I have had it happen to me three separate times now that when running a 12% SDS-PAGE gel, the dye front stalls/slows two-thirds through the gel and the ladders and my protein eventually catch up to it and starts to run together; I am using a Bio-Rad 250 kD Kaleidoscope standard and after transferring I could clearly see the 25, 20, and 15 kD standards laying on top of one another. After I transferred, I briefly stained with pounceau-S and there was no protein below where dye front and ladders had stalled. My PI had me probe for our protein of interest, which runs at 21 kD, to no avail.
Technical details: For the first two instances, prior to running, I had poured the gels 48 hours prior and wrapped in wet paper towel and left them @ 4 degree as I had done before. The samples themselves I had originally prepped them in RIPA plus inhibitors and then approximately 24 hours prior to loading and running the gels I made each sample to load 50ug protein using RIPA as diluent and a 2X reducing loading dye containing SDS and BME and then just prior to loading I further denatured the samples with a 20 min incubation at 70 degrees C.
The third instance I had this happen I had cast the gels 24 hours prior and prepped the samples for loading just prior to loading as I described before.
The only thing that I have changed recently is that I have started using autoclave tape to tape the outside edges of the plates before pouring to minimize the leakage due to our aging gel casts, which I don't know if that would necessarily have an effect.
Henry,
The tape per se is irrelevant as long as it is removed or razor cut to allow for free current flow. Electrophoresis is a complicated process as you are discovering. Parsimony suggests that your problem lies with your homemade gel casts. My experience points me towards your initiator/catalyst/polymerization reaction as the problem. The variables are vast. Ammonium persulfate degrades in the presence of water. TEMED turns to ammonia plus other inhibitors of catalysis. Urea likewise degrades. Acrylamide degrades to acrylic acid and other inhibitors of polymerization. Bad Tris and bad glycine are candidates to poor polymerization. Oxygen introduced into solutions will inhibit polymmerzation, use a vacuum pump to degas. Detergents remaining on plates are a concern. I recommend using commercially prepared gels. If you must go "old school", read Methods in Enzymology 1978-1983 to learn how to properly prepare homemade gels. Also, Biorad has white papers from that era by excellent chemists. A commercially prepared gel costs about $8 in USA if you form a relationship with a corporate rep. You will have to make many hundreds of homemade gels to recoup the start up costs of doing it yourself. Read the old literature and do a cost benefit analysis. Your PI will agree that purchasing robot made gels is the way to go! I am old school until technology wins and in this case it does!
Michael
Michael,
Thank you very much for your response. You have given me much to consider and to discuss with my PI.
Henry
Henry,
We routinely cast our own gels and I have seen a similar problem in the past couple of years. Running a 15% gel will still resolve the lower mwt protein standards down to 11KDa, but the 10 and even 12 % gels seem to have the standards below 25kDa all running together at the dye front. I get around this by just using a higher % gel when I want to see the small proteins, but if you do find the cause I'd be very interested to know.
Nicola
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