Lecture 4: Special cases Flashcards
How do we deal with extracellular protein PTMs?
Extracellular proteins are difficult because they often have disulphide bonds or glycosylation. They are also often multi-domain.
• We can use Origami E. coli to avoid glycosylation.
• We can use insect cells for simple glycosylation or mammalian cells if we want it to be more complex.
• Sometimes we need to regulate glycosylation if we want to make homogenous protein.
• We can add inhibitors which block the processing such as Kifunensine.
• Some mutant cells lack processing enzymes such as GnTI-.
• Or we can cleave glycans away using enzymes such as endo H.
• Extracellular proteins are purified from the culture medium, which has a very high volume. We need to separate and concentrate it by using tangential flow systems and buffer exchange.
• Tangential flow systems flow the sample past a permeable membrane. Molecules which are small enough can permeate into the new solution.
• Buffer exchange is based on SEC.
How do we perform refolding?
Refolding is used to refold inclusion bodies. Inclusion bodies are aggregates of misfolded, insoluble protein which are sometimes produced by E. coli.
• Inclusion bodies are first solubilised using a denaturing agent like guanidine hydrochloride or urea. The molecules are able to alter water structure and reduce the hydrophobic effect.
• A reducing agent like β-mercaptoethanol can break disulphide bonds.
• Solubilised material can be centrifuged to remove aggregates
• A His tag will still work if the protein is denatured. We can use it to purify the protein.
• We can gradually decrease the concentration of the denaturing agent to allow refolding. This method reduces aggregation, proteins are held on to the column until they are refolded.
• Another option is to elute the protein while still denatured and then dilute it into buffer which lacks the denaturing agents. The protein will have multiple opportunities to refold.
• The activin growth factor is expressed as an inclusion body in E. coli. It is then washed and solubilised before being diluted into refolding buffers and incubated for a week. The refolded protein is then purified using reverse phase and gel filtration chromatography.
How can we protect proteins from proteolysis with inclusion bodies?
Some proteins are especially susceptible to this. We want them to be expressed as inclusion bodies.
• Fusion of a ketosteroid isomerase (KSI) tag will accomplish this.
• This insoluble protein drives the fusion protein into inclusion bodies.
• The fusion protein can be purified under denaturing conditions.
• The KSI fusion is designed with met residues between peptides. It can then be cleaved by cyanogen bromide, which cuts at the C-terminal side of the Met in order to release the peptides.
• The peptides can finally be purified by RP chromatography.
How do we purify membrane proteins?
Membrane proteins are inherently a very difficult set of proteins to deal with.
• The proteins have evolved to function in a membrane, with lateral pressure from the lipid bilayer.
• We use detergents to extract them from the membrane environment.
• Detergents have hydrophobic and hydrophilic parts. They assemble into micelles in solution (when the critical micellar concentration is reached).
• Harsh detergents have small cmc values and short chains.
• Mild detergents have large cmc values and long chains.
• The cell is broken apart. We then use differential centrifugation. Low speed is use to pellet large cell fragments, nuclei and intact cells. We then centrifuge at a higher speed to pellet the membrane fragments. This second pellet contains the protein of interest.
• Detergents solubilise the membranes and then pack around the protein and the lipids.
• Membranes are incubated with the detergent for an hour and centrifugation is used to remove unsolubilised material.
• Solubilised protein is purified with standard techniques, but with the detergent present at > CMC in all buffers.
How can we design membrane proteins to make them more stable?
Even with the aforementioned techniques, membrane protein purification is tricky. We can make various modifications to improve the process of purification.
• This was done with the β-adrenergic receptor. Flexible loops and N and C termini were truncated. Glycosylation sites were removed.
• A major way to improve stability is thermal stabilisation. Each residue is mutagenized in turn and the proteins were tested for ability to survive a temperature increase.
• We can test stability with fluorescent size exclusion chromatography (FSEC). The protein is expressed with GFP fused to the C-terminus. Gel filtration is used to assess whether a protein is correctly folded.
• We screen different homologues, ligands, mutations or detergents to test which conditions give the most stable protein.
• We can use high throughput methods to test many of these combinations at once to see which gives the most stability. High throughput methods use automation and other tricks.
