I recently ran a 2D gel electrophoresis, however my spots were a wee bit faint, so i increased the exposure of the image. This resulted in the appearance of small pixels (dirty spots?) across my gel that could interfere with the identification process later on.
The only variable that I can think of in my protocol is that i used the oriole stain that has been used once before. This may have resulted in poor staining, but does it explain the pixelation when the exposure is increased?
My samples are very valuable and I can't afford a rerun. I have attached the image of my gel. Any suggestions?
This happens with Flamingo as well and it seems to be directly related to the amount of protein in the gel. The stains are colloidal and if they don't bind to protein, the molecules bind to themselves and precipitate. I've never explored how to fix this because it only happens with low loads of protein, but I would try halving the concentration of the stain and REALLY shaking the stain in a 50ml tube before pouring it on.
That could work, but I don't think the gel would survive the shaking and would tear into pieces. I haven't tried repeated washing either. I recall a throwaway line in a paper once by Malcolm Ball that Flamingo binds to Protein-SDS complexes but have never seen it mentioned anywhere else. You could try washing the gel in fix solution (40% MeOH, 10% acetic acid) or 40% MeOH and see if that removes excess SDS and colloidal stain. I have no idea if that would work. If you could remove the precipitated stain, the stain bound to the protein would still be there so you wouldn't need to re-stain.
I think the issue you need to address is the dynamic range of the sample. There are a number of high abundance proteins there and, unless they are the ones you want to quantitate the differences in, you won't see or be able to quantitate the lower abundance spots.
Lastly, I don't think Oriole is MS compatible, so if you need to identify by MS, you will have to switch to Flamingo or Sypro Ruby.
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