Why would b-actin run differently using the same lysate but on a different gel?
I solubilized rat brain tissue using the Ambion PARIS kit according to their protocol. I ran 20ug of protein on a 10% gel, transferred to pvdf, etc. I first probed for phospho-jnk, then stripped the membrane using Abcam's recipe (SDS, Tris, BME), reprobed for total jnk, stripped again, and reprobed for b-actin (Thermofisher). No issues whatsoever. Great b-actin band in the right place, about 42kDa.
I then took the same lysates, load 20ug of protein onto each of 2 gels (12% this time), transferred to pvdf, etc., and then probed 1 for p-jnk and 1 for p-erk. I stripped them both, probed for total jnk and total erk, stripped again and probed both for b-actin (same dilution). This time b-actin shows up faintly around 42kDa but also has a much bolder band around 50-52kDa on both membranes. Can anyone tell me why this would happen? The only change was from a 10% to a 12% gel, as far as I know. I considered that perhaps there was ineffective stripping but that 50kDa band was not present previously on the ERK blots. Could something have gone wrong in the preparation for loading the samples? Can lysate taken from tissue rather than cells be unpredictable? I have used this same b-actin antibody before quite successfully with endothelial cells. Please advise. Thank you!
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