useing a uniqeu enzyme for both insert and vector digestion - (Oct/05/2011 )
here is the situation:
I am trying to sub clone a fragment from pTZ-57R/T plasmid into another one.
1- the plasmid that contains the insert fragment has been digested with SalI enzyme in order to digest out the insert fragment.
2- after the completion of digestion process, the desired band has been purified from the agarose gel.
3- the ultimate vector has been digested with the same enzyme and after the completion of the digestion process and checking the lineariziation of the vector on agarose gel, it has been treated with alkaline phosphatase and then purified.
4- both vector and insert have been incubated in 37 centigrade degrees for 15 mins. prior to ligation reaction.
5- ligation reactions were set up with different I:V ratios( 2:1, 3:1, 6:1) and incubated in different situations( 16 centigrade degrees or 4 centigrade degrees) for an overnight period of time.
6- the following day, chemically competent bacteria has been transformed with these ligation reactions and after an overnight incubation in 37 centigrade degrees, no colony was seen on the plates.
how can the ligation efficiency be induced in such situations?( when the only option is the use of an unique enzyme for both vector and insert)
all the best
I don't think there is a problem with your restriction enzyme or the digestion, but it would appear there is something inefficient with your ligation reaction. Thank you for providing your vector-to-insert ratios, but could you provide the rest of your ligation conditions? Also, do you cool your vector and insert on ice before adding the rest of the ligation reaction components? Many of the components are heat sensitive and if you add them to a tube that has been pre-heated to 37 degrees, you can lose some activity. Last, what kind of competent cells are you using? Do you know what the approximate tranformation efficiency of these cells are?
Let us know and we'll try to help.
Best of Luck.
1- a usual ligation reaction I set up looks like this :
1- apo.150 ng( I have tried 50 ng as well) vector, 1-3 microlit
2- relative ng of insert to keep it within aforementioned molar ratios with Vector , 5 or more microlit
* both vector anb insert are eluted in distilled water.
3- 10/5 X T4ligase buffer, 2/4 microlit( the stock buffer has been previously dispersed in order to avoid repetitive freez/thaw, DTT smell is dominant and I avoid vortexing the buffer)
4- T4 ligase, 1/5 units/microlit. 1- 2 microlit.
5- d.W up to 20 microlit.
2- yes, both vector and Insert will be cooled either on Ice or left to reach the room temperature before proceeding to the next step.
3- lab-made chemically competent Ecoli, I have used different strains: DH5alpha, JM110, GM2163
thank you in advance
Do you have a transformation control? what about a ligation control? Do this work?
How do you transform your cells?
frankly, I do not check the competemts everytime I make a batch, and I assume because I always use the same method and material, they work well, the only control I use is the amp negative plates on with I test the growth of competent but not transformed bacteria.
the method I use for transformation is sambrook's protocol:
1- mix the ligation reaction with 200 microlit of competent bacteria and incubate on ice for 30 min
2- incubate in 42 centigrade degrees for 90 sec.
3- transfer on ice immediately incubate for 1-2 min
4- add 700 microlit of prewarmed LB, incubate in 37 centigrade degrees for 60 min
5- plate on LBagar Amp posotive, incubate overnight.
6- check for colonies
I dont use a ligation control.
I honestly don't see anything wrong with your protocols, and I'm sure they have worked for you in the past. I would just do a quick tranformation control with any standard vector you have around (pUC, pGEM) just transform a few dilutions of plasmid (0.1 ng, 0.01 ng, 0.001 ng) and plate 10 or 100 uL of each, you should be able to tell really quickly if your cells are in good shape. I always like to do a quick test with new batches of cells, it just makes me feel better that there's one less thing to wonder about.
If that isn't your problem, then I am stumped. I'll keep thinking about it, but definitely let us know if something works.
Best of Luck.
It just won't work!! this was the third time this week I did the colony PCR and none were positive, I was wondering, I use pharmaceutical water for injection in all reactions, could it be the water? Shall use another source, say, millipore?
I would definitely not be using that water. Who knows what they have in it as preservatives, or even ions. It certainly isn't anything like pure water from a Millipore polishing system.
I would suggest strongly that you test your cells for competence. Ligation problems, in my experience, are almost always problems in the DNA being used, or (more commonly) low competence cells. If you have good DNA and good cells, the ligation is rarely a problem. Ligation buffer can, with many freeze/thaw cycles or age cause difficulties.
I have just transformed competent bacteria with a control plasmid to see if they are working properly. any suggestions about the water? can I use millipore water directly out of the purifier? or should I filter it, like with 0.2 micron filter?
There is a huge difference between transforming prepared plasmids and ligation products. Competent cells adequate for subcloning prepared plasmids are often useless for ligation reactions. You need to measure the competence. To do this, you need to do 10x serial dilutions of known concentration plasmid DNA and transform. Efficiencies of at least 10^8 colonies per ug of plasmid DNA are needed for good results in ligations. Typically this is tested by transforming 1 ul of dilutions of DNA at the 10 pg/ul or 100 pg/ul level, and counting colonies (accounting for the amount plated, etc.). For a 10 pg/ul sample, you should get at least 100 colonies if the entire volume is plated.