My lab has purchased some of the Super Sep precast Phos-Tag gels from Wako (#198-17981) to try and determine the phosphorylation state of a kinase we're interested in. We have had very little luck getting these gels to work (high numbers of nonspecific bands, distorted bands, no separation of phosphorylated protein from normal form). As a control, we know that phosphorylation of a different kinase increases under the conditions we are testing (we can confirm with a phospho-specific antibody), but we have been completely unable to visualize a specfic band shift on the Phos-Tag gel for our control kinase.

I have been following the EDTA chelation protocol after running the gel before transferring (20 min 3X in 10 mM EDTA) and making sure there has been enough space between samples and the ladder. Our lysis buffer for isolating the whole cell lysates uses PBS w 1% Tx-100, could the PBS be interfering with the gel? Are there any specific conditions for running the gel that need to be followed to get good separation of phospho vs. non-phospho? Should I use our phosphospecific antibody as a control to see if it picks out specific bands on the phos-tag gel or will the reagent interfere with antibody binding?

We would really like to get this system up and running in our lab but we have yet to get a run to show a result with our positive control protein. Any advice would be greatly appreciated!