pcr product digestion problemdigestion problem
Posted 15 November 2012 - 02:29 AM
I have an insert is 200 bp.it is a pcr product. I did pcr purification and it is 138 ng/ul and 158 ng/ul. my vector is 6000 bp and 580 ng/ ul. I want to clonning but I am not . My restricition digestion enzymes NotI and PacI (FERMANTAS 10u/ul). I can not set of the amount of enzyme and vector and ınsert. how do I adjust the amount of reaction? fermantas is recommend
10X Buffer G
4-fold excess of NotI
4-fold excess of PacI
Incubate at 37 but ı can't understand.please help me.
Posted 15 November 2012 - 12:04 PM
Use n (amount you want) =concentration x volume.
Check that Not1 and Pac1 are compatible for double digest and using appropriate buffer.
Posted 15 November 2012 - 06:00 PM
From their PCR product digestion protocol PDFs, they recommend 10-20 units of NotI to digest 0.1-0.5 ug of PCR product. So you would need to add 4x that much (40-80 units) for 0.1-0.5 ug of PCR product in Buffer G. They recommend the same amounts for PacI, so you'd need to add 40-80 units of both enzymes for 0.1-0.5 ug of DNA.
For your vector, from the main enzyme datasheets, it looks like they recommend 10-20 units per 0.5-1 ug of DNA. So you would need to use 40-80 units per 0.5-1 ug of vector DNA for that reaction in Buffer G.
Because of the poor activity of both of the enzymes in that buffer, I wouldn't try to do a double-digest with the enzymes you have. If you can't change the restriction sites you're using, an alternative is using their FastDigest enzymes, which are all supposed to work in the same buffer (according to their site). It looks like they have FastDigest versions of both NotI and PacI.
Otherwise, I'd try to change the enzymes to ones that can perform a double-digest in the same buffer. Trying to cut in a buffer that neither enzyme likes is asking for trouble, in my opinion.
Posted 07 December 2012 - 04:45 AM
Posted 07 December 2012 - 04:40 PM
For a ligation, I would mix roughly 100 ng total DNA of vector+insert in 10 µl total volume for the reaction. For a first try, most people seem to suggest a ratio of 3:1 molar excess of insert over vector. If that doesn't work, but you're sure your enzymes are cutting, you can try changing the ratio lower or higher (from 1:1 to maybe 10:1 insert:vector).
For that calculation: ng insert = 3*(ng vector)(bp insert)/(bp vector)
In your cloning, with a 200 bp insert, 6000 bp vector, I would use: ng insert = 3*(90 ng vector)(200 bp)/(6000 bp) = 9 ng insert with 90 ng vector. (I roughly estimate the amount of vector to use by taking the length of the vector (6000) and add 3x the length of the insert (600) (total 6600), then figure out what % of the total the vector is (6000/6600 is ~90%)) There may be an easier way to calculate it, but this is how I do it.
I have also been told that if you run your DNAs on a gel, if you can see the band at a given volume loaded, then you can use that amount in a ligation (whether it is the insert or the vector). So if you loaded 2 µl of your insert and vector DNAs, and can faintly see bands for both of them, then that's a good amount of DNA of each to try in a ligation. I've never done my ligations that way, but if you have no way to determine the concentration of DNA in your samples, then that might be worth trying.
Edited by John Forsberg, 07 December 2012 - 04:42 PM.