Hi lab people!
I am having a problem with my agarose denaturing RNA gels. I started using the standard protocol with 2,2M of formaldehyde in the gel, but got suspicious it was too concentrated and tried to go as low as 1M without any improvements. Its suppose to be used for northern blotting so I would like to have sharper bands. It is particularly funny since in my previous lab it use to work just fine. Following is the protocol summary and gel picture (loaded 5ug of total RNA).
Anyone has any suggestions or solved a similar problem before? Thanks a lot in advance!
Gel: 1g agarose + 80 mL water. Heat-dissolve, adjust volume to compensate evaporated water, cool to around 60°C, add 10mL 10X MOPS and 10mL of formaldehyde 37% (12,3M).
Denaturing loading buffer (1mL always fresh): 500uL formamide, 170uL formaldehyde (same as above), 130ul 10X MOPS, 200uL LB RNA.
LB RNA: 30% Ficoll, 1mM EDTA, 100ug/mL Ethidium Bromide, Bromophenol Blue.
For loading the gel I use 1 volume of RNA mixed with 3 volumes of denaturing loading buffer, heat at 65°C for 5 minutes, snap cool in water-ice and load on 10 min. pre-runned gel.
Thanks again for any suggestion!!!