I've been recently given the tast of identifying the affinity of my protein towards a synthetic oligo 19bp long. Previous work on the protein suggests an EMSA on a 2% agarose gel, run at 70V for 40min. Ethidium bromide was added to the agarose gel before cooling to resolve the bands quicker.
I initially observed bands in the protein+DNA lane, however as we increased the concentration of the protein, the DNA band below didn't seem to get any feint.
So I ran a lane with just protein (~100uM), and got this consistent band, heavier than the DNA control but consistent with the DNA+Protein lanes. We thought it might be a DNA contaminant so we ran it through a heparin column, but the band was still present.
We tried running mutants of the protein with which the crystal structure has been resolved with no DNA contaminants, and the same band was observed, but with varying thicknesses. (See image)
Denaturing the protein with Guanidinium-HCl, which I thought would liberate the 'mystery band' only caused a pretty much insoluble complex which wouldn't migrate through the gel.
I honestly don't know what's causing the band: since the gel already has EtBr I would assume that anything that fluoresces is DNA.
Where is that band shift? D: