I dug around on the net and found this method to remove lipids from
proteins:
More precisely, from denatured proteins. That's what methanol/chloroform
phase does for most proteins.
"Wessel & Fluegge (1984), Anal. Biochem. 138:141-143. Itґs a methanol/
chloroform precipitation and gives you a pellet that is easily
redissolved. The method was especially devised for removing lipids or
detergents, so it should be perfect for you."
-- http://www.bio.net/bionet/mm/methods/1996-December/052513.html
By far the best method of concentration/desalting/de-lipidizing proteins
for SDS gels. I've used it extensively over the years. Even then, the
efficiency of precipitation drops off very significantly for most small
proteins at low [protein].
Is this still the preferred way? I do not want to use reagents that
are *themselves* likely to denature my protein. Has anyone tried
cyclodextrins?
Lots of people did. They work. So if you have protein that you can easily
immobilize, washing the matrix extensively with b-cyclodextrin will do the
trick. But immobilized cyclodextrins are not readily available for
reasonable price. So for untagged protein your next bet would be various
detergent removal sorbents available from Calbiochem, Pierce, Bio-Rad and
likely many others. All of these WILL bind your protein to various extent,
but usually not completely because they are also work as size exclusion.
I'm specifically trying to strip sarcosyl. I want to do
it completely.
What's the definition of completely? If you are lucky and your protein
binds to cation exchangers, simply washing the column with >20 CV of low
salt buffer (even better with non-denaturing concentrations of alcohols or
glycols) usually will decrease sarcosyl concentration by ~ 100X. Pretty
much the same if your his-tagged protein is bound to IMAC sorbent.
- Dima
