I'm expressing recombinant protein in tobacco and trying to get some crude extracts for SDS-PAGE/Western blotting. I began by using a relatively gentle protein extraction buffer (50mM HEPES, 1mM EDTA, 10mM KAc, 5mM MgAc, 1mM DTT, 2mM PMSF) but my protein of interest was not extracted (checked by western)
After realising that my recombinant protein might be membrane-associated, I used a more stringent extraction process. I ground up my leaf material first in 62.5mM Tris-HCl, 10% glycerol, 5% BME, 2% triton, and removed the supernatant, then ground up the pellet in the same buffer, but with 1% sds instead of triton. When doing Western analysis I found that my protein of interest was released in either one or the other of these buffers (I have a few different constructs on the go).
However, I want to do a bradford on these samples and some of the reagents aren't compatible in their current concentrations, and in order to dilute them down enough they would be too low for the assay to pick up.
My next step was to acetone-precipitate my samples and resuspend them in a more suitable buffer. However, I found it really hard to get my pellets back into solution, and when I ran what I managed to resuspend on a gel, it was just a big smear, while before precipitation I had lovely discrete bands.
I've also tried methanol/chloroform and TCA precipitation on these samples but each time all I get is a smear.
Could there be something in my original buffers that is stopping me getting my samples back nicely or am I making some other mistake?
Thanks for any insight you can give me.
Edited by vojera, 26 September 2011 - 02:31 AM.