Dear CCP4BB, Forgive me for soap boxing, but yesterday the first structure of a GPCR/Gprotein complex was released (PDB: http://www.rcsb.org/pdb/explore/explore.do?structureId=3SN6, article: http://www.nature.com/nature/journal/vnfv/ncurrent/full/nature10361.html). Recently a student preparing for their dissertation asked this board for its opinion about the most significant recent structural accomplishment, and among many things the progress on GPCRs was mentioned (albeit as being cute, I think). Reading this work from Kobilka and Sunahara's two groups, I am floored by what it must have taken to achieve this - particularly if you know how hard and how long people have tried (both past and present) to get a GPCR/Gprotein complex structure. It is my opinion that this structure was something of a holy grail for the GPCR community. So, even if you don't usually follow the developments of membrane crystallography, I wanted to invite your attention to this historical achievement in the GPCR field; and I hope you will join me in congratulating the scientist involved in this work. Cheers~ ~Justin
Hi Alex, I read Filip's comment about volume not as a path length argument, but about concentration uncertainty in mixing small volumes to dilute a sample down before measuring it (?). I have never had to make a dilution for my nanodrop (my proteins are usually not that concentrated), but I could see his point if I did have to. As for the variance between samples, I don't know about 25%, but I have observed multiple readings to have variance. I always take 3 readings on my nanodrop and then average them to deal with the variance I see. I don't mind doing this because the instrument is so fast, and I don't mind the cost at 6 ul of sample total. The most variance I have seen is usually in spin columns, where I will be doing a buffer exchange from a storage buffer (sometimes at ca. 20% glycerol) into an assay or xstal buffer, and I have wondered to myself if the variance I see could be due to incomplete mixing of a protein sample betwen a viscous buffer at the bottom with the rest of the buffer. I don't know how often other people find themselves in a situation where they may be sampling their 2 ul from a micro-environment that is not homogenous with the rest of the sample, but with small volumes I think that be a problem. Food for thought. Filip, I would buy a nanodrop. It is much better than a Bradford/cuvette and your students will love you for it. Cheers~ ~Justin Quoting aaleshin aales...@burnham.org: Filip, 25% accuracy is observed only for very diluted (OD280 0.1) or concentrated samples. But those sample a rarely used for ITC or CD. The concentrated samples require dilution but a regular spec does it too. Since the light passway is very short in Nanodrop it is accurate with more concentrated samples, which we crystallographers use, so Nanodrop is ideal instrument for our trade. If the drop is within recommended volume like 1-2 ul for our model, its size has a very small influence on the measurement. Cuvettes will give a better accuracy provided you clean them properly. I hated those times when I had to measure a concentration because of a need to wash a cuvette. In a biological lab they are always dirty. We switched to plastic disposable cuvettes for that reason... Alex On Jun 16, 2011, at 1:06 PM, Filip Van Petegem wrote: 25% is not acceptable for ITC or CD experiments though... I was just sharing our bad experience with a demo nanodrop we had. Even if evaporation is not an issue, one has to take pipetting errors into account when dealing with small volumes. The relative error on 1-2ul is a lot bigger than on 50ul. Unless you want to pre-mix 50ul and use a small quantity of that, which defeats the purpose of miniaturization... It all depends on your applications and sample availability, but if you want a very accurate measurement, miniaturized volumes just won't get you the same accuracy. Cuvettes will give a better accuracy provided you clean them properly. Just some water or EtOH is *not* enough... Filip Van Petegem On Thu, Jun 16, 2011 at 12:52 PM, aaleshin aales...@burnham.org wrote: I also like our Nanodrop, but I do not recommend using it for Bradford measurements. The 25% accuracy mentioned by Flip is pretty good for biological samples. Using 50 ul cuvette in a traditional spectrophotometer will not give this accuracy because cleanness of the cuvette will be a big issue... Alex On Jun 16, 2011, at 12:43 PM, Oganesyan, Vaheh wrote: I completely disagree with Filip’s assessment. I’ve been using nanodrop nearly 5 years and never had inconsistency issues. If you work at reasonable speed (if you put a drop there then lower the lever and click measure before you do anything else) there will be no issues. At very high concentrations the accuracy and therefore consistency may become lower. Concentrations between 5 and 10 mg/ml should be fine. The instrument is pricey though. Vaheh From: CCP4 bulletin board [mailto:CCP4BB@JISCMAIL.AC.UK] On Behalf Of Filip Van Petegem Sent: Thursday, June 16, 2011 3:34 PM To: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] Nanodrop versus Nanophotomter Pearl versus good old Bradford. Dear Arnon, the Bradford method is not recommended for accurate measurements. The readings are strongly dependent on the amino acid composition. A much better method is using the absorption at 280nm under denaturing conditions (6M Guanidine), and using calculated extinction coefficients based on the composition of mostly Tyrosine and Tryptophan residues (+ disulfide bonds). This method is also old (Edelhoch, 1967), but very reliable. One thing about the nanodrop: smaller volume = more evaporation. On the demo we've had, I was so unimpressed with the precision (25% variability between two consecutive measurement) that we didn't consider this instrument at all. So unless you just want a 'rough' estimate, I wouldn't
Hi Harvey, Well, knowing nothing about your protein, allow me to ruminate anyway... It sounds like you are exploring the possibility of a metal ion or other cofactor being lost. This is a reasonable first thing to check, but your buffer exchange steps should allow small cofactors (smaller than most proteins that is) to pass through your membrane and away from your protein. This suggests that your loss of activity is due to the loss of something the size of your protein. Four things come to mind right away. 1) The least exotic possibility I can think of is maybe your protein is inactive all along according to your assay (your assay could have a problem in it, I would suggest trouble shooting your assay as a first step). This could result in your relatively dirtier prep falsely reporting activity because of another protein component (i.e. an impurity) that is active according to your assay, and then lost later during your purification. 2) This next idea seems unlikely, but you asked so... Could there be another protein component missing that is necessary for activity that you don't know about? This protein would be lost during purification resulting in an inactive form or your protein. 3) Probably another red herring here... Maybe your protein is not stable without lots of other proteins around. I have personally seen proteins that go to pot at low concentrations, but are very stable at high concentrations, for which this sort of reasoning is invoked. You could try adding Arg or other amino acids to keep it folded. 4) Is your protein active in a cleaved form? I have seen kinases with competent kinase domains in the absence of regulatory domains. If you run an activity assay that included the cleaved form of your protein, and then lose this cleaved form later after purifying away the cleaved protein, it would appear that you have lost activity. The most important advice I can give you is to pay attention to what your assays are really telling you, not what you think they are telling you because of useful assumptions we all make, but what the data really reports. For example, your activity assay shows no activity, the problem could be your protein, or a component of the assay, it is a bad idea to assume the protein is the only place something could be wrong. A factual analysis will hopefully allow you to trace back what you really know and where things could be going wrong. Hope this doesn't give you too many gooses to chase, hopefully somewhere in here is a spark to help you reason yourself out of your problem. Cheers~ ~Justin Quoting Harvey Rodriguez h.rodriguez.x...@gmail.com: Dear all, Recently, I came across an obstacle on the purification and acitivty measurement of my protein. My protein was expressed with an C terminal His tag in the HEK 293T cells and purified by nickel affinity, anion exchange and size exclucion chromatography. For every purification step, I preserved some sample to test the activty. Strikingly, the protein retains activity after nickel affinity column even for three days but lost almost all the activty immediately after Mono Q and SEC. Therefore, I speculated that something (metal ion or co-factor) binding to the protein was striped by the Mono Q column. Then I skipped this step and only use the SEC for further purification. However, the protein is still not active no matter what buffer I use, eg. Tris,Hepes or PBS. The protein I purified by nickel column is also in the PBS buffer and no additive was added. Buffer exchange in the concentrator doesn't affect the activity of the protein. Can anyone explain why anion exchange or size exclucion chromatography destroy the activity of the protein? Any comment or proposal is appreciated! Harvey
Dear Community, In trying to trouble shoot an experiment I have become interested in the cellular process that regulates the insertion and proper orientation of membrane proteins. I am looking for references for how a GPCR is correctly oriented during expression (i.e. the extra cellular domain ends up extra cellularly oriented instead of a 50/50 mix in and out), my intuition is that there must be an N-terminal sequence that directs this process, but I am having no luck finding information on what this sequence is for GPCRs, what players are involved or how orientation is thought to be controlled. Any suggestions? This is all spurred by my wanting to use phage display with a protein that binds to the intracellular side of a GPCR, but of course that is the hard side to present to the outside of a cell so I need to figure out how to flip these guys around. I have thought about adding a new TM helix before TM1 (or removing TM1) to flip these guys, but was hoping there might be another way around that doesn't involve such massive architectural rearrangement such as simply clipping the N-terminal sequence responsible for proper orientation (if such a thing exists). Cheers~ ~Justin
Dear Community, I am looking for a tool that can convert a potential MR search model to match the residue number and a.a. type of my actual protein. Specifically, I have a homolog structure, with slightly different start and stop residues, and several non identical a.a.s relative to my protein. Can anyone direct me to a tool that can renumber the a.a.s in my search model, and stub nonidentical a.a.s to Ala? I recall there is a program in CCP4 that will renumber, leaving the user to change a.a. type, but this problem seams like such a common one that I am surprised there is not a utility that would perform both functions. Cheers~ ~Justin
Hi Liu, If I understand your question correctly, youre asking how different do two structures need to be for one to be new'. If by new you mean a new fold, then the answer is NO. Your structure and the homolog have the same fold. However, if your structure is the first structure of a protein in a new class, then your structure is a new insight for that reason (e.g. it is the first structure of a Unobtainium-metalloprotease). If it is not the first structure of a protein from a new class, lets say a previous structure of Unobtainium-metalloprotease has been solved using H. sapiens' sequence, but your protein is the first D. melanogaster ortholog solved, then your structure is a new insight for that reason. So, in a nut-shell, I guess what I am saying is that your protein is not a new fold, but is almost certainly new by some qualification, and you will know best what that qualification is. I hope that helps, cheers and happy holidays~ ~Justin On 20/12/2010 10:49, Liu Zhao wrote: The structure of my protein is as shown as the purple one. Another one ,as shown as green,is homologous .But the structure of my protein can't be obtained by using molecular replacement. And both structures have much different, especially in B chain. If my structure is a new one? thank you for help.
Hi Sebastiano, I have had some experience with protein:protein complexes with KD ~ 10-1 uM, kinetic characterization and trying to purify a complex of these proteins using SEC. While I would say that if you have reliable evidence from SPR that you have a fast on (high Kon), then you must have a fast off (high Koff) because by definition KD = 10 E-6 = Koff/Kon. However, I have observed several systems where you have a KD ~ 10-1 uM, but the kinetics are not fast on/fast off. In my experience, I have never seen anything in the crystal structures of the weak affinity complexes I have solved that would coorelate B-factors to Kon/Koff, and while it might be tempting for you to draw this comparison in your structure, I would warn that this is too large a leap without further (non-anecdotal) evidence. As a further note, during SEC purification of complexes, I have observed that you generally have to have the complexes at at least 5 to 10-fold higher initial concentration if you want to purify the complex, which you are only pushing with your 80-100 uM high end concentration. A colleague of mine once told me this is due to a 5 to 10-fold dilution effect upon addition to the column, but I have never verified this nor read any primary source that validated this so I cannot supply a reference (others might be able to help here). Good luck and cheers~ ~Justin Quoting Sebastiano Pasqualato sebastiano.pasqual...@ifom-ieo-campus.it: Hi all, I have a crystallographical/biochemical problem, and maybe some of you guys can help me out. We have recently crystallized a protein:protein complex, whose Kd has been measured being ca. 10 uM (both by fluorescence polarization and surface plasmon resonance). Despite the 'decent' affinity, we couldn't purify an homogeneous complex in size exclusion chromatography, even mixing the protein at concentrations up to 80-100 uM each. We explained this behavior by assuming that extremely high Kon/Koff values combine to give this 10 uM affinity, and the high Koff value would account for the dissociation going on during size exclusion chromatography. We have partial evidence for this from the SPR curves, although we haven't actually measured the Kon/Koff values. We eventually managed to solve the crystal structure of the complex by mixing the two proteins (we had to add an excess of one of them to get good diffraction data). Once solved the structure (which makes perfect biological sense and has been validated), we get mean B factors for one of the component (the larger) much lower than those of the other component (the smaller one, which we had in excess). We're talking about 48 Å^2 vs. 75 Å^2. I was wondering if anybody has had some similar cases, or has any hint on the possible relationship it might (or might not) exist between high a Koff value and high B factors (a relationship we are tempted to draw). Thanks in advance, best regards, ciao s -- Sebastiano Pasqualato, PhD IFOM-IEO Campus Dipartimento di Oncologia Sperimentale Istituto Europeo di Oncologia via Adamello, 16 20139 - Milano Italy tel +39 02 9437 5094 fax +39 02 9437 5990
Hi Francis, I might save you some time by telling you up front you should just go back and purify your compound to remove the impurity, you dont even need to read the rest of this, just go. Along the lines of what Savvas was saying, with any equilibrium binding assay between two direct competitors (Y is the impurity and Z is your analyte), if you are working at concentrations above the KD then the resultant complexes (XY and XZ) will partition according to their relative association strengths (dG) and concentrations. So, if Y and Z have equivalent dG values, then the concentration of XY ([XY]) and [XZ] will be a function of [Y] and [Z], if [Y]=[Z] in this circumstance then [XY]=[XZ]. If dGy dGz or [Y] [Z], then you are in the clear. This is why going back to purify Z from Y is a good idea. Now,the great thing about ITC is of course that you can get dG, dH and -TdS in one experiment, but this is also going to bite you in the butt here since you will simultaneously be determining dG, dH and -TdS for both Y and Z, which leaves you will more unknowns that you have data to solve for unless you independently know [X], [Y], [Z] and dG, dH and -TdS for XY or XZ. In fact, the circumstance where you know [X], [Y], [Z] and dG, dH and -TdS for XY or XZ is what Savvas is describing with displacement assays, and unless I am misunderstanding your situation it sounds like you dont know these parameters. For that reason I would not qualify this as a displacement assay, but instead just as a poorly controlled experiment . Now, you might be able to do the experiment with pure Y binding to X to determine dG, dH and TdS, then perform the proposed experiment with impure Y and Z as a displacement binding, but this is going to still be a headache because your uncertainty will be greater, you will not have as accurate a measure of [Y] and [Z] as when they are pure, and since your your direct signal (dH) is going to be from the formation of both XY and XZ (dHtotal = dHy + dHz) S/N will be equal to or less than the experiment with pure Y or pure Z (my nanny used to say 'dont do good experiments with bad reagents, youll just waste time', she was very wise). Hope that helps, cheers~ ~Justin Quoting Savvas Savvides savvas.savvi...@ugent.be: Hi Francis I guess it depends on how much residual high-affinity binder you have in the mixture and what the difference in affinity is between Y and deriv-Y. Another issue is of course whether Y and derY compete for the same binding site and have the same stoichiometry. A well designed displacement ITC experiment and comparisons thereof with ITC data for your high-affinity binder should lead to some good answers. Knowing the ratio of Y vs deriv-Y in your starting compound solution will be an advantage. A very useful reference in thinking about and carrying out displacement ITC in our group has been the one by Velazquez-Campoy and Freire. This article was specifically written to address the application of displacement titrations in ITC. We have applied this approach to address several types of questions concerning interactions in the uM-pM range. Velazquez-Campoy A, Freire E. Isothermal titration calorimetry to determine association constants for high-affinity ligands Nat Protoc. 2006;1(1):186-91. Best regards Savvas Savvas Savvides Unit for Structural Biology @ L-ProBE Ghent University K.L. Ledeganckstraat 35, 9000 Ghent, Belgium Ph. +32 (0)472 928 519 http://www.LProBE.ugent.be/xray.html On 24 Aug 2010, at 17:11, Francis E Reyes wrote: Hi All I'm curious the effect of small impurities in commercially synthesized compounds on ITC and its analysis. Say if compound Y is the high affinity binder, but you make a derivative that differs from a single functional group from Y (you used Y to make this new compound) and you never are able to completely get rid of Y. How does this affect the analysis of determining the derivative's affinity by ITC? References or personal experience is appreciated! F - Francis E. Reyes M.Sc. 215 UCB University of Colorado at Boulder gpg --keyserver pgp.mit.edu --recv-keys 67BA8D5D 8AE2 F2F4 90F7 9640 28BC 686F 78FD 6669 67BA 8D5D
Hi Ajit; One of our CRT monitors broke recently, and in the context of bemoaning the loss to a friend I was told that LCD monitors will not work for stereo viewing. I understood the reason to be related to the difference in refresh rates (?), with LCD's not being fast enough so that the viewer is left seeing ghosts. The effect, which I have not seen first hand, was described to me as capable of making most hapless stereo viewers very ill, very fast. I would encourage you to plumb the depths of knowledge on this subject further, but that is my simple understanding. Best wishes~ ~Justin Quoting Ajit Datta adat...@jhmi.edu: Hello everyone, Sorry for a non-CCP4 related question again. Can anyone let me know how to make stereo work on linux with Zalman monitor with Coot? Is it as simple as what we do with CRT monitors? Or do we need something else? We presently use CRT monitors on a Quadro FX 4600 graphics card. I would like to move to LCDs. Thanks for all inputs Ajit B.
Thanks to everyone for the info on Zalman monitors, sorry to have muddied the waters for you Ajit. Best wishes~ ~Justin Quoting Justin Hall hallj...@onid.orst.edu: Hi Ajit; One of our CRT monitors broke recently, and in the context of bemoaning the loss to a friend I was told that LCD monitors will not work for stereo viewing. I understood the reason to be related to the difference in refresh rates (?), with LCD's not being fast enough so that the viewer is left seeing ghosts. The effect, which I have not seen first hand, was described to me as capable of making most hapless stereo viewers very ill, very fast. I would encourage you to plumb the depths of knowledge on this subject further, but that is my simple understanding. Best wishes~ ~Justin Quoting Ajit Datta adat...@jhmi.edu: Hello everyone, Sorry for a non-CCP4 related question again. Can anyone let me know how to make stereo work on linux with Zalman monitor with Coot? Is it as simple as what we do with CRT monitors? Or do we need something else? We presently use CRT monitors on a Quadro FX 4600 graphics card. I would like to move to LCDs. Thanks for all inputs Ajit B.
Dear All; In response to my Anisotropic Diffraction In Refinement, which asked for suggestions for how best to proceed with refinement with an anisotropic data set, I received a large number of responses which overwhelmingly suggested using the UCLA Anisotropy Server (http://www.doe-mbi.ucla.edu/~sawaya/anisoscale/). The Anisotripy Server treats scaled/truncated data sets (I used Scala and the old Truncate program). Fo and SigFo are analyzed with respect to resolution in three dimensions and the data treated in three steps: 1) An elliptical resolution boundary is determined and applied. 2) A purely anisotropic B-factor is applied to the Fo and SigFo data to cause the data in all directions to fall off equally. 3) A negative isotropic B-factor is then applied to the structure factors to force the fall-off in the strongest direction to match that of the original data, effectively meaning that the data are not scaled to the mean but the weaker data are scaled up to match the strongest data. Application of a elliptical resolution boundary is justified because the resolution boundary from common integration programs (Denzo and Mosflm for example) is spherical where diffraction for anisotropic data is ellipsoidal. A spherical boundary would result in the inclusion of numerous poorly measured reflections in the higher resolution shells which effectively makes these data more noisy. Imposing an ellipsoidal resolution boundary is equivalent to removing noise from the higher resolution bins and is simply the anisotropic equivalent of the normal resolution limit truncation. However, I was confused by the second and third steps. The second step of application of anisotropic scale factors is appropriate if the refinement program does not include anisotropic scaling in its calculation of Fc, however modern refinement programs do this. Pavel Afonine touched on this in his CCP4BB general posting in response to my original posting where he noted that anisotropic scale factor[s] that [are] part of the total structure factor take care of this (https://www.jiscmail.ac.uk/cgi-bin/webadmin?A2=ind0909L=CCP4BBT=0F=S=P=8362). For the third step, applying a negative isotropic B-factor to modify the Fo is equivalent to sharpening the peaks in your maps and this can be useful. However, applying the correction to Fo will also result in an inappropriate decrease in the average temperature factor of the resulting model. Since B-factors are used as a measure of the coordinate error of an atom, modifying your Fo means these low B factors will tend to confuse the users of that model into thinking its quality is better than it really is. If a sharper map makes identification of model errors easier, the map can be sharpened when it is calculated, without affecting the parameters in the PDB file. The latest versions of Coot, for example, allows you to sharpen any map that it calculates. I brought these points to the attention of the Anisotropy Server director (Michael Sawaya), who is now working to provide an option to omit steps 2 and 3 for users who do not what their structure factors modified. My thanks to everyone who responded to my original question, and to Dale Tronrud and Michael Sawaya in particular for valuable discussion.
Dear All; I am working with a data set which is anisotropic. The resolution limits are ~ 2.75 by 3.45 A. The I have integrated (using Mosflm) the data out to 2.75 A, the data therefore includes a mix of real (I/sig 1) and imaginary (I/sig ~1) data past the 3.45 A resolution bin. I am concerned that the presence of the poor quality data in the outer shells will cause my good data to 2.75 A resolution to be down weighted in refinement. Since anisotropic resolution limits do not seem to be an option, are there other tools that would allow proper weighting for this situation? Cheers~ ~Justin Hall Oregon State University