I am working with an RNA primer extension assay to elucidate the activity of a polymerase with Urea-PAGE gels.
Unfortunately, there are not many ladders for low MW ssRNA gels so am using a low MW ssDNA ladder from IDT. I know it won't correlate perfectly but was hoping it would be close enough.
I am getting polymerization but am struggling to rationalize why i am getting bands so much higher than the full extension band would be. I was also hoping for any assistance in reducing smearing (15 well gel at 1mm thickness using 20% acrylamide in 1x TBE; I am also already pre-running/rinsing the wells and loading 9μL).
I have also seen a mix of papers using formamide based or urea based loading buffers to quench the reaction -- currently i am using urea but is there a benefit of one over the other?
Any help advice would be appreciated. (I have attached a gel for reference) As a note, I am focusing on the 12.5nM RNA lane -- this was just a titration I had done previously of RNA but illustrated my questions the best.
small RNA of the same size can have different migration depending on their nucleotide sequences (the presence/absence of a phospate group at one end can affect migration too) so I would try to have the expected RNA as control (maybe from an in vitro transcription (T7) assay? or ordered from a company (chemical synthesis)?) or at least an RNA of the right size) Other bands could be due to stopped extensions or to incomplete denaturation of the RNA and the DNA-RNA hybrid. your samples should be denatured before running (50% desionized formamide or 5M urea). it could also be good to run your gel hot (prerun to get it warm, rince the wells to remove urea then load your samples. you can make a RNA ladder by labelling the 5' end of an RNA then treated it with low concentration of NaOH for a short time so you will get random cleavages on the molecule; with the right time and concentration you will get a RNA ladder with 1 base intervals..
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