I am working with a kinase that seems (by fluorescence imaging) to shuttle back and forth from membrane localization to more cytoplasmic localization. We are able to manipulate this localization experimentally.
I have many fusion tagged variant constructs for expression of my kinase in HEK293 cells (Human Embryonic Kidney). By fluorescent microscopy, the full length kinase dead variants become very strongly membrane localized, while kinase active variants become cytosolic.
I have seen membrane fraction Vs cytosolic fraction protocols that look very easy to follow, provided the cells are growing in suspension.
I have been growing my cells in adherent culture on 6well plates. These cells are reasonably well adherent, but not so much that they can not be detached with gentle scraping.
I am hoping to find a simple protocol to use to compare the cytosolic fraction to the membrane fraction in HEK cells expressing kinase dead Vs kinase active forms of my kinase. I predict that Western blotting will show an enrichment in the membrane fraction of kinase dead variants and a reciprocal cytosolic enrichment of kinase active variants.
Please be detailed as far as formulations, volumes, handling, and temperature and all of that kind of thing.
I am looking for any simple and cheap method or methods.
Thanks,
Neal
Mem-Per from Pierce (89842) works well as it separates membrane from cytosolic proteins. You do get about 10% membrane proteins in the cytosolic fraction. Knowing this, you can plan your experiments to further demonstrate localization.
Hi Neal Edward Beeman
Adding to Cornelius Krasel 's answer, before loading the pellet fraction, DO NOT boil the samples in SDS loading dye. Instead, resuspend the pellet in ~0.5-1% SDS, vortex for 1 minute and incubate at room temperature for 15 minutes on a rocker, followed by addition of SDS loading dye and vortexing for 30 seconds. After this, spin down the pellet, to remove all lipid content, as its presence in the gel hinders transfer and blot development.
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