the blunt end needs fast kinetics to try and ligate the very transient encounters of the blunt ends. For this reason I had best results, when mixing the two approaches, first ligating O/N at 16 °C and then transferring in the morning the reaction at r.t. (the classical ligation in the drawer!) for the whole day, transforming in the late afternoon.

Finally, I would pick-up the suggestion of seemless, ligation-free cloning, of the last post, but suggest to have a look at NEB's Gibson Assembly or Clontech's InFusion approaches. I have to say with large inserts or large plasmids they work very efficiently, and although they appear more expensive to start with, in the end avoiding the repetition of the cloning so many times turn out to be much cheaper...

Good luck.

Hi Marcos,

For this size of inserts we usually use mega-primer PCR. Reliable, doesn't need a lot of material (2PCRs and a DpnI digest) and always works!

good luck,

Christine

We have successfully cloned >10 kb fragments onto various sized plasmids (3-7 kb). Your combined insert-vector size is 9 kb so this should be easily possible.

You have a fairly comprehensive, excellent  answer from Dominique Liger above but I will add a couple more points:

1. If you are gel purifying your digested fragments, then do not vortex the DNA at any stage. Vortexing the gel fragments of the resin with the DNA will cause it to shear and reduce yield.

2. Once you add the ligation to competent cells, do not vortex at any stage. 

3. If your competent cells are home made, try using the rubidium chloride preparation method. It seems to work better for larger constructions. This method is available from many sources e.g. http://www-cryst.bioc.cam.ac.uk/hyvonen/methods/competent_cells

Good luck!

What's the ligase you used? Just a reminder, blunt end ligation is not efficiently catalyzed by T7 DNA Ligase. So you'd better use T4 ligase.

Lin

Thank you so much for all your replies ! They were really useful indeed! We are evaluating changing the approach now!

First if you do not see any colonies at all, then you should use more DNA, or better cells. A working ligation means that you spread 10% of the suspension on an agar plate, and see 20-25 colonies in the negative control reaction (ligation without the insert). Usually that way I see more than two hundred colonies on the ligation If you do not see them at all, that means something is not working right. Try the same reaction with more DNA!

The phenol extraction is quite unneccessary, and most likely reduces efficiency, if any trace amounts of phenol is left it can damage the proteins. You should digest the insert and plasmid, run the DNA on an agarose gel, and cut out the bands using a blue-light illuminator (a common problem is that using an UV illuminator damages the DNA itself, so if you spend a lot of time cutting out the bands that can be the problem) and simply isolate DNA with a cheap silicagel matrix kit, there are many on the market. After this treatment the insert and the vector are ready to be ligated.

The ligation can be made to work, but the blunt and sticky ends need different temperatures. First you should ligate the blunt end at a lower temperature ON and then add fresh enzyme and ATP and ligate it again at 18C for 2h.

All the above colleagues has covered most of the point facing in ligation or problem associated with it. My question is why you not ligating at one restriction site, i know its cloning efficiency is fewer then double digested ligation. As of my cloning experience I cloned in single restriction position and PCR gel purified insert after Dpn1 Digestion, 1:2 molar ratio vector:insert was always successful with 50ng of vector in total volume of 20ul o/n at 16oC or you can try some other ratios as well. For transformation, only using 3-5ul of ligated reaction in 50ul of competent cells. 

Best of Luck

Another thing that popped into my mind: You can add PEG to the reaction to increase macromolecular crowding to facilitate the ligation of the blunt end.