Our skills in custom protein purification were gained over many years of work with protein pharmaceuticals, drug targets, biological system components and biochemical reagents. All Protein Purification Services start with the analysis of physico-chemical and biological properties of a target protein resulting in the development of tailored procedures for its extraction, purification and characterization. Our purification strategy aims to achieve a homogeneous active protein preparation in two to three purification steps. This goal is reached by a thorough selection and optimization of the capture step, incorporation of a gel filtration step to remove aggregates, degradation products and other contaminants, selection of buffer conditions that stabilize biological activity and prevent product degradation. We routinely work with antibodies, antigens, enzymes, growth factors, DNA-binding proteins, membrane proteins, blood proteins and many more. Although most proteins require an individual approach, we are confident that we can handle your protein. We will purify it cost-efficiently, characterize it according to your specifications and deliver it to you in an active and application-compatible form.
CUSTOM PROTEIN PURIFICATION FEATURES
- purification of
- fusion and tagged proteins from recombinant sources
- native proteins from natural sources
- antibodies of different isotypes including IgMs
- membrane proteins
- common protein purification scales range from 0.001g to 5g
- proteins are purified according to client-specified purity
- protein purification methods from
- client-supplied protocols
- published protocols
- improved protocols from client-supplied or published protocols
- de novo protocols tailored to client's requirements
- any protein purification mode can be used
- gel filtration
- affinity (broad-spectrum)
- hydrophobic interaction
- refolding from inclusion bodies
- protein purification method development for transfer to a GMP facility
- dedicated columns are used in each project
- efficiency is provided by the automation and precision of AKTA systems from Amersham BioSciences (currently GE)
ANALYSIS OF INTERMEDIATE AND FINAL PURIFIED PROTEINS
SDS-PAGE and/or dot/Western blotting are routinely used for fraction analysis final protein purity is determined by densitometry from Coomassie-stained gels final products are supplied with a certificate of analysis, purification report and, if applicable, a batch record
- SDS-PAGE and/or dot/Western blotting are routinely used for fraction analysis
- final protein purity is determined by densitometry from Coomassie-stained gels
- final products are supplied with a certificate of analysis, purification report and, if applicable, a batch record
ALL CUSTOM PURIFIED PROTEINS ARE
- supplied with a certificate of analysis tailored to client's specification (read more Protein Characterization)
- provided at specified protein concentration
- formulated in a buffer that protects from protein degradation due to proteolysis, oxidation and shear stress
- dispensed into specified aliquot sizes
AUXILIARY PROTEIN PURIFICATION SERVICES INCLUDE
- protein refolding
- endotoxin removal (read more Endotoxin Removal)
- tag removal
- labeling & conjugation (read more Protein Labeling & Conjugation)
Useful Tools for a Protein Purification ProjectCalculate Protein Physical Properties (ProtParam)
Ammonium Sulfate Precipitation Tool
Solution Concentration Tool
Solution Dilution Calculator
Chromatography Volume and Flow Rate Calculator
Phosphate Buffer Calculator
pH Buffer Calculator
Predict Protein Folding Rate from Sequence
Selected Latest Developments in Protein Purification
Strategies for Optimizing the Production of Proteins and Peptides with Multiple Disulfide BondsYunqi Ma et al., Antibiotics (Basel). 2020 Sep; 9(9): 541.
Bacteria can produce recombinant proteins quickly and cost effectively. However, their physiological properties limit their use for the production of proteins in their native form, especially polypeptides that are subjected to major post-translational modifications. Proteins that rely on disulfide bridges for their stability are difficult to produce in Escherichia coli. The bacterium offers the least costly, simplest, and fastest method for protein production. However, it is difficult to produce proteins with a very large size. Saccharomyces cerevisiae and Pichia pastoris are the most commonly used yeast species for protein production. At a low expense, yeasts can offer high protein yields, generate proteins with a molecular weight greater than 50 kDa, extract signal sequences, and glycosylate proteins. Both eukaryotic and prokaryotic species maintain reducing conditions in the cytoplasm. Hence, the formation of disulfide bonds is inhibited. These bonds are formed in eukaryotic cells during the export cycle, under the oxidizing conditions of the endoplasmic reticulum. Bacteria do not have an advanced subcellular space, but in the oxidizing periplasm, they exhibit both export systems and enzymatic activities directed at the formation and quality of disulfide bonds. Here, we discuss current techniques used to target eukaryotic and prokaryotic species for the generation of correctly folded proteins with disulfide bonds.
The rapid “teabag” method for high-end purification of membrane proteinsJenny Hering et al., Sci Rep. 2020; 10: 16167.
Overproduction and purification of membrane proteins are generally challenging and time-consuming procedures due to low expression levels, misfolding, and low stability once extracted from the membrane. Reducing processing steps and shortening the timespan for purification represent attractive approaches to overcome some of these challenges. We have therefore compared a fast “teabag” purification method with conventional purification for five different membrane proteins (MraY, AQP10, ClC-1, PAR2 and KCC2). Notably, this new approach reduces the purification time significantly, and the quality of the purified membrane proteins is equal to or exceeds conventional methods as assessed by size exclusion chromatography, SDS-PAGE and downstream applications such as ITC, crystallization and cryo-EM. Furthermore, the method is scalable, applicable to a range of affinity resins and allows for parallelization. Consequently, the technique has the potential to substantially simplify purification efforts of membrane proteins in basic and applied sciences.
Covalent Functionalization of Bioengineered Polyhydroxyalkanoate Spheres Directed by Specific Protein-Protein InteractionsJin Xiang Wong et al., Front Bioeng Biotechnol. 2020; 8: 44.
Bioengineered polyhydroxyalkanoate (PHA) spheres assembled in engineered bacteria are showing promising potential in protein immobilization for high-value applications. Here, we have designed innovative streamlined approaches to add functional proteins from complex mixtures (e.g., without prior purification) to bioengineered PHA spheres directly harnessing the specificity of the SpyTag/SpyCatcher mediated protein ligation. Escherichia coli was engineered to assemble PHA spheres displaying the SpyCatcher domain while simultaneously producing a SpyTagged target protein, which was in vivo specifically ligated to the PHA spheres. To further demonstrate the specificity of this ligation reaction, we incubated isolated SpyCatcher-coated PHA spheres with cell lysates containing SpyTagged target protein, which also resulted in specific ligation mediating surface functionalization. An even cruder approach was used by lysing a mixture of cells, either producing PHA spheres or target protein, which resulted in specific surface functionalization suggesting that ligation between the SpyCatcher-coated PHA spheres and the SpyTagged target proteins is highly specific. To expand the design space of this general modular approach toward programmable multifunctionalization, e.g., one-pot construction of immobilized multienzyme cascade systems on PHA spheres, we designed various recombinant bimodular PHA spheres utilizing alternative Tag/Catcher pairs (e.g., SnoopTag/SnoopCatcher and SdyTag/SdyCatcher systems). One of our bimodular PHA spheres resulted in simultaneous multifunctionalization of plain PHA spheres in one-step with two differently tagged proteins under in vitro and ex vivo reaction conditions while remaining functional. Our bimodular PHA spheres also showed high orthogonality with the non-target peptide tag and exhibited decent robustness against repeated freeze-thaw treatment. We demonstrated the utility of these approaches by using a fluorescent protein, a monomeric amylase, and a dimeric organophosphate hydrolase as target proteins. We established a versatile toolbox for dynamic functionalization of PHA spheres for biomedical and industrial applications.