In the denaturing RNA gel electrophoresis, by the action of used formamide or formaldehyde the secondary structures of RNA (hair pin loop, pseudo double helix etc.) are removed and further restricted to form again as long as the denaturing agents like formaldehyde is present . So, as there is no more secondary structure then how can EtBr still able to bind it? Can somebody please let me know the mechanism behind it?
EtBr also stains single-strand nucleic acid (RNA and DNA) with about 10-20x lower efficiency to that of double-strand nucleic acid.
once the gel is done running:
1) fix in 10% acetic acid for 15 min (when the bromophenol blue turns yellow it's done)
2) stain for 15-30 min in 0.5-1x TBE + 1ug/ml EtBr.
3) destain in 0.5-1x TBE for ~10 min
easy as 1-2-3...and you should be good to go.
the detection limit is ~25ng/band (obviously this will vary depending on the length of the particular RNA you're trying to detect).
also - if you're using a fancy image analyzer to take a picture of the gel you can jack up the signal intensity (if the signal-to-noise is too high - destain for longer). or if you can, increase your signal by loading more RNA/well.
Denatured RNA still has double-stranded regions due to base-pairing interaction, which allows EtBr to intercalate.
In addition to the previous two answers, I would like to add that although RNA (or even ssDNA) can loop back into itself to form secondary structures, even after treatment with denaturing agents, such an action occurs at very low rates. ssDNA or RNA treated with denaturing agents is ~10 times less likely to form secondary structures, which is why EtBr-based detection of nucleic acids is a lot less sensitive and protocols such as this one recommend "... 10 times more nucleic acid for equivalent detection." Hope this helps clarify things a bit more.
Are you using glyoxal or formaldehyde for RNA denaturation? I noticed that formaldehyde quenches the ETBr signal. If you store the RNA gel at 4C overnight, the signal is much brighter. As an alternative you can stain the RNA gel with methylen blue e.g. 0.02% Methylen blue in 10 mM Tris-HCl, pH 7.5 (may be reused several times). Destain the gel with deionized water until the rRNA bands are visible.
I routinely stain RNA with EtBr on urea denaturing gels. Unless you have a very short oligo, it is far more sensitive than UV shadowing (which we use for preparative purposes). I have always assumed that the RNA retains some base pairing and base stacking of single stranded regions that allow ethidium to fluoresce.
Denatured RNA forms double stranded zones also in the form of what is called the Hairpin formation, EtBr intercalates there and can be detected by Gel electrophoresis . Moreover, heat denatured RNA can refold to Hairpin formation easily than chemical denatured RNA. In so far as the qualitative detection of RNA is concerned the degree of renaturation will not make much difference .
There are some molecules with very low secondary structure; for instance homopolymers of A's or U´s are not stained by EtBr after been fractionated by electrophoresis in native gels.
@Arvis Sulovari : Thank you for the answer.
@Stefan Neuenschwander : I am using a formaldehyde denaturing gel. Thanks a lot for the idea of using methylene blue staining. I would surely try it next time.
EtBr intercalates between the nitrogenous bases and forms stacking interactions with them.
I denaturate RNA by dimethyl sulfoxide and glyoxal and run the electrophoresis in phosphate buffer system. Then I remove the glyoxal by 50 mM NaOH and at the same time stain the gel by EtBr.
I am also using a formaldehyde denaturing gel like you, but I add EtBr into the sample RNA. RNA loading buffer: 50% glycerol, 1mM EDTA, 0,4% bromophenol blue and 1mg/ml ethidium bromide. Add 2 mkl buffer per 10-20 mkl RNA sample (RNA plus sample buffer) after denaturing RNA at 70g C . Gel may be photographed immediately after the runnig.
EtBr also stains single-strand nucleic acid (RNA and DNA) with about 10-20x lower efficiency to that of double-strand nucleic acid.
once the gel is done running:
1) fix in 10% acetic acid for 15 min (when the bromophenol blue turns yellow it's done)
2) stain for 15-30 min in 0.5-1x TBE + 1ug/ml EtBr.
3) destain in 0.5-1x TBE for ~10 min
easy as 1-2-3...and you should be good to go.
the detection limit is ~25ng/band (obviously this will vary depending on the length of the particular RNA you're trying to detect).
also - if you're using a fancy image analyzer to take a picture of the gel you can jack up the signal intensity (if the signal-to-noise is too high - destain for longer). or if you can, increase your signal by loading more RNA/well.
I am trying to denature double stranded RNA, in 1*TE buffer, using 50 mM NaOH and putting it at 37C for 1 hour. When I run 1% standard agarose gel with EtBr (running buffer: 0.5% TAE with EtBr), I don't see single stranded RNA band in the gel image even after staining the gel in EtBr. Can somebody please inform me where might be the mistake?
A standard protocol for converting RNA to nucleotides is 300 mM KOH for 30 minutes at 37 degrees C. You may be converting the RNA to short chains of nucleotides using NaOH and they will run ahead of the BromoPhenolBlue tracking dye
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