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 ratio fails for you.
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.
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. 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.
Visit us at igem. Inefficient Digestion You should never assume that your digest worked as expected. Negative and Positive Ligation Controls It's easy to forget or skip controls when you're doing restriction digests and ligations. Ligation Ratios Generally, a insert:backbone ratio will work well for two-part BioBricks assembly.
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. With easy-to-use hardware and an open software platform, Opentrons automates manual lab work and empowers collaborative research for hundreds of life scientists. Opentrons is used by scientists at 90 percent of the top 10 largest pharmaceutical companies and 90 percent of the top 50 biology research universities.
In any experimental procedure, getting the controls right can save you a lot of work when things go wrong by allowing you to troubleshoot the source of the problem. DNA ligation is no different. In this article, we explain how to set up a ligation reaction with a complete set of controls, and use them to troubleshoot the cause of your ligation problems. Ligation reactions fall into two categories, depending whether you are trying to join blunt or sticky DNA.
Although the set-up is slightly different, the controls are basically the same for both types of reaction. In this article, we will focus on ligating a single insert into a vector.
The ligation reaction begins with a linearized vector fragment, to which an equimolar or molar excess of insert is mixed, along with DNA ligase in an appropriate buffer.
The ligase joins the vector and insert pieces creating a covalently closed, circular vector from the initial linearized DNA. I normally use 50ng of vector, then calculate the amount of insert required for one molar equivalent using the equation shown on the right.
So, my ligation mix for a 5 kbp vector and a 1kbp insert would look like this:. Adding a molar excess of insert can increase the chances of ligation, depending on the insert size, so it is often advisable to try a few vector:insert ratios. You can get more info on how ligation works here.
After ligation, competent E. The fact that E. The number of colonies obtained from transformation with covalently-closed, circular DNA depends on the competency of the cells and the amount of DNA present. Assuming we use the same competent cell prep for our ligation and controls, the number of colonies formed will be dependent on the amount covalently-closed DNA we have in each reaction i.
And now onto the controls. To orientate you, below is a summary of the controls we will focus on:. The vector preparation is a crucial part of the cloning procedure. If even a small amount of undigested vector carries through into the ligation, this will be transformed efficiently and will produce colonies.
More information on preparing vectors to reduce background colonies can be found here. Another source of background is re-ligation of the vector. If the linear vector has compatible ends e. A phosphatase treatment will reduce the possibility of re-ligation and reduce background, but it is often not perfect. Whatever the source, it is vitally important to know how much background there is in a ligation reaction. To determine the amount of background, a parallel ligation reaction should be set up, with the insert replaced by water.
This is control 1. Any colonies formed from transformation of this control will be the result of undigested or re-ligated vector, and subtracting the number of colonies formed from this control from the number of colonies formed from the ligation will give an idea of whether the ligation reaction has been successful.
The ideal scenario is that there are few or no colonies on the control plate and tens, hundreds, or thousands of colonies on the ligation plate. If your ligations are going well, control 1 is sufficient. However, if you are experiencing problems, the controls described below will help you pinpoint the source. Control 2 is similar to control 1, but without the ligase. Colonies formed from transformation of control 2 can only be the result of undigested vector, not vector re-ligation, since there is no ligase present.
If you are obtaining a high number of background colonies, comparing controls 1 and 2 will allow you to determine whether the problem is due to undigested vector or vector re-ligation. The other side of the coin as that you get very few, or no, colonies. This could be because of issues with the competent cells or transformation procedure, use of the wrong antibiotic, or problems with the ligation reaction, such as poor ligase activity.
The remaining controls allow you to distinguish between these possibilities.
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