what is the molar ratio between plasmid/insert?

Hi, sorry should have added that too. I was using vector:insert rations of 1:1, 1:3 and 1:5 using 40-50ng vector. For the 1:3 ratio that worked out something like 40ng vector to 270ng insert. I though this might be too much DNA for the competent cells thats why I added 1ul. Using 1ul gave me some colonies while adding 5ul gave me now colonies.

I advise you to use a ratio of 1:7 or 1:10 (total 100 ng between vector and insert). Then you can use only 3/5 microliters of ligation to transform competent cells.

Im acually setting up ligations today. So will try this ration and see how it goes. Thanks!

Adding to the above comments - you could try adding PEG to the reaction mixture (say PEG-8000 at 5-10%) - it effectively increases the concentration of your mix and picks up the kinetics of the ligation.  I also always ligate overnight at 16 degrees, not 4 degrees. I'm not sure what competency your 'Bioline Gold' cells are, I tend to use super/ultra comp cells that are about 10E9 cfu per ug... Also, assume your ATP is nice and fresh too!

Hope that helps

is it possible to use cut-ligation?: cut the vector with smaI, fill up the sticky end of the ncoI(/ecorv) cutted insert, mix vector and insert 1:10 + 1/10 volume ligase buffer, 1 ul T4 ligase (neb), 0.5 ul SmaI, x ul water. incubate at 25°C for 1 hour. religated vector will be recutted and is available for new ligation. attention should be paid that there are no smai restriction sites within the insert. I often used this method, worked well in the past. good luck!

Why don't use Xma  instead of Sma?

Hi Richard, thanks for your response. 

I am using the neb quick ligation and blunt ligation kits that have peg6000 in think. I also try to do the ligation in the smallest volume possible (5-10ul) to help the vector find an insert. The competent cells I use are 10e9 cfu per ug. I usually aliquot my ligase buffer and defreeze one aliquot per reaction I set up. I also sometimes add extra atp from 10mM atp to make sure the reaction goes forward. None of this has helped so far and my ligation kits where brand new so I don't think the ligase was off. Either way I have cut one of my vectors and will try to religate it as a positive control for the ligase. Thanks for the help.

Hi Andrei, thanks for your response and apologies for not being more specific.

Is not a pcr product, both vector and insert are digested with a sticky and blunt enzymes.

The image you see is the digested products after gel extraction and purification. That's why there are no further controls. These controls I run during the separation of the digested samples. These controls look fine. I have tried few times now. You can clearly see that the vector and insert have Been cut well because it releases various fragments of smaller sizes compared to linearised or uncut vector. 

I usually dephosphorylate the backbone as you point out. But a senior investigator in my lab convinced me that if I cut with different enzymes it shouldn't religate. In theory he is right, however whenever I get colonies after transformation they always are religated vector to our surprise. Some people say that using phosphatases can inhibit the ligation, but don't ask me the specifics of why :s. I usually dephosphorylate the vector. The insert in theory doesn't need phosphorylation as it has been cut with enzymes. However, as I stated , I have also tried to dephosphorylate the vector and phosphorylate the insert with not much success either.

As for the ligations. I always carry the cut control with no insert and ligase, and the same reaction without ligase. No insert with ligase gives me 3-5 colonies sometimes and no colonies when I dephosphorylate the vector. the cut vector with no ligase gives no colonies, confirming no uncut present. You reduce your background with the dephosphorylation.

Let me know if I followed you correctly as I'm replying from my mobile. Thanks very much for all the pointers. What ratios would you do with such a big insert in such a small vector?

Hi Annett, thank you for the help.

Yes I thought of this method too, haven't tried yet. I think is a very cool method. If i

Sorry my post was too long Annette. Correct me if I misunderstood your method. 

Im cloning into a pentr vector that has a ccdb suicide gene in. Therefore I need to cut the vector with ncoi and ecorv to remove it and replace it with an insert. Having the ccdb should reduce background of linearised religated and uncut as the bacteria will die. I could do the same digestion and keep smai in the ligation so if reduce the rilagated vector (with no ccdb, I know it's weird with different re sites but it does happen and do get colonies). However, since i get very few colonies in the negative ligation control and usually no colonies or 2-5 colonies in the insert:vector ligation plates I thought perhaps is best to not be so stringent. Will give it a go though as I need to try everything. 

Thanks very much for the help.

Hi Petre,

There is no xmai/smai in my destination vector. So I cut blunt with ecorv in the backbone and with smai in the insert holding vector. I would use xmai too since it cuts at 37 and its sticky but can't in this case I think.

Thanks

Strangely enough colonies start growing on the transformation plates after 48h at 37C instead of 24h. I was wondering if this is because the backbone is kanomycin resistance. I let the bugs recover for a good hour with shaking. I was temptend once and picked some colonies after 48h but it seemed to be religated plasmid. Should I pick colonies after 48h or only stick to 24h always? colonies look just like normal ligation colonies. Sometimes after 24h you can see very little colonies growing. Incubator is set to 37 and not 30.

If you keep having problems I would consider trying Gibson cloning. Just PCR your insert (Or vector) with vector/insert overlapping flanking sequences, mix with the vector (or insert) at the recommended molar ratio and use the NEB Gibson cloning kit. It costs about $18/reaction but is incredibly efficient and worth the money in my opinion. For simple cloning you get literally thousands of colonies from 1/10th the reaction volume. Our lab moved over to this method for almost all of our cloning this year and it has saved us hours and hours of time due to its high efficiency and propensity to work first time.

Hi Daniel,

Thanks! I will consider that if I get another complicated cloning. FInally I got my cloning to work a couple of weeks back. I basically ended up using lots of ligase and it went in. i can provide more details if anyone is interested.

Thanks to everyone for their ideas and input they were very helpful.

Hi Patrick,

I also have the same problem, except my vector is 5000 bp and the insert is 2000 bp. I have alredy tried the same processes that You wrote mounths ago, and now I find this topic. Could you please tell me what was your solution?

Thanks,

Sari

@ Patrick Ovando-Roche how you got it can you share it now?