Please specify the vector, REs that yu are using and the length of the insert. 

SD

What is your insert:plasmid molar ratio? It sounds to me like it could be too high.

If you go over 5:1, you run the risk of "capping" both ends of your cut vector with individual copies of your inset, which would prevent recircularization of the plasmid.

Also, extra insert DNA can sometimes contribute to a carrier effect in which the slight molecular crowding afforded by the presence of higher amounts of insert DNA can encourage carryover singly-cut vector molecules to recircularize at a higher rate than they do in the no-insert controls.

It kind of sounds like both of these things are happening in your reaction. What method are you using to quantify your DNA prior to ligation?

- For ligation, the following molar ratios (vector: insert) should be used:

1. two 'blunt' ends (1:5)

2. one 'blunt' end, one 'sticky' end (1:4)

3. two 'sticky' ends (1:3)

- Make sure that your vector has been completely digested before setting up the ligation reaction.

- Make sure that your ligase buffer has not undergone too many freeze-thaw cycles. If it has, you may need to add some ATP (0.5 mM, in my experience) to your reaction.

Even if there are a few clones with the insert, this should be enough for many applications (except for library construction) if you have a good method to screen them and identify the useful ones.

Anyway, if you want to improve cloning efficiency, you can

1. If you are using a PCR amplified insert, check the primer design and  make sure that there are additional nucleotides flanking the restriction sites to allow proper insert digestion.

2. Optmize your digestions of both vector and insert  by using more enzyme and longer digestion times.

3. Improve the separation of digested from undigested vector by purifying the digested band from an agarose gel after each digestion.

4. Try different insert-vector ratios during ligations to determine the optimal one.

5. Dephosphorylate the vector after the second digestion.

It seems that one of your RE is not working efficiently. Hence, put Restriction digestion reaction for vector and add phosphatase so as to avoid vector vector ligation in case you get single RE digested vector. Elute the vector after gel electrophoresis. Now proceed with the ligation reaction using RE digested 1:3 molar ratio of vector and insert.

Thank you everyone for useful comments.

I am trying/failing in multiple cloning strategies! So here is one example - I am using MfeI and NotI (6/8bp cutter respt.) in pCAG vector (~5.5Kb). The insert is PCR amplified with extra 3-5 bp attached to improve RE cutting at the ends. Digestions are being done overnight with 10U of enzyme for every 5ug of parent. I gel extract both insert and vector after digestion. Most of cloning strategies involves digestion of vector at multiple cloning sites without release of a large insert; hence gel separation of cut vs uncut is not feasible

I am definitely considering the molar ratio to be the problem. Our nanodrop has been erratic and what I am considering to be a 1:3 ratio might be way off! Will try to titrate and retry ligations.

@Govinda: Very interesting to hear about 'molecular crowding'. Had no idea. Thanks!

Will update you guys after I try these suggestions.

-Anand

Gel separation of cut /uncut is possible and recommendable, at least after the first digestion (its better to do both digestions separately), even if the released fragment is small. Electrophoretic mobility of the digested vector usually changes and even if you don't see the undigested product, gel isolation of the major digested band should help, by removing traces of undigested vector that can have  a big influence on the results. The enzyme amount can also be increased, it was not so high.

About the molar ratio, its difficult to predict the ideal one, and there could also be concentration estimation problems.  That's why its good to try several ratios for difficult cloning.

Hi Anan

A couple of months ago I strugled with cloning too, so I would like to give you some of the tips that were given to me:

- Plasmid migration will be different wheter it is cut or not, with vector extracted from bacterias via Miniprep the different is very noticeable. 

- Nowadays RE are very efficient, multiple digestion works fine. 

- If you decide to purify the digested inserts/vector, take into account that UV exposure may harm your DNA.

- Take Nanodrop concentrations as.... orientative, ours give us about twice the concentration of Qubit's measurements ...

- Test your ligase buffer, it may have run out of ATP. 

- Test 1:3 and 1:5 molar ratios, the NEB calculator is really useful for it.

This may sound unscientific... but I got the best results when I knew that everything worked fine separately and started to only purify (and measure after it) my PCR products (without gel electrophoresis), the less I touched after digestion, the better efficiencies I got. 

Once again thank you all for your generous time in helping me troubleshoot.

I have an update: I got my first successful result today!

I SAP'd my vector and that made all the difference. Clearly the migration of single cut parent was far too similar to the double cut plasmid and I was cutting out both together. I was just flummoxed by the absence of colonies in the vector only ligation controls. However, given the 'molecular crowding' phenomenon described above @Govinda - I might be able to rationalize my results! I did try three different ratio from 1:1 to 1:10. Surprisingly, 1:1 gave the best results! Means our nanodrop is completely out of wack or this is a oddity. Will be testing more cloning strategies and will give a final update within the next few 2 weeks.

tl;dr: SAP your vector if you are seeing strange cloning outcomes.