The goal of the protein purification process is to obtain highly pure, stable and active protein for downstream experiments. The exact nature of the downstream applications will determine the purity level you need to obtain, the compatible buffer/storage conditions and the necessary quality control tests. For example, protein that will be used for in vitro biochemical or structural biology experiments will need to fulfil different conditions than proteins that will be used for in vivo immunization or other immunological experiments. Therefore, it’s important to have an idea of the requirements of your planned downstream applications before starting a protein expression and purification experiment.
Generally, the protein purification process consists of the following steps:
If you’re not sure what would be the best design for your protein expression and purification experiment, don’t hesitate to contact the PEPCF staff for some help or advice.
EMBL PEPCF offers the following services regarding protein purification:
The way you will further process your cells after the protein expression, depends on the type of proteins you are working with. Usually, the first step is centrifugation to separate the cell culture medium from the cell pellet. When your protein is secreted into the cell culture medium, you’ll continue with the supernatant after centrifugation. For intracellular proteins, you’ll proceed with the cell pellet. For cytosolic proteins, you’ll lyse the cells using your method of preference. After cell lysis, an extra centrifugation step is required to separate the cell debris from the soluble protein fraction, which is then used for the first purification step. For membrane proteins, in most cases the membrane fraction is isolated after cell lysis. When you plan to purify proteins that are expressed in the E. coli periplasm, you can subject the cells to an osmotic shock to specifically isolate the E. coli periplasmic fraction.
Generally, a combination of various chromatographic techniques is used during the protein purification process. In many cases, the first step will be an affinity chromatography step, depending on the affinity tag you have chosen during the construct design. If a protease cleavage site has been included between the affinity tag and the protein of interest, this specific protease can be used to remove the affinity tag either immediately after the affinity chromatography step or later on in the purification process. To increase purity, a second chromatographic step such as ion exchange chromatography or hydrophobic interaction chromatography can be used. As a final polishing step, a size exclusion chromatography is usually performed, as this also immediately serves as a quality control step to assess of the oligomerization state of the protein(s) of interest.
|Biorecognition (ligand specificity)
|Tandem purification: compatible buffers?
|Ion exchange chromatography (IEX)
|Protein stable in low salt?
|Size exclusion chromatography (SEC)
|Size and shape
|Dilution of sample
|Hydrophobic interaction Chromatography (HIC)
|Protein not precipitated by high salt?
|Reverse Phase Chromatography (RPC)
|Protein can be destabilised by organic solvent! (mainly used for peptides)
During the purification process, you can monitor the results of all individual steps by SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis). Coomassie staining is still one of the most popular methods to visualize the protein bands on the gel, although other technologies such as silver staining, fluorescent staining and stain-free visualization (dependent on the presence of Trp residues) can be used as well. To verify the identity of your protein of interest western blotting with a specific antibody (or an antibody against the protein tag) can be performed as well. Although western blotting possesses a high specificity and sensitivity, antibody quality can be a bottleneck. Our preferred method of confirming protein identity is mass spectrometry, either via in-gel analysis of a protein band or in-solution analysis of the protein sample. For this quality control step in the protein production process, we collaborate with the EMBL Proteomics Core Facility.
The most commonly used method for determining the protein concentration is measuring the absorption at 280 nm in a spectrophotometer. At this wavelength, the aromatic amino acids Trp and Tyr exhibit strong light absorption, although cysteine groups forming disulfide bonds also absorb, but to a much lesser extent.
The law of Lambert-Beer describes the relationship between the absorbance (A) of a protein and the specific extinction coefficient (ε), the concentration (c) and the path length (L) of the incident light:
A = ε x c x L
A: absorbance of the sample
ε: molar extinction coefficient of the protein in M-1 cm-1 (can be calculated using the ProtParam server)
c: protein concentration in M
L: pathlength of the light in cm
If other components are present in the sample that also absorb at 280 nm (nucleic acids, chromophores, detergents etc.), other concentration determination methods such as Bradford, BCA, Biuret and Lowry can be used as well. These assays generally make use of the binding or formation of a chromphore in the presence of soluble protein and then measuring the absorbance of this chromophore to determine the concentration. Each of these assays has their own specific advantages and disadvantages regarding accuracy, robustness or compatibility with various buffer components though.
After you have invested all the hard work to express and purify your target protein, you should not forget to think about how you want to store your purified sample. The optimal storage method and conditions strongly depend on the protein’s specific characteristics and stability, the time you need to store it for and the planned downstream applications. In general it is important to avoid storage conditions that are close to the stability limits of the protein (e.g. extreme pH or pH values close to the isoelectric point of the protein). Furthermore, you also want to avoid the addition of compounds that might interfere with your downstream application and would therefore have to be removed prior to performing your experiments.
For determining buffer conditions in which your protein is stable, thermal shift assays such as thermofluor or nano-DSF can be used. This way you can screen various buffering reagents, different pH conditions and a variety of additives.
For short-term storage (~24h), most proteins can be kept at 4ºC. For long-term storage, protein samples are typically kept at -20ºC or -80ºC.
Protein storage at -20ºC usually requires the addition of 50% glycerol to your sample to avoid freezing at this temperature. If we plan to store a protein at -20ºC, we generally run the final size exclusion chromatography step in 2x storage buffer and then dilute the sample 1:1 with 100% glycerol. Alternatively, the protein sample can also be dialysed against the storage buffer already containing 50% glycerol. Proteins stored at -20ºC are often stable for several months, although the exact time frame is protein-dependent and should be determined experimentally.
Protein samples stored at -80ºC will be frozen. As repeated freeze-thaw cycles generally have a negative influence on protein samples, it’s best to prepare small-sized aliquots that will be used up during the course of an experiment. 5-10% glycerol or other additives that protect against the effect of freezing and thawing can be added as well. After preparing your protein sample aliquots, it’s important to flash-freeze them in liquid nitrogen before moving them into the -80ºC freezer for long-term storage. Many proteins are stable for months to years when stored in appropriate conditions at -80ºC, but the exact time frame again varies from protein to protein and should be determined experimentally.
If your protein of interest cannot be properly folded inside the cell, it might accumulate in insoluble intracellular aggregates called inclusion bodies. Proteins inside inclusion bodies are mostly in a inactive state.
Generally, we try to avoid having to purify recombinant proteins from inclusion bodies, as in vitro refolding can be complicated and often requires extensive screening of refolding conditions. Even if some soluble protein can be obtained after refolding, it’s imperative to assess your protein is properly folded, not aggregated and biologically active.
If a proper refolding protocol can be established, there can be some advantages to protein production in inclusion bodies as well:
The first step in the purification of insoluble protein from inclusion bodies is the isolation and solubilization of the inclusion bodies. After cell lysis and centrifugation, the inclusion bodies will be present in the pellet. Usually, a washing step with a low concentration of chaotropic agents (0.5 – 1.0 M of urea or guanidinium hydrochloride) or with detergents (e.g. 1% Triton-X100) is performed to remove contaminants. The washed inclusion bodies are then solubilized in a buffer containing 8 M urea or 6 M guanidinium hydrochloride. Often a reducing agent is added as well to keep the cysteine residues in a reduced state and break incorrect disulfide bonds that might have formed during the preparation. After solubilization, another centrifugation step is required to remove any remaining aggregates. The solubilized inclusion body fraction can then be used for further purification and refolding.
Some protein chromatography methods (for example IMAC) can be carried out under denaturing conditions as well and can therefore be used for the purification of denatured protein from solubilized inclusion bodies. The compatibility of the chromatography resin with chaotropic agents is usually described in the manufacturer’s manuals.
Various methods exist for the in vitro refolding process, such as dialysis, slow or rapid dilution and chromatographic refolding. The most optimal refolding method and conditions are protein-dependent and need to be established experimentally for each individual protein you wish to refold. If the protein contains disulfide bonds, the refolding buffer also needs to be supplemented with a redox system. The addition of a mixture of reduced and oxidized forms of a low molecular weight thiol reagent can provide the appropriate redox potential to allow formation and reshuffling of disulfide bonds. The most commonly used redox shuffling reagents are reduced and oxidized glutathione, although other options (e.g. cysteine and cysteamine) exist as well.
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Wingfield P.T., Palmer I. and Liang S.M. (2014) Folding and Purification of Insoluble (Inclusion Body) Proteins from Escherichia coli. Curr Protoc Protein Sci. 78: 6.5.1-6.5.30
Singh A., Upadhyay V., Upadhyay A.K., Singh S.M. and Panda A.K. (2015) Protein recovery from inclusion bodies of Escherichiacoli using mild solubilization process. Microb Cell Fact. 14:41
Hoffmann D., Ebrahimi M., Gerlach D., Salzig D. and Czermak P. (2018) Reassessment of inclusion body-based production as a versatile opportunity for difficult-to-express recombinant proteins. Crit Rev Biotechnol. 38(5):729-744