Lecture 5: Assessing Protein Identity Flashcards
How can we check protein purity?
We known that our protein is correctly purified if there is a single band on an SDS PAGE gel and a gel filtration column. After we know that there is one protein, we need to check that it is the correct protein.
• N-terminal sequencing.
• Amino acid analysis.
• Mass spectrometry.
• We can also check folding with circular dichroism. We can also see if the correct secondary structure is adopted.
How does N-terminal sequencing work?
N-terminal sequencing (also known as Edman degradation) is a technique for sequencing the N-terminus of a protein.
• The sample is prepared by running a gel and doing a Western blot with Coomassie.
• Amino acids are removed one at a time.
• Phenylisothiocyanate (PITC) is added to a peptide to form phenylthiocarbamyl (PTC)-peptide. This is done at pH 8.0 and at 45 degrees Celsius.
• The PTC group destabilises the N-terminal peptide bond. Acid treatment leads to the cleavage of this bond.
• An amino acid derivative (ATZ-derivative) is formed.
• The cleaved amino acid is converted into a more stable (PTH) form.
• The process is automated.
• After each cycle, the resultant amino acid can be separated by HPLC (high performance liquid chromatography) and identified.
• 5 residues is usually enough to confirm identity.
What is amino acid analysis?
Amino acid analysis can be used to assess the amino acid content of a sample.
• We can identify a protein by comparing amino acid ratios.
• This gives a good measure of the amount of protein which has been purified.
• The protein is hydrolysed into its component amino acids.
• Acid hydrolysis is the most common method. Salt and buffer are removed. The protein is then sealed in an argon atmosphere with HCl vapour. Tubes are incubated at 110 degrees for 24-72 hours.
• Some amino acids are damaged during this process.
• Trp and Cys are destroyed.
• Asn is converted to Asp so there is one peak for both.
• Gln is converted to Glu so there is one peak for both.
• Ser and Thr are partially destroyed so collect time points at 24, 48 and 72 hr then extrapolate back.
• Ile and Val hydrolyse slowly, so measure after 72 hr.
• After hydrolysis, the amino acids are added to PITC. There is a PTC label on each amino acid.
• Amino acids are separated by a C18 HPLC column. PTC can the be detected through 254nm absorbance of the PTC label.
• The column is calibrated using known quantities of amino acids.
• We need about 100-300 pmol of protein to perform AAA. The sample is normally supplied in solution, ideally without buffer salts.
How can we use proteolysis for looking at domain boundaries?
Proteolysis can be used to test whether our domain boundaries are correct.
• The original boundaries are educated guesses.
• We can use limited proteolysis to trim the protein or cut away stable fragments.
• We treat the pure protein with small quantities of proteases and take samples at different time points.
• We can use SDS PAGE to study the results and find protease resistant protein fragments. These can then be studied with NTS or MS.
• When we identify stable fragments, we may want to go back and redesign new domain boundaries.
• Conformational changes may change protease accessibility. For example arrestin cleavage changes in response to different peptides from the vasopressin receptor.
How can we select better buffer conditions?
The first time we purify a new protein, our choices of solution components are an educated guess.
• Once we have a pure protein sample, we can start experimenting with conditions.
• One of the best high-throughput methods of accomplishing this is thermal shift.
• Thermal shifts use fluorescent dyes which bind to hydrophobic areas, like the core of a protein. Examples of dyes are ANS and SYPRO orange.
• We can increase the temperature and look at the melting temperatures. We can use a real time PCR machine to perform this (it can run a temperature ramp and detect changes in fluorescence).
• We then experiment with different buffer conditions and see which ones will increase the melting temperatures by stabilising the protein.
• This can be done in a 96 well format, each one with different buffer components.
• We can also use thermal shift to screen for ligands which bind to proteins. These can be native ligands or small molecule inhibitors to use as therapeutics.
How can we select better buffer conditions?
The first time we purify a new protein, our choices of solution components are an educated guess.
• Once we have a pure protein sample, we can start experimenting with conditions.
• One of the best high-throughput methods of accomplishing this is thermal shift.
• Thermal shifts use fluorescent dyes which bind to hydrophobic areas, like the core of a protein. Examples of dyes are ANS and SYPRO orange.
• We can increase the temperature and look at the melting temperatures. We can use a real time PCR machine to perform this (it can run a temperature ramp and detect changes in fluorescence).
• We then experiment with different buffer conditions and see which ones will increase the melting temperatures by stabilising the protein.
• This can be done in a 96 well format, each one with different buffer components.
• We can also use thermal shift to screen for ligands which bind to proteins. These can be native ligands or small molecule inhibitors to use as therapeutics.
How can we check protein heterogeneity?
We want to see how homogenous our sample is. This could be based on charge, PTMs, degree of proteolysis, oligomerisation or aggregation.
• Charge heterogeneity can occur for a number of reasons. It could be variation in PTMs (with charged groups like phosphates) or it could be from proteolysis removing charged residues.
• We can test heterogeneity with isoelectric focusing.
• IEF gels have a pH gradient which is created by placing the anode in an a cidic buffer and the cathode in a basic buffer.
• The gel contains ampholytes. Ampholytes have different pI values. They move within the gel until they reach a pH equal to their pI. This reinforces the pH gradient across the gel. Some ampholytes are set into the gel after the gradient forms.
• When the protein is in a part of the gel when pH < pI, it will be positively charged and move towards the anode.
• When the protein is in a part of the gel when pH > pI, it will be negatively charged, and it will move towards the cathode.
• When the proteins reach a pH = pI, they will stop. The gel is run until all proteins reach equilibrium position.
• IEF can separate proteins based on phosphorylations or N-linked glycans or other small modifications.
What is 2D gel electrophoresis?
2D gel is an IEF which is followed by SDS PAGE.
• The mixture is loaded onto an IEF gel.
• When IEF is complete, the gel is loaded onto SDS PAGE.
