The final step in the construction of a recombinant plasmid is connecting the insert DNA (gene or fragment of interest) into a compatibly digested vector backbone. This is accomplished by covalently connecting the sugar backbone of the two DNA fragments. This reaction, called ligation, is performed by the T4 DNA ligase enzyme. The DNA ligase catalyzes the formation of covalent phosphodiester linkages, which permanently join the nucleotides together. After ligation, the insert DNA is physically attached to the backbone and the complete plasmid can be transformed into bacterial cells for propagation.
The majority of ligation reactions involve DNA fragments that have been generated by restriction enzyme digestion. Most restriction enzymes digest DNA asymmetrically across their recognition sequence, which results in a single stranded overhang on the digested end of the DNA fragment. The overhangs, called "sticky ends", are what allow the vector and insert to bind to each other. When the sticky ends are compatible, meaning that the overhanging base pairs on the vector and insert are complementary, the two pieces of DNA connect and ultimately are fused by the ligation reaction.
The example below depicts the ligation of two sticky ends that were generated by EcoRI digestion:
Usually, scientists select two different enzymes for adding an insert into a vector (one enzyme on the 5' end and a different enzyme on the 3' end). This ensures that the insert will be added in the correct orientation and prevents the vector from ligating to itself during the ligation process. If the sticky ends on either side of the vector are compatible with each other, the vector is much more likely to ligate to itself rather than to the desired insert. If you are in this situation, it is important to treat the digested vector backbone with a phosphatase before performing the ligation reaction (phosphatase removes the 5' phosphate and therefore prevents the ligase from being able to fuse the two ends of the vector together).
Protocol: Standard Insert + Vector DNA Ligation
Before setting up the ligation reaction itself, it is important to determine the amount of cut insert and vector to use for the ligation reaction. The volume of vector DNA and insert DNA used in the ligation will vary depending on the size of each and their concentration. However, for most standard cloning (where the insert is smaller than the vector) a 3 insert : 1 vector molar ratio will work just fine. We recommend around 100ng of total DNA in a standard ligation reaction. Use a ligation calculator to easily quantify how much vector and insert DNA to use.
- Combine the following in a PCR or Eppendorf tube:
- Vector DNA
- Insert DNA
- Ligase Buffer (1μL/10μL reaction for 10X buffer, and 2μL/10μL reaction for 5X buffer)
- 0.5-1μL T4 DNA Ligase
- H2O to a total of 10μL
Note: If the DNA concentrations are low such that you cannot get all 100ng of DNA, buffer and ligase into a 10μL reaction, scale the reaction size as necessary - being sure to increase the amount of buffer proportionally. 1μL of ligase should be sufficient for larger ligation reactions.
Note: Because ligase buffer contains ATP, which degrades upon freeze/thaw cycles, it is a good idea to take a fresh tube, thaw it one time and aliquot individual tubes of 5, 10 or 20μL for storage at -20°C. Whenever you need to set up ligations in the future you can thaw a new tube that you know has only been thawed once before.
Note: Always do controls. See Tips and FAQ below for details.
Note: Try different vector to insert ratios to optimize the ligation reaction. See Tips and FAQ below for details on optimization.
Note: For many ligation reactions, especially if using "high concentration" ligase, 5min at room temperature is enough. For trickier ligations (such as ligation of annealed oligos) the efficiency of ligation can be improved by incubation at 37°C.
Tips and FAQ
Do controls: When doing ligations you should ALWAYS do a vector alone + ligase control. This will allow you to verify that the vector was completely digested and if phosphatase treated, that the phosphatase treatment worked. This control should, in principle, be free of colonies, but the reality is that it will have some amount of background. What you want to see is that your vector + insert ligation has many more colonies than your vector alone ligation.
Additional controls are encouraged, but may only be required for troubleshooting failed ligations. The following table indicates the various controls:
|Uncut vector||-||Checks viability of competent cells and verifies the antibiotic resistance of the plasmid|
|Cut vector||-||Background due to uncut vector|
|Cut vector||+||Background due to vector re-circularization - most useful for phosphatase treated vector|
|Insert or water||+||Any colonies indicate contamination of intact plasmid in ligation or transformation reagents|
Optimizing the Vector:Insert Ratio: Although a 3:1 insert to vector ratio is usually sufficient, you can optimize the amount of insert and vector to improve ligation efficiency in situations where the 3:1 ratio is not working or when doing more complicated cloning. While 3:1 will get you in the ballpark for average size genes and vectors, this ratio is really meant to refer to the molarity of DNA ends available for ligation. Simply put, there are only two ends on any given piece of DNA no matter how long it is, and therefore we need to adjust the amount of DNA used in a ligation based on the length of the DNA to get a proper ratio of 3 available insert ends for every available vector end. Ligation calculators are easily found on the web. Just enter the concentration, lengths of your insert and vector, and what ratio you want, and it will tell you exactly how much of each to use.
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