Purifying and characterizing proteins

Transcription

1 Purifying and characterizing proteins Gel filtration chromatography Often called size-exclusion chromatography, this method separates proteins by their size (which is closely related to their molecular weight). A sample is run through a column that consists of beads, usually of agarose or polyacrylamide, which have have small holes that small proteins are able to enter but larger ones cannot. Because small proteins are able to enter the holes and travel through the beads, they take longer to reach the bottom of the column than larger proteins, which pass between the beads. The amount of protein coming off the bottom at any time can be crudely determined by taking a small sample and measuring it's absorbance of UV light at 280nm, which is absorbed by tyrosine and tryptophan residues in the protein. Gel filtration is better for separating large amounts of protein than gel electrophoresis, but is not as good at separating proteins of very similar molecular weight and is therefore not very useful for determining the mass of a protein. Ion exchange chromatography This method is used to purify proteins according to their overall charge, which depends on their amino acid composition. At ph 7, a protein will usually bind to a negatively charged group such as carboxylate if the net charge on the protein is positive. The more positive charges the protein has, the more tightly it will bind. Conversely, a negatively charged protein will bind to an amino group or something else with a positive charge. The crude protein solution is run through a gel containing agarose or cellulose beads with covalently attached charged groups, and proteins with opposite charge bind to those groups while proteins with the same charge elute. There are two ways of eluting the bound protein; by adding salt or by changing the ph. The ions in salt bind to the protein and neutralise it's charge, so that the protein is released from the gel. Alterations in ph can give finer control over the release of proteins and allows crude separation according to the isoelectric point (pi) - the ph at which a protein has no net charge. (Remember - proteins become more positively charged as the ph decreases due to the high H + concentration.) If we have a negatively charged protein bound to a positively charged matrix at ph 7, then decreasing the ph to 5 will elute all the proteins with pi values between 5 and 7. Those proteins with pi values higher than 7 will have been positively

2 charged when they were added to the column and would have eluted immediately, while those with pi values lower than 5 will still have a negative charge at ph 5 and remain bound to the column. By decreasing the ph of the column slowly using different buffers, it is possible to collect fractions containing proteins with progressively lower pi values. A sudden change in ph releases much more protein at the same time, and gives less pure samples. Affinity chromatography Affinity chromatography is used to give quite pure solutions of proteins, providing that the desired protein binds strongly to a known ligand. Generally, few proteins will have binding sites for the same ligand as the desired protein, which makes this a very powerful method. It is especially useful when used in conjunction with other methods, to separate out all the proteins with the same specificity but different net charge or molecular weight. The crude solution of protein is passed through a column containing beads (usually agarose) which have covalently attached ligand molecules, or an analogue of the ligand. Proteins that do not bind the ligand elute, while proteins that recognise the ligand bind to it and do not elute. When all the unbound protein has been washed through, the bound protein can be eluted by adding a solution containing free ligand. This competes with the solid phase-bound substrate for the binding sites on the protein, so when a protein binds a free ligand molecule is washed through.

3 For example, glutathione-s-transferase binds glutathione. This can be used to purify transgenic proteins. Also, where no substrate is available, specific antibodies to certain proteins can be immobolised on the column, then the protein is released by reducing the ph to about 4. The use of histidine tags and GST fusions is explained in the section on expressing recombinant genes. SDS-PAGE Polyacrylamide gel electrophoresis (PAGE) is a very useful method for separating small amounts of protein according to molecular weight, and allows measurement of the MW in comparison with marker proteins. Protein samples for PAGE are treated with SDS (sodium dodecyl sulphate), an anionic detergent and powerful denaturant. Addition of SDS to a protein destroys the tertiary structure by disrupting all the hydrogen bonds and Van der Waal's interactions, so that the protein is essentially a linear polypeptide. Many molecules of SDS bind to the denatured proteins, which gives them a large net negative charge. The number of SDS molecules that can bind to a protein is dependent only on the size of the protein, so pi values and tertiary structures have virtually no effect in PAGE. Protein samples are loaded in wells at the top of a polyacrylamide gel, which is sandwiched between two glass plates. A marker sample containing several proteins of known molecular weight is added to one or more wells to provide a standard. The gel is then suspended in a buffer and a voltage (typically 200V) is applied so that the bottom of the gel is positive relative to the top. The proteins, which are negatively charged, migrate through the gel at a rate which depends on their molecular weight. Because larger proteins can bind more SDS and are therefore more negatively charged, they might have been expected to migrate faster than small, less charged, proteins, but this is not the case because the polyacrylamide gel is like a sieve and impedes the progress of larger proteins. However, the extra charge does have an effect and the large proteins travel further than they would with less charge, so the scale is not linear. Usually, a sample dye is added to the protein solutions. This dye runs to the bottom of the gel slightly ahead of the smallest proteins, so that electrophoresis can be stopped when the samples have run far enough to get a good separation but not so far that some is lost from the bottom of

4 the gel. When the process is complete (illustrated on the left hand side of the diagram below), the gel is removed from the buffer and glass plates and soaked in a protein stain, a dye which binds to proteins but not to polyacrylamide. Coomassie blue is commonly used, because it gives distinct blue bands and can show up bands containing as little as 0.1µg of protein. A stained gel is shown below. The gel can then be coated in ovenproof film and baked dry for future reference. An alternative to Coomassie blue is autoradiography, which is used to visualise bands of radiolabelled protein on a gel. To determine the molecular weight of the unknown proteins, a graph of distance travelled against molecular weight is drawn using the results from the marker protein. The mw of any protein in another well is then found by drawing a line from the y (distance travelled) axis to the point where it intercepts the standard line, and reading off the x (molecular weight) value. Isoelectric focusing and 2D electrophoresis Isoelectric focusing is similar to SDS-PAGE, except that no SDS is added to the sample so all the proteins are in their native state. They are then electrophoresed on a polyacrylamide gel with a ph gradient. Proteins will then migrate across the gel to a point where the ph matches their pi, at which point they will stop as they have zero net charge (and are unaffected by the electric field) at their pi value. The voltage is applied so that the end of the gel with high ph is negative relative to the end with low ph - so an acidic protein (negative charge at ph 7) will travel towards the end with lower ph and positive charge, until the ph equals it's own pi. The ph gradient is formed before the protein is added by electrophoresing a mixture of small multicharged polymers called ampholytes, each of which has a slightly different pi. A stained isoelectric gel looks much like the SDS-PAGE gel above. Isolelectric focusing is often used in conjunction with SDS-PAGE to give a very powerful method of protein characterisation, by separating the sample of proteins first by isoelectric point and then by molecular weight. This method is powerful enough to positively resolve over 1000 proteins from a total protein sample of E.Coli, and is sometimes used in forensics to analyse samples and determine their source.

5 First, a protein sample is separated in one direction using isoelectric focusing. This lane of the gel is then placed horizontally across the top of an SDS-PAGE gel, so that the proteins are denatured in situ by the SDS, then a positive charge is applied to the bottom of the SDS-PAGE gel, so that the proteins in the isoelectric gel now run vertically down the gel as for normal SDS-PAGE. The result is a gel containing lots of protein spots (see illustration below), with pi measured horizontally and molecular weight measured vertically.