since I want to do a northern blot, I have to perform a denaturing agaose
gel electrophorese with total RNA of Paramecium.
My problem is, that I don't even get the typical bands (28S and 18S rRNA)
for total eukaryotic RNA. I isolated the RNA with Trizol and prepared a 1,1%
agarose gel with Formaldehyd (3-4ml 37% per 40ml gel solution) in a northern
running buffer (20mM MOPS, 5 mM NaAc, 1mM EDTA). The run was at 50-70V.
I denatured the RNA samples at 65°C for 5-10 min.
I used a ssRNA ladder in its original purchased loading buffer (7M urea,
ficoll,...), which showed a perfect run in this gel. In contrast, my RNA
samples in this loading buffer showed a pattern of different bands with one
very dominant (at ca 3500 nt).
When I used another common loading buffer receipe(50% formamide, 25%
formaldehyde,...), I got a smear and some weak bands. I can nearly exclude
that the RNA that I used was not severely degraded, since I had it on a
bioanalyser a few month ago (stored at -70°C).
What might be wrong in my system?
Thank you very much!