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Plasmid Ligation does not work-suggestions? - (Aug/03/2007 )

Hi everyone,
I am new to molecular biology research and am attempting to construct a plasmid. I am having a problem which I'll describe here. Any advice would be welcome!

Here is a description of my problem:
I am trying to remove a gene from a plasmid created previously by someone else in my lab. Then, I would like to replace that gene with another, different gene (a thioesterase). The original plasmid contains ClaI and EcoRI sites in the proper places, which I have digested. Gel analysis shows two strands of the proper length. I also have a PCR product of the thioesterase which I would like to insert, with the EcoRI and ClaI sites included in the primers. I believe my digestion reactions are working properly.

My ligations do not seem to work. After isolating both the plasmid backbone and the thioesterase from a gel, I have attempted multiple ligation reactions using anywhere from 1:1 to 1:6 vector:insert ratio. I use T4 DNA ligase from invitrogen.

When I analyze these reacions on a gel compared to a negative control (the backbone and insert alone), there is one difference: the insert seems to be ligating with itself. My insert is about 900 bp, and I see bands at 900 bp, fainter at 1800 bp, and still fainter at 2700 bp. I also see a band where the plasmid backbone is, but I do not see a band where the completed plasmid should be.

I have also attempted a positive control to see if my ligase is working. I digested the original plasmid, ran it on a gel, isolated the backbone and the original gene, and tried to religate them back together. The positive control did NOT work. Instead, I had the same problem: the original gene ligated with itself to form a double-length strand, but did not ligate with the plasmid backbone.

My first guess was that ligase wasn't working, but the insert ligating with itself seems to contradict that. Does anyone have any idea what is going on, or how to track down the source of the problem? I am pretty lost and don't know how to proceed. I would be very grateful for any suggestions!


Describe how you are doing the ligations. The EcoRI site has an AATT overhang, which has a low Tm. This requires a relatively low annealing temperature to efficiently ligate. Try an overnight ligation at 14C, or (easier) at 4C. Other common problems are too much DNA, too much ligase, UV damage to DNA, impurities in the DNA, endonuclease activity, bad ligase,bad ligase buffer. It's hard to help more without knowing exactly what you are trying. Your experimental approach of trying to ligate the pieces independently and running the result on a gel is good. We find that heat killing the ligase before gels makes a difference. You can enhance the test by adding small amounts of one or the other of the enzymes to the ligated results prior to running them on the gel. The enzyme recuts the corresponding sites, leaving (in principle) only double length fragments. The ratio of single to double length fragments is an assay for the ligation efficiency of the other enzyme cut ends. You may need to adjust the salt concentration of the T4 ligation buffer to allow the enzymes to cut well. We adjust it to 25 mM, which is lower than optimal for the enzyme, but not so high as to inhibit the T4 ligase.


What kind of difference does heat killing the ligase make, good or bad? I have been doing that at 65 deg C before running gels. Also, since you mentioned too much DNA, I was wondering what are considered good upper limits for amount of DNA of both insert and backbone?

I've been running ligation reactions at RT (usually about 26 deg C) so I'll have to try putting them in the refrigerator overnight. Here is an example of the kind of recipe I typically use, although it varies depending on DNA concentrations I have available.

1 uL ligase
2 uL buffer
4 uL insert ( at 25 ug/mL, 900 bp)
3 uL vector backbone ( at 88 ug/mL, 6600 bp))

I have run them anywhere from 1 hour to 10 hours with no positive results... any disadvantage to leaving them for too long, or should I always be running them overnight?

Since it's hard to see these on gels, I ran 10-15 of them then reconcentrated them using phenol-chloroform-isoamyl alcohol extraction and ethanol precipitation.



For ligations, I would use 20ng-30ngs of vector DNA and its corresponding 1:3 ratio insert. Try to calculate your DNA accrodingly. Atltimes, I would use a total of 200ngs, which is not normal. In your case, the total DNA is too high which could be the problem.


Heat killing the ligase makes the gels much easier to read. Final concentrations of DNA in ligation reactions (favoring recircularization) should be 1-10 ng/ul. You need much less ligase than you are using. You are adding about 400 units, and need about 10 for sticky ended ligations. The extra DNA can cause two problems -- concatamers, but more importantly, if it has any inhibitors such as EtOH or phenol, you are not diluting them. Try a reaction with 0.3 ul of each of your DNA samples, 0.2 ul ligase, 2 ul of T4 ligation buffer, and the rest of the 20 ul DI water. You can transform 1 ul of this directly into 50 ul of competent cells. You may be able to do this with a RT ligation for 10 minutes, but I'd also set up an overnight one at lower temperature.


How wil you calculate the 3:1 molar ratio of insert:vec? I've read tat its concerned with pmol concentrations of DNA termini. Can you explain me how to calculate tat stating an example?


QUOTE (buddie @ Aug 6 2007, 07:24 PM)
How wil you calculate the 3:1 molar ratio of insert:vec? I've read tat its concerned with pmol concentrations of DNA termini. Can you explain me how to calculate tat stating an example?

you can calculate it for example here;sp=Sligations

-Ned Land-