Dear all Thanks to for all the responses. I would continue my question for getting more advice.
I expressed a dimeric multi-protein (600 KDa). This protein was purified (affinity chromatography) in 4 different common buffers and run with the same SEC-buffer. We observe clear single size-exclusion peak referring to the dimer. We then measure activity of these 4 different protein yields in a well established assay. What we observe: Only one out of 4 has high activity and even one sample has NO activity at all. In all cases, the HPLC-SEC-MALS signal using 'protein sample from activity assay' confirms unequivocally a dimeric state. The observation of dimeric peak without activity means that the protein have a state that is either mis-folded - either locally or partially. Are there any *sensitive* methods to detect this behaviour - minor structural changes? Can circular-dichroism detect these changes? or any other methods for a large multi-domain protein? Many thanks, Markus Previous responses: ----------------------------------- I would not say *active* in the case of an enzyme, but probably *folded*. An enzyme may have many conformational states, some of which may represent inactive states, which will not be distinguished with gel filtration (because their hydrodynamic radii will be roughly the same), unless the inactivation involved unfolding and aggregation of the protein. -------------------------------- Is your enzyme pH sensitive? For example, if it has a histidine in the active site and most of the buffer conditions you are testing are below pH 6, you may be looking at a well folded protein that just isn't active because you've protonated the active site residue. Or it could be that the buffers you are testing are binding to your protein and sterically interfering with your substrate? It doesn't mean that your protein isn't folded or even inactive if you have just blocked the binding site, merely inhibited. There could be all kinds of reasons that changing buffers could change the activity of the protein without unfolding the protein itself. Another example is that people often use phosphate buffer in purification, but if the enzyme requires a Mg, you could be inadvertently pulling that out of the enzyme by using phosphate buffer (or using sulfate with an enzyme that requires Ca, etc). I'm sure it is possible that there are many enzymes in the PDB that are clearly well folded (have good structures) that are not in their fully active states due to the crystallisation conditions used to obtain the crystals. We are usually capturing a single state of a protein which usually has to be mobile to perform its enzymatic function. ----------------------- You can speak for yourself, but not for me. I do not assume activity from a gel; that's what assays are for.Different buffers: it could be you have a cofactor, perhaps a metal. The best practice is to document what you do in your publications to the extent that a reader could duplicate your results. ----------------------- There are lots of examples in the PDB of incorrect structures. And a single peak on SE doesnt guaruntee correctly folded protein. What were the differences between the buffers? pH, ionic strength and additives all matter for enzyme activity, and many buffers do bind to active sites thus affecting activity (despite the general attempt to use large molecules which are unlikely to bind in the cases of the Good buffers). All that being said, the idea of a single, correctly folded conformation of an enzyme/protein is an oversimplification used in textbooks rather than the more complicated picture held by experts in the field. -------------------------- I believe the strong assumption in the community is that a clear single peak of appropriate Mw is a clear indication of pure protein, worthy intensive crystallization efforts. Whether it is active is another question and this should be measured.For your analysis, it is not important in which buffers the protein is not active, but whether the protein you purified is active in the buffer (maybe without precipitant) you used for crystallization.A single apo structure is usually not enough to determine the catalytic mechanism of an enzyme, you usually need some substrate-, transition state- product- (analog) structures as well. If your protein is active in the crystallization buffer and the ligand complexes make chemical sense, you can be pretty sure that you have crystallized the right conformation.If your protein is not active in the crystallization buffer, you must critically analyze the structure, if it makes chemical sense and if you can explain the absence of activity (e.g. pH far from optimum; inhibitor bound in the active site). I am currently working on an enzyme who's active site loves all kinds of substituted and unsubstituted phosphates, sulfates etc. so it is not active in a wide range of buffers like phosphate, MES, MOPS, HEPES etc. However, the crystal structures still represent the active conformation, the active site is just blocked by some buffer component.Other proteins (proteases) can only be crystallized in an inactive form, since active they chew themselves to pieces. ######################################################################## To unsubscribe from the CCP4BB list, click the following link: https://www.jiscmail.ac.uk/cgi-bin/webadmin?SUBED1=CCP4BB&A=1
