180 kDa protein is getting stuck at the stacking gel and separating gel interface.
Stacking gel-3.5%
Separating gel-12.5% (this percentage is chosen because we run a 18kDa protein along with the 180kDa protein. With a 10% gel, the 18kDa protein runs with the gel front and therefore detection gets problematic.)
The standard runs perfectly and so do the lower molecular weight proteins. On staining the PVDF membrane for the 180kDa protein we see massive bands at the interface.
Samples are treated with 3X lamelli buffer and boiled for 5 mins prior to loading. We have also rechecked the pH of the buffers and they seemed fine.
Any inputs will be appreciated!
It likely forms higher order complexes/aggregates with other proteins and stuck at the interface. Try loading the samples sooner. I mean after boiling, cool it in ice for 2-3 mins, pipette it and load immediately. Basically you're reducing the processing time which will in turn reduce formation of complexes/aggregates again after boiling.
Hi Megha,
I would suggest that you ether use a gradient gel 4-14%, or simply use one gel with 6-7% for your high molecular weight protein and a second gel with 12.5% for the 10kDa. That would probably be the easiest and fastest way to get rid of the problem.
I also assume that you encounter problems in western blot if you are going for both protein from the same gel, since the blotting conditions differ for proteins with these big diffences in molecular weight.
Best,
Kai
That's one of the telltale signs of protein aggregation. Try including urea in the Laemmli buffer, and derivatizing free sulfhydryls with iodoacetamide or any other thiol-specific reagent to block the reformation of disulfide bonds.
are u adding any reducing reagent in Laemmli buffer. Add DTT or BME freeshly in Laemmli buffer, mix content properly (may be vortexed), and keep sample at RT for 5 min, then boil sample for 5min. Cool down sample at RT and load on 10% gel. In take out the gel before dye front comes out of gel. staking of 5% is good enough.
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