Hi, I recently purified an eukaryotic protein from BL21 cells. Upon examining the fractions, it seems that this protein is found predominantly in the insoluble pool. As such, I need some advice on the protocol to purify the His-tagged fused protein. I believe they are aggregated in the inclusion bodies as I could see black spots in the insoluble pool. According to the EMBL purification guidelines, 1% triton and 1M Urea are added to the pellet for washing. However, can someone help clarify if these reagents solubilise the rest of the cell components and not the inclusion bodies? In addition, will this treatment cause the release of proteins from inclusion bodies?
The EMBL protocol I saw can be obtained from this link: http://www.embl.de/pepcore/pepcore_services/protein_purification/extraction_clarification/solubilisation_renaturation/
Also, please do share other protocols for protein purification from inclusion bodies.
Thanks in advance!
Hi Adam,
since my proteins of interest are expressed in inclusion bodies (using BL21 DE3) I use the following protocol (which works really fine) adapted from Pratt et al. (2006) Nature Protocols Vol1 No.2 (works for membrane proteins, soluble proteins, standard peptides, etc...)
Please find attached my lab protocol as well as the original publication.
Good luck,
Heidi
Article Multiplexed absolute quantification for proteomics using con...
The triton and urea solubilize almost all cell components. The triton will break the cell membrane and the urea will unfold all proteins present.
In cases such as yours, sonication is performed on bacterial cells expressing the protein in a buffer containing 1.5% triton and optionally lysozyme, usually in a phosphate buffer. This will usually clean away all impurities and leave the inclusion body after a few rounds of sonication and centrifugation. Usually, once the inclusion bodies are obtained in isolated form, they are solubulized with guanidine HCl or some other denaturant (maybe urea).
From here, it depends on your protein. If you are using a His tag then you use nickel beads for purification. After this you will have to refold the protein. It may be advisable to perform some sort of liquid chromatogrpahy to further purify the protein after or before refolding, depending on your protein.
Hi Adam,
since my proteins of interest are expressed in inclusion bodies (using BL21 DE3) I use the following protocol (which works really fine) adapted from Pratt et al. (2006) Nature Protocols Vol1 No.2 (works for membrane proteins, soluble proteins, standard peptides, etc...)
Please find attached my lab protocol as well as the original publication.
Good luck,
Heidi
Article Multiplexed absolute quantification for proteomics using con...
Hi Adam,
sorry, didn´t attache the recipes of the used buffers - here they come:
Used buffers:
Binding Buffer (=wash buffer): Elution Buffer:
20 mM phosphate buffer pH 7.4 20 mM phosphate buffer pH 7.4
0.5 m NaCl 0.5 m NaCl
20 mM imidazole 500 mM imidazole
6 M guanidine hydrochloride 6 M guanidine hydrochloride
Cheers, Heidi
Thank you all for your reply!
Heidi & Joanne: Do you all perform re-folding upon purification with nickel beads? It seems that there is no such step in your descriptions. Or perhaps I am missing something out?
In addition, I observed in some protocols that dialysis is used to gradually reduce the amount of urea or GuHcl from the system. I wonder if this decrement is crucial in the renaturation of proteins?
Hi Adam,
after IMAC (Ni-beads) you HAVE to dialyse your elution fractions to get rid of high salt and denaturant concentrations. I usually to this via dialysis in 50 mM ABC, 1 mM DTT for 3 x 2hours at 4 °C (using commercially available dialysis cassettes). This step also causes re-folding of your protein what is very important if you want to study some enzyme kinetics, etc......where you protein has to be refolded in it`s natural structure.
Hi,
maybe I can help to clarify the question regarding the use of urea in the washing of the inclusion bodies. As stated above, the detergent should help solubilising membrane fragments that are attached to the IBs and the urea should wash away other components that are bound unspecifically to the IBs (as it says in the protocol, e.g. proteases). The conc. of urea is usually not high enough to solubilise IBs so it helps as an initial purification step of the IBs. This washing step sometimes gives you really pure (>80%) protein which can then be solubilised in buffer with 6M guanidinium or 6-8M urea and further purified e.g. by IMAC or ion-exchange if you are using urea (IEX doesn't work with 6M Gua because it's charged!).
Please note that in some cases the protein can be present in a fully or partially folded form inside the insoluble material, i.e. it is "just" aggregated (or trapped/attached to other component in the pellet like DNA or membranes) and not fully denatured. That kind of material can sometimes be solubilized to a large extent already with 1%TritonX-100 and/or 1M urea, so keep the supernatant of the first washing step and check by SDS-PAGE (you might lose most of your protein, if you just discard this wash fraction!!).
Last not least, one short comment to a method that some people use, where 6M guanidinium is used directly to solubilise the cell pellet. The problem here is that you carry over all the proteins and other stuff (DNA, RNA, membranes) that can cause trouble during the purification step. You would lose all the advantages that separating IBs under non-denaturing conditions offer (DNase digestion works, almost pure target protein in IB, thus low background).
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