Cloning trick: ligation of multiple inserts

[2013.02.26 Edit: A number of people are finding this through Google searches. I don’t have an updated post on the topic, but if you’re trying to assemble multiple DNA fragments then I suggest looking into Gibson Assembly. NEB sells* a dead-simple mastermix, which is a bit pricey per reaction (I just make my reactions half the size) but comes out to cheap when you take into account the cost of labor (so long as your PI values your time…).]

I’ve spent the last couple months building a plasmid library, and in the process I thought of a trick. Ligations, perhaps the worst part of cloning, are notoriously finicky reactions. The goal is to take several pieces of linear DNA, where the ends of the pieces can only connect in a certain way, and then use an enzyme (T4 Ligase) to sew them all together into one piece (in my case, a circular plasmid).

Figure 1. Ligase (2HVQ.pdb) rendered in PyMOL. Click to see a crappy animated GIF!

I needed to insert three fragments at once into a single backbone. In my ignorance (from my lack of experience) I thought ligating four fragments should work just as well as two, so I just threw them all together and ran the reaction. The result was a mess, and when I tested 40 different clones afterwards not a single one was correct. So I started adding them one piece at a time which, obviously, was going to take three times as long.

The next morning while in the shower (one of the best places for random ideas) I thought that, though most of what was in my test tube was not the desired product, there was most likely some tiny population that actually was the correct fragment. The problem was just that putting my products into E.coli and then trying to find the correct thing wouldn’t work if only 1/100 (or less!) of the ligation products were correct. But PCR is wonderful for making tiny amounts of DNA into large amounts, so I thought, why not just try to use PCR to get the correct thing out of the mess? Since each product is made out of some combination of the input fragments, the lengths would be discrete and therefore visible and, potentially, separable on a gel. I just needed to be able to copy up enough that I could see DNA run on a gel, and then I could just purify out the one that was the right length.

Figure 2. Each differently-colored fragment is being ligated into the gray backbone. The primers (black arrows) flank the entire insert.

I designed primers that would flank the combined insert and did PCR, and did indeed get a band of the desired length! After gel purification I was able to ligate the entire insert into the backbone without any trouble.

You might be asking, “Well isn’t this now a two-step process, just one less than the three required otherwise,  and so barely worth the effort?” Almost, except that there is no need to transform the multi-fragment ligation mixture. So it’s more like 1.5 steps. Plus, the ligation, PCR, gel purification, final ligation, and final transformation can all be done in a single day!

Though my labmates were surprised when this worked, it turns out I was a year too late to publish this idea :(. If you have access to Analytical Biochemistry, see the paper by An, Wu, and Lv called “A PCR-after-ligation method for cloning of multiple DNA inserts“. If you aren’t that lucky, I’ve already told you everything you need to know!

12 thoughts on “Cloning trick: ligation of multiple inserts

  1. The only problem with this apporach is that the PCR step could introduce errors into your product which could or could not be very bad (lets say a stop codon in your marker gene or gene of interest). You seem pretty savey though so you probably calculated the probability of an error based off the fidelity of your polymerase!

      1. I’ve done some large fragments (~10kb), but never in a situation where I needed perfect fidelity along the entire sequence. I sequence the portions that strongly matter (usually <1.5kb, so a few Sanger runs) and then rely on selection in bacteria or other functional assays to ensure that any backbone mutations don't matter.

  2. Awesome! Nothing saved me when I tried to do multiple insert ligation, not even fusion PCR. This seems to make my life hopeful again!

  3. Do i need to do restriction digestion again after PCR so as to ligate my desired product into the vector???

    1. Probably, depending on how you designed your cloning strategy. If you pool your inserts together for ligation without a vector, you’ll still (probably) need to trim the blunt ends. If you used a vector, you’ll end up with a huge pool of incorrect constructs that PCR won’t help you with. You could, however, transform this pool and then check for proper insert length in a large number of colonies. That would work fine if the proper ligation will represent a good fraction of your products. Otherwise you would indeed want to gel-purify and trim the PCR product that comes from the mixed ligation.

      If you’re just starting your cloning project, it’s worth looking into Gibson Assembly and planning your cloning strategy around that method.

    1. Depends. I usually just dephosphorylate if I have trouble getting it to work, or if I have a particularly small insert that will be in at high concentrations.

      But for multiple inserts, you definitely can’t dephosphorylate everything. At most it would need to be every other insert. I think. Top-of-my-head answer.

  4. I am trying to ligate two inserts (insert 1 cut with AgeI and ApaI, and insert 2 cut with AgeI and ClaI) and then amplify by PCR the ligated product with the same flanking primers, but it doesnt work. I made a gradient PCR but nothing. Do I need to add the vector to the ligation mix, since I only need the ligation product, or the vector helps to the right ligation?

    1. You shouldn’t need the vector, but I’ve never compared the with/without vector cases. So it may work better, I have no idea!

      Those enzyme overhangs should be fine. Did you gel-purify the digested fragments and make sure they were the right size? If you digest out of PCR products instead of out of a vector, you may need to add some bases to the 5′ end of the primer to give the enzyme something to hold onto.

      Finally, if you’re doing double-digests it’s possible that the buffer conditions were not suitable for both enzymes at once. I’ve gotten messed up in the past when using NEB’s high fidelity enzymes because they sometimes use different buffers than the regular ones. If you are using NEB enzymes, they’ve also recently changed all of their buffers, so it’s worth a double-check.

      Sorry I can’t be of more help!

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