Troubleshooting/Restriction Digest and Ligation

Now that you know your transformation efficiency and it's above 1 x 108 CFU/µg DNA, we can work on other possible problems if you're still not getting great results from your cloning. This page is focused on common problems researchers have with ligations and restriction digests.

Restriction Digest Troubleshooting

Inefficient Digestion

You should never assume that your digest worked as expected. It's always good practice to check a small amount of your digested product on a gel prior to ligation to make sure your DNA was properly digested.

    Run a gel: After you cut your DNA (both insert and backbone), you should check the size on a gel. Run a DNA agarose gel with your digested plasmid alongside a lane of the uncut plasmid. Your uncut plasmid should appear to run smaller due to the supercoiled nature of uncut plasmid. Your cut plasmid should run higher since it's no longer supercoiled after being cut by the restriction enzyme(s). You should do this for your insert as well, especially if it's a segment of DNA you are cutting out of one plasmid to insert into another.

Ligation Troubleshooting

Negative and Positive Ligation Controls

It's easy to forget or skip controls when you're doing restriction digests and ligations. I strongly urge you to always run controls for these reactions as they will give you a lot of information and allow you to more easily troubleshoot your cloning problems. By running a few extra reactions, you can potentially save yourself many hours and days worth of troubleshooting later. Due diligence when it comes to cloning is well worth the effort, I promise you!

When comparing your backbone + insert ligations, you ideally want to see many more colonies than from the backbone control ligation (cut backbone with ligase, in table below).

Control Ligase Results
Cut backbone no (-) Colonies will give you an idea of the background due to uncut vector due to inefficient restriction digest of the backbone
Cut backbone yes (+) Colonies will give you an idea of the background due to the re-circularization of cut backbone
Cut insert yes (+) Colonies indicate contamination of intact/uncut plasmid in your ligation and/or transformation reagents
Ligation Ratios

Generally, a 3:1 insert:backbone ratio will work well for two-part BioBricks assembly. You can also try other ratios (1:1, 2:1, and up to 7:1) if the 3:1 fails to yield good results. New England Biolabs has a useful online Ligation Calculator that can help you determine how much of your insert DNA you need to add in various gram values.
In order to use this tool, you need to know:

  1. the length of your Insert DNA
  2. the length of your Vector DNA (or backbone)
  3. the amount of your Vector DNA you have for your reaction (labeled Vector DNA mass). This is most often entered as nanograms or micrograms.
You can set which scale you want to work in (example: nanograms or micrograms) and it will give you a series of different ratio outputs you can try to optimize your ligation if the 3:1 ratio fails for you.

Too many colonies

Sometimes with ligation reactions you can end up with a lawn of bacterial growth where it's impossible to select a single colony. While you may think this means your reaction worked really well, it actually indicates a problem with your restriction digest.

  1. No antibiotic: The most common cause of a lawn of bacteria after ligation is plating the transformation on an agar plate with no antibiotic added. This will allow any cells, with or without your ligation product, to grow up overnight on the plate.
  2. Uncut backbone: Another possible cause of too many colonies is from having a large amount of uncut plasmid backbone in your reaction. This makes your transformation in essence a plasmid transformation and you get far too many colonies on your plate. Remember this is one of the controls you should run for your ligations - setup a ligation reaction with your cut backbone without ligase. This control will tell you if large amounts of uncut plasmid is the cause of your problem.