Hi!
Can anyone give insights as to which of these two methods is better?
We're trying to express a recombinant protein in E.coli. It is quite difficult to express this protein in its native soluble form as its monomer has 4 intrachain disulfide bonds and one Cys residue to form a homodimer. Other signal peptides such as OmpA has been tried, but it still ended up in insoluble inclusion bodies.
I've read about the introduction of SOX+disulfide isomerase, and I'm thinking maybe it's worth a try. But I would like to hear from people who have firsthand experience using this method on multiple Cys-containing proteins.
Thanks in advance^^
To form the disulfide bonds, you have to either use a strain engineered to have less-reducing cytoplasmic environment (Origami strains and others http://wolfson.huji.ac.il/expression/bac-strains-prot-exp.html), or export your protein to the periplasm. A third alternative is the production of inclusion bodies and refolding in vitro in the presence of a redox shuttle. It is more tedious than in vivo folding, but can work where the in-vivo folding methods fail. In the worst case, you may have to go to an eukaryotic host, e.g. yeast.
None of these methods is a priori better than the others, and for a new protein each may need quite a bit of optimization. MBP fusions can keep your protein in solution, giving it more time to fold properly, but may also just lead to the formation of soluble aggregates instead of insoluble inclusion bodies. Unfortunately, there in no one procedure that works best for all disulfide-containing proteins.
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