I am trying to express my protein of interest by IPTG induction. I can not see any difference between uninduced and induced sample in SDS-PAGE after induction for 4 hours in 28 degree. Final concentration of IPTG is 1mM. I used BL21(DE3) strain and PET30a as expression vector
There can be multiple problems/factors. Assuming your vector is ok (no mutations in the important regions) the following immediately came to my mind:
1. Toxic protein; can be checked by measuring OD (no or low growth after induction indicates toxicity)
2. slow induction; prolong induction period to 16h (or more)
3. low expression level; check availability by activity test (for enzymes) or western blot (e.g. anti-His antibodys)
5. wrong induction temperature, some proteins are only expressed at low temperatures; check expression at 16°C
4. damaged strain; do a positive control with a working plasmid and the “same” strain
Thomas Krause Thank you. Later I already tried with low temperature and prolonging time period but did not work out. I will work on with other possibilities you suggested.
Hi there,
Induction might not be strong enough to be noticable on SDS/PAGE. In order to be sure you should either perform a WB using a specific antibody (anti protein or anto tag) or perform a micropurification and see what you get. Without these experiments you can't conclude there is no expression...
Dominique Liger So I should use the induced sample to do western blot? And can you please provide any reference or protocol for micro purification? Thank you for your suggession.
Hi again,
For His tagged protein, you can use either the commercial NiNTA spin system from Qiagen which is a bit of expensive (https://www.qiagen.com/us/products/discovery-and-translational-research/protein-purification/tagged-protein-expression-purification-detection/ni-nta-spin-system/) or adapt the protocol using NiNTA resin as a batch and centrifuge.
Hello,
My proteins of interest tend to mis-fold if induced too strongly and end up socked away in inclusion bodies, never to return. Depending on your protein, this might be the case for you as well. To get around this, we do a very slow induction, which seems to allow more of the expressed protein to fold correctly. We use pET vectors in E. coli, modified Terrific Broth as media, and induce when OD reaches ~0.8 with 250uM IPTG (though some look like I should try an even lower concentration), then shake at 18C fort 20-24 hours. I know another group working with the same family of enzymes doesn't add IPTG at all, just uses a leaky promoter, and swears by that (I tried it once, but it actually gave worse results for me. But again, only tried once, so the prep just might've been off for any number of other reasons). More recently, I've been playing with culture aeration to try to achieve better bacterial growth, and hopefully, more expressed protein.
Also, how are you lysing your cells? My enzymes tend to be somewhat short-lived at the best of times, and seem to be intolerant of certain lysis methods. For example, chemical lysis and/or sonication kill the enzymes on the spot, as does very high salt, or anything that heats the sample or causes it to froth. We get the best results using an old French Press (which I hate, because it is a pain in the you-know-where), and more recently, a cell disruptor another lab near us purchased, which I think uses the same principles at the French Press. Lysozyme doesn't seem to hurt our enzymes, but doesn't seem to improve our results much either. Anyway, that's another thing you could maybe play around with. Good luck!!
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