I got a really messy SDS-PAGE gel colored after transfer with Coomassie blue. I'm wondering also why the sample stopped running although the blue line with the samples was much lower than the lowest protein line in the picture (picture taken with the UVP camera where you can't see it in color).
The samples in the picture are freshly prepared rat brain samples.
As the lysis buffer I use RIPA buffer
(50 mM Tris-HCl, pH 8.0, with 150 mM sodium chloride, 1.0% Igepal CA-630 (NP-40), 0.5% sodium deoxycholate, and 0.1% sodium dodecyl sulfate) with protease inhibitor cocktail (PIC) on ice.
After homogenising the dissected brain structure with the RIPA lysis buffer +PIC, I checked the protein concentration with the BCA method and after calculations I made sure I have 20 ug of total protein per well.
But even my beta-actin didn't give any signal at all (both primary and secondary antibody has worked fine in the past) this time.
The gel contained a 10% separating (1.5M Tris, pH 8.8) and 4% stacking gel (0.5 M Tris, pH 6.8.
As the running buffer I used the classical recipe:
250 mM Tris
1.92 M glycine
1% SDS
The electrophoresis took about 2.5 h (kinda slow!?). I set the electricity for 1 gel 20 mA.
My transfer buffer is the Bjerrum and Schaffer-Nielsen Buffer:
48mM Tris pH9.2
39 mM glycine
20% Methanol
which is apparently better for semi-dry transfer (1 h, 1mA per 1 cm2, so around 40 mA for one gel to membrane transfer).
Any suggestions in which direction I should look into?