Re: [ccp4bb] Crystallisation of a minority fraction monomers
I just wanted to disagree with Roger's word choice, but not his argument (this is a flame-free response). Forget about packing and packable as there is no outside force doing the work. The molecules are just falling into a local energy minimum where favorable intra- and intermolecular interactions predominate. It is difference in the behavior of the ensemble versus of a solubilized, dispersed species (be it monomer or dimer). It is a phase behavior issue. Concerning Sebastian's case, while it is uncommon, the idea that a monomer has a crystalline phase state while the dimer does not is perfectly reasonable, and the crystals of the monomer grow due to mass action. I am sure the number of verified examples of this are limited. However, there are many cases where dimeric and tetrameric enzymes can be shown to be fully saturated with one or another bound substrate in solution, but show one or more empty active sites in the crystal. I know of several cases where this occurs, showing that selection of the species with the best set of favorable intra- and intermolecular interactions occurs. Regards, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 8, 2015, at 9:28 AM, Roger Rowlett rrowl...@colgate.edu wrote: The problem with crystallization is that is selects for the least soluble, most packable species. Sometimes that works against what you would like to know. That could include oligomerization state as well as conformational state. For example, some of the allosteric carbonic anhydrases stubbornly crystallize only in the T-state, despite crystallization conditions that are known to preferentially stabilize the R-state, and for which the predominant R-state population can be confirmed by other methods. Cheers, ___ Roger S. Rowlett Gordon Dorothy Kline Professor Department of Chemistry Colgate University 13 Oak Drive Hamilton, NY 13346 tel: (315)-228-7245 ofc: (315)-228-7395 fax: (315)-228-7935 email: rrowl...@colgate.edu On 4/8/2015 9:07 AM, Sebastiaan Werten wrote: Dear all, we are currently working on a protein that is known to exist in a monomer-dimer equilibrium. At the high concentrations used for crystallisation assays, the dimer is predominant and the monomer practically undetectable. Nevertheless, one of the crystal forms that we have obtained contains the monomeric species, not the dimer. I was wondering if anyone is aware of similar (published) cases, and if the phenomenon as such has been discussed in detail anywhere? I did literature searches but so far couldn't find anything. Any pointers would be much appreciated! Best wishes, Sebastiaan Werten.
Re: [ccp4bb] Crystallization of a minority fraction monomers
Thierry, I need to point out there is no outside work as it is one system, but with multiple phases. Protein and nucleic acids are not true crystals in the classic sense, but highly hydrated ordered colloids (in the 1930's some called them crystalloids because bulk water is such a major and critical component, unlike small molecule crystals). It is colloidal physical chemistry at work. Thus, the water argument for a force does not hold, rather the system just comes to an energy minimum where two stable phases are formed (one being the crystal). My complaint is that we use terms that imply the wrong physical behavior, which then obscure the true issues. For example, every protein is packable from a purely physical standpoint; physical shape is not the issue, but the balancing of favorable and unfavorable interactions is. Crystallization is a balance between many global and local interactions. Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 8, 2015, at 10:52 AM, Fischmann, Thierry thierry.fischm...@merck.com wrote: Some counter-arguments to Michael : There is an “outside force doing the work”: macromolecule crystallization except rare exceptions is driven by competition for water molecules between the macromolecule and the precipitant. The exceptions are crystallization against low salt buffer, in which case the process is driven by hydrophobic “forces”. And “packable” may play a role. A molecule which is of such shape and surface charge distribution that there is no way to pack it in a regular lattice will never crystallize. Regarding the dimer vs. monomer debate, crystallization acts as a purification step. It seems perfectly plausible that crystal growth would “select” the monomeric state if dimers cannot be included in the growing crystal lattice, regardless of whether one is more soluble than the other. It all comes down to the initial crystal seed favored by the crystallization conditions. On a separate note, protein which forms dimers in solution trend to be more soluble in dimeric state than as monomers because dimerization usually buries a significant hydrophobic patch of molecular surface. If crystallization was only “selecting for the least soluble” oligomeric state we would rarely crystallize proteins as dimers. Crystallization is such a confusing process J Thierry From: CCP4 bulletin board [mailto:CCP4BB@JISCMAIL.AC.UK] On Behalf Of R. M. Garavito Sent: Wednesday, April 08, 2015 10:04 AM To: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] Crystallisation of a minority fraction monomers I just wanted to disagree with Roger's word choice, but not his argument (this is a flame-free response). Forget about packing and packable as there is no outside force doing the work. The molecules are just falling into a local energy minimum where favorable intra- and intermolecular interactions predominate. It is difference in the behavior of the ensemble versus of a solubilized, dispersed species (be it monomer or dimer). It is a phase behavior issue. Concerning Sebastian's case, while it is uncommon, the idea that a monomer has a crystalline phase state while the dimer does not is perfectly reasonable, and the crystals of the monomer grow due to mass action. I am sure the number of verified examples of this are limited. However, there are many cases where dimeric and tetrameric enzymes can be shown to be fully saturated with one or another bound substrate in solution, but show one or more empty active sites in the crystal. I know of several cases where this occurs, showing that selection of the species with the best set of favorable intra- and intermolecular interactions occurs. Regards, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 8, 2015, at 9:28 AM, Roger Rowlett rrowl...@colgate.edu wrote: The problem with crystallization is that is selects for the least soluble, most packable species. Sometimes that works against what you would like to know. That could include oligomerization state as well as conformational state. For example, some of the allosteric carbonic anhydrases stubbornly crystallize only in the T-state, despite
Re: [ccp4bb] asymmetric homotrimer in the asu
Dear Hay, And your point is? I am not trying to be snarky (although I am just starting my morning coffee), but to bring up the fact that CCP4BB readers need more info to comment on your case, like space group, local interactions, and how packed is tightly packed. I have had two cases of trimers, as my students initially called them, that were actually a dimer and a half. The half dimer had its mate in another ASU. Can it be a biological monomer that just happened to crystallize 3 monomers to an ASU? Non-symmetric homo-oligomers are rare, but sadly cannot be absolutely confirmed by crystallography alone, but by good old biochemistry. The PISA website (http://www.ebi.ac.uk/msd-srv/prot_int/pistart.html) can give you estimations of the strengths of the interfacial interactions, but they are mere estimates. What does gel filtration say or cross linking? Does it fit with the biology/biochemistry expected of this protein? Anyway, have fun with your structure, but use a lot of skepticism in your interpretation. That will help you convince the reviewers. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Dec 11, 2014, at 7:27 AM, Hay Dvir hd...@tx.technion.ac.il wrote: Dear all, We have a structure of a rather tightly packed homotrimer protein in the ASU with no apparent crystallographic or non-crystallographic rotational symmetry between monomers. Attempting to establish the biological assembly, we are very interested to hear about additional similar cases you might know of. Thanks in advance, Hay --- Hay Dvir Ph. D. Head Technion Center for Structural Biology Technion Haifa 323, Israel Tel: +(972)-77-887-1901 Fax: +(972)-77-887-1935 E-mailhd...@technion.ac.il Website http://tcsb.technion.ac.il
Re: [ccp4bb] unknown densities
Yamei, This is not unusual, particularly for many proteins that bind nucleotide derivatives, especially GDP/GTP binding proteins, as Nat said. If it is GDP that is tightly bound at high occupancy, it should be quite easy to identify because of the pyrophosphates and the guanine ring. To build into, pop in a GDP molecule from another GDP/GTP binding protein structure; there is GDP .cif files in the CCP4 and Phenix libraries. Just be aware that there are at least two major conformers seen regarding the guanine ring (syn and anti). While in GDP/GTP binding proteins the ring conformer I believe is anti (the ring not over the ribose), in GDP-sugar enzymes, it can be syn (the ring over the ribose). Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Dec 8, 2014, at 10:15 PM, Yamei Yu ymyux...@gmail.com wrote: Hi all, We crystallised a small GTPase expressed in E. Coli and found some densities in GDP/GTP binding site. We didn’t add any GDP/GTP or GDP/GPD homologues during protein expression, purification and crystallisation. The resolution is not high, we couldn’t tell what it is. Is there any method to detect what it is? Thanks! Best wishes! yamei Yamei Yu State Key Laboratory of Biotherapy／Collaborative Innovation Center of Biotherapy, West China Hospital, Sichuan University,Chengdu,610041, P.R.China Tel: 15882013485 Email: ymyux...@gmail.com ymyux...@163.com yamei...@scu.edu.cn
Re: [ccp4bb] fastening crystal formation
Although three months is a long time, it is no completely unheard of, and does not require the invocation of proteolysis. The longest time I have heard of is ~1 yr, so count yourself lucky. However to get good advice, as well as to use it, you need to ask yourself several questions: 1. What kind of crystals are they? Protein, salt, etc? If they are salt, don't pursue this condition. 2. How many crystals did you get? One or 2 in a drop or a microcrystal shower. And of what kind? Single, well shaped, rosettes, needle clusters, or something that looks crystalline. Screen more broadly around your initial hit. 3. How many times have your tried to repeat this? Once, twice, or more? Did you try setups in duplicate? If so, did you get reproducible results? Have you actually screened around these conditions, varying each component systematically (PEG, salt, pH, buffer, etc.)? 4. What method did you use? And in what kind of container? For one thing, we don't completely trust the integrity of our setups for longer than 2 months. All containers leak water slowly, so when crystals take longer than 2 months to grow (a) the real conditions are at much higher values than you naively think (i.e., the drop has dried out more than you expected) or (b) other components are crystallizing, for example a zinc salt. It depends what else is in your protein buffer, as well. To quicken protein crystallization (which is not always a good thing), increase your protein concentration (by 1.5-2x) and/or PEG concentration (such as screening up to 40% PEGmme 550). Sadly, crystallization is a combination of thermodynamic and kinetic factors: you can get crystals (sometimes a single crystal only) when just outside the truly optimal conditions, but this may be only a sporadic event. You got to keep screening. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 31, 2014, at 6:05 AM, Vijaykumar Pillalamarri vijaypkuma...@gmail.com wrote: Dear all, I am trying to crystallize a 30 kD protein. Protein crystals are formed after 3 months. The crystals are formed in the following condition: 0.01M Zinc sulphate 0.1M MES monohydrate pH 6.5 25% v/v PEG monomethyl ether 550 Please suggest me how to grow these crystals faster. Thanking you -- Vijaykumar Pillalamarri, UGC-JRF, C/O: Dr. Anthony Addlagatta, Senior Scientist, CSIR-IICT, Tarnaka, Hyderabad, India-57 Mobile: +918886922975
Re: [ccp4bb] off-topic;Crystal cannot dissolved in buffer
Xiao, You could be the victim of the dreaded PEG cross-linking effect. One of the unfortunate by-products of keeping PEG stock solutions in water is that they will form peroxides and aldehydes. They will slowly cross-link the surface of some crystals. However, it is dependent on the nature of your protein's composition of surface residues, so not every protein crystal does this. I had one case where PEG4000 grown crystals would be resistant to dissolving and would easily bend (just like Herman related); the thinner rods would spring back straight. After placing the crystals into buffer known to dissolve them, I poked the crystals hard and the insides squeezed out like toothpaste, leaving an empty sack behind. The bottom-line is that fresh crystals diffracted better than old crystals because of this cross-linking. Suggestions: 1) make your PEG stocks up fresh or store them in the freezer as aliquots. 2) Remove oxidized PEGs from your stocks: see Ray et al. Biochemistry 1991, 30, 6866-6875 and Jurnak, J. Cryst. Growth, 76, 577-582, 1986. 3) Check to see if freshly grown crystals behave better. Best of luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Sep 25, 2014, at 7:53 PM, Xiao Xiao victor41...@gmail.com wrote: Hi everyone, Sorry for an off-topic question. I got a problem with crystal dissolving. Basically I got crystals of my protein in various conditions, most conditions contain PEGs but different salts. These crystals has very similar shape, so it should not be salt. Now I am trying to dissolve the crystal to make sure it is my protein, by SDS-PAGE and N-term sequencing. I washed the crystals in its original crystallization buffer few times then transfered them into regular buffer (500mM NaCl, 50mM HEPES 7.5) with or without 10mM DTT, however the crystal didn't dissolve. I then tried to heat it then add SDS loading buffer to run a gel, I did see very small amount of protein on the gel, at the correct position, but it's not enough for N-term sequencing. Is it normal for a protein crystal? And does anyone have any suggestion for dissolving such crystals?
Re: [ccp4bb] Removing PEG3350
Reza, If your protein is not too small (20 kDa), use a spin-column (i.e., desalting column) with G-25 sephedex. It is CHEAP, fast, and the recovery is good. We have even used them to adjust buffer concentrations or to remove micellar detergents; we have used protein concentrations up to 10 mg/mL, prior to crystallization. You will need a clinical centrifuge, G-25 Sephedex (reusable) equilibrated in your desired buffer, glass wool, 15 mL plastic conical centrifuge tube (reusable), 5 mL syringe barrel (reusable). Load the Sephedex into the 5 mL syringe barrel to make a gel bed ≥ 5x the sample volume Put into the 15 mL plastic conical centrifuge tube and spin for 2 min to remove excess buffer Remove the excess buffer from conical centrifuge tube Add your sample (0.2 mL to 1 mL) to the gel bed, then spin for 2 min to collect your cleaned sample. The sample will be slightly diluted by 10-20% depending on conditions. We have our students in our undergraduate biochem class do this all the time to remove ammonium sulfate from protein samples for assay. If they can do it. Or you can buy premade stuff to do the same thing. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 19, 2014, at 9:55 AM, Reza Khayat rkha...@ccny.cuny.edu wrote: Hi, Does anyone have a protocol for getting rid of PEG3350 from a protein sample? Best wishes, Reza Reza Khayat, PhD Assistant Professor The City College of New York Department of Chemistry, MR-1135 160 Convent Avenue New York, NY 10031 Tel. (212) 650-6070 www.khayatlab.org
[ccp4bb]
Prishant, Remember that concentrating by almost any method is a non-uniform process. In your case, right at the membrane the concentration is much higher than in the surrounding solution. As Chris says, frequent efforts to keep the solution well mixed can prevent precipitation. As you mix, look through the tube towards a light to see if you see schlieren patterns, indicative your protein concentrating at the membrane. In the best case, the protein concentration may shoot up too high at the membrane, which induces precipitation, and mixing more will help. In the really worst case scenario, your protein may not be very soluble under the conditions you have chosen (i.e., pH, salt, ionic strength). If so, you may need to rethink buffer conditions for concentrating in this manner (E.g., more salt, different buffer pH, chaotropic salts like LiCl, etc.). Good luck and don't despair yet. This happens quite often sometimes. Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 19, 2014, at 3:55 PM, Chris Fage cdf...@gmail.com wrote: Hi Prashant, I typically stop the centrifuge once in awhile and pipet up/down to prevent the sample from over-concentrating. Depending on how sensitive the sample is, you may want to do this once every 10-60 min. Hope this helps, Chris On Tue, Aug 19, 2014 at 1:42 PM, Prashant Deshmukh prashantbiophys...@gmail.com wrote: Hi, i am concentrating my protein using centricon filter, but it is precipitated soon. Please help me solving this problem. Thanks. Prashant Deshmukh Dept. of Biophysics, NIMHANS, Bangalore 560 029, E-mail:prashantbiophys...@gmail.com Mob.No.: +919620986525
[ccp4bb]
Avisek, Besides the advice that Bernhard and Remy have given you, beware of strange packing arrangements with low symmetry space groups and higher order oligomers. We have had several cases in P21 and C2 where tetramers pack with 6 or 8 molecules in the asymmetric unit (ASU), but could not be solved by MR using a tetramer model. Why? The true packing element was a dimer portion of the tetramer. Hence, the ASU with 6 monomers contained a tetramer and a half; the full unit cell then contained 3 tetramers, with the 2-fold axis of one tetramer sitting on a crystallographic axis. Likewise, the ASU with 2 monomers contained 1 tetramer and 2 half tetramers; the full unit cell then contained 2 tetramers, with the 2-fold axes of 2 tetramers sitting on a crystallographic axis. In each case, using the tetramer as a probe failed, but monomers and the appropriate dimers worked. The take-home message is don't get fixated on the tetramer as the only possible MR probe. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 17, 2014, at 2:54 AM, Avisek Mondal avisekmonda...@gmail.com wrote: Hello everyone, i am struggling with a problem.. My crystal was diffracted at 1.9A in P21 spacegoup with unit cell parameter a=87.7 b=93.9, c=111.78 ,beta=94.98 which contains 16 molecules per assymmetric unit (Molecular weight of the Protein+DNA =56 KDa. actually it is a complex of 40Kda tetramer and 23bpDNA ) .In solution, it shows tetrameric in nature The crystal structure of its homologous structure has been reported earlier (50%identical in amino acid seq.) and it was also a tetramer and its unit cell (also P21) was approximately 4 times less than mine. It showed 4 molecules per assymmetric unit. I didn't get any molecular replacement. All the programms are takings very long time to do it. Although the single crystal is untwinned i think it is a case of pseudosymmetry. Please help me if you have any good suggestion regarding molecular replacement other than experimental phasing.
Re: [ccp4bb] PEG dependent structural changes in crystal structure
Shiv, PEG1500 and PEG3350 are not the same because of the nature of their synthesis and manufacture. They are a polydisperse and semi-purified products of an ethylene oxide condensation. So while they are described as H-(CH2-CH2-O)n-OH, the value of n and its ranges are quite different. Some of the PEG molecules in both with be the same size, but PEG1500 will have a lot more small PEGs than PEG3350; it is the small PEGs that might bind more tightly to proteins. On top of this, there is the manufacturing issue. All PEG is made in bulk and then fractionated; different manufacturers have slightly different recipes and purification schemes (which are trade secrets, so good luck in finding more information from any company). Take a look at an old paper by Fran Jurnak (J. Cryst, Growth, 76, 577-582, 1986) for the trials and tribulations of working with PEG from different manufacturers of PEG (as well as how the purify it if you really get worried). She got at least 4 different space groups, and the key was the differing amount of phosphoric acid used in the neutralization step in PEG production. EACH batch of PEG from the same manufacturer can be different! So Herman's comment about pH should be considered. But as Herman said check the enzyme activity of your protein in each and get different batches of PEG to test. Is it a particular size of PEG or a PEG contaminant? Remember that stock solutions you may buy from Hampton or other X-ray specialty companies can be equally suspect because of where they get the bulk product to begin with. Even with purified PEGs (i.e., cleaned up of metal contaminants, aldehyde, and peroxided) age. So think about how old your stocks are. One last comment, just about your terminology. I would avoid the use of the phrase improper crystal packing. There is nothing really improper about it, just unfortunate from your point of view. The crystal packing is just the natural consequence of physical interactions in solution that you have prepared. You just have to be lucky sometimes. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 4, 2014, at 9:20 PM, shivendra singh shivendr...@gmail.com wrote: Dear All, I have been working on a protein which initially got crystallised in condition having PEG1500 as precipitant. The space group was P21 and got solved with reasonable Rfree. Analysis of its structure showed large deviation and very distinct active site architecture along with disorderedness in one of its long loop (no density) in comparison with the expected result, based on related homologous structures. The structure does not seem to be active with one of its active site residue moved apart from other catalytic amino acids. Also the substrate entry tunnel looks distorted. The purified enzyme used for crystallisation showed optimum activity in vitro. This led us to screen it again for some other crystallisation condition and got another crystal hit in condition having PEG3350 as precipitant. Rest of the components of crystallisation cocktail were same. The data belonged to P212121 space group. The regions which were disordered or distorted in earlier case were observed to be ordered and in their expected orientation and position. The enzyme is not reported to be in different structural or functional states as observed. I am wondering how the protein from the same batch showed two distinct structural organizations in conditions with varying PEGs. What may cause it to follow such transition. Whether it has some significant functional aspect or just a result of improper crystal packing. Thanks. Shiv
Re: [ccp4bb] equivalent osmotic pressures for xtal transfer
Frank, It is not just osmotic factors that need balancing, but the fact that polymers and salts don't always mix well. PEG and high salt can form 2-phase systems that don't disperse well, particularly in a crystal. Check out Ray et al. (Biochemistry 1991, 30, 6866-6875) where Bill Ray and and others from Purdue needed to exchange ~2.1 M ammonium sulfate in phosphoglucomutase crystals to various PEGs just to do the kind of experiments you suggest. It can be a pain, but osmolarity matching was not the big issue. As Bill was an excellent, old-time physical biochemist, he developed the system based on first-principles. You may also look up the work of Pierre Douzou and the cryoenzymology. Douzou collaborated with Petsko on some protein crystal work doing the same kind of exchange to reduce the crystal's freezing point. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 26, 2014, at 3:42 PM, Frank von Delft frank.vonde...@sgc.ox.ac.uk wrote: Hi all - The Google fails me, so I'll the CCP4BBoogle: Can anybody conjure up some of the references that describe the details of how to get a (say) PEG solution to have the same osmotic pressure as a (say) salt solution - in particular, this is for transferring a crystal from it's original salt condition into a more ligand-friendly PEG condition. I know there was something from the late nineties, where specific lookup tables were either shown or mentioned; but I've not been able to track it down. Thanks! phx
Re: [ccp4bb] Heavy Atom Phasing
Rhys, If crystals grow reproducibly and within a reasonable timeframe, I would always do co-crystallization, particularly if the HA is a good anomalous scatterer. We have had good success with this method, including a recent membrane protein structure. Even if you get crystals that are not isomorphous with the native, SIRAS is easy to do these days. Also broaden your soaking screens of HAs (if you haven't already), to include mercury compounds, which don't always need a Cys to bind well and love hydrophobic nooks and crannies, and metal clusters (like tantalum bromide). As this is a beta-barrel membrane protein, iodine might also be a way to go. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 27, 2014, at 5:48 PM, RHYS GRINTER r.grinte...@research.gla.ac.uk wrote: Hi All, I thought I might put a question to the community, with the hope of getting some tips of the best way to proceed with my heavy atom phasing problem. I'm working on solving the structure of an integral beta-barrel membrane protein of approximately 100 kDa. I've crystallised protein, growing some very flimsy needle like crystals, and collected datasets to around 3.1 A. I then produced selenomet derivative protein and repeated crystallisation trials in the same conditions and also repeated broad screens, however the derivative protein failed to produce crystals that diffracted beyond 10 A (in fact it barely crystallises at all). So I've moved on to heavy atom soaks and have had some success with tetrachloroplatinate and tetranitroplatinate compounds, in that the crystals didn't dissolve (as they did with gold and samarium compounds) and diffracted to some degree. I collected SAD data to around 6.5 A from these crystals and there seems to be anomolous signal. However, while I get a good CC of 0.4 from HYSS in phenix with this dataset and the phaser EP FOM is 0.56, the maps before and after DM are uninterpretable. I'm guessing the quality and resolution of the data I collected just aren't good enough (the data is reasonably anisotropic). I performed the metal soaking, by taking a small amount of the platinate salt and adding it to the crystallisation drop as the crystals are extremely fragile and don't stand up well to handling through a soaking or cryo solution. Leaving the crystals to soak for 48 hours and then, freezing them directly. The solution is on the border of cryoprotection (the conditions has PEG2000MME and PVP and the precipitants), but with native crystals this doesn't seem to be a parameter which affects diffraction. The crystals are very variable in performance, so while I feel that the heavy atom soaking has compromised their diffractability to a degree, inherent variation may play a part. What I was wondering is if some one with more experience than me found themselves in this position, how would they proceed? Questions which spring to mind are, how much heavy atom compound do people add and how long do they soak for? Is there anyway I can squeeze something out of the anomalous data I have, given I have 'reasonable' native data, or will poor quality data give spuriously positive statistics for heavy atom phasing? And are there any tricks people have experienced to improve performance of crystals like these (aside from the usual seeding, additives, different detergents etc which I have spend a fair bit of time on optimization already). Thanks in advance, Rhys
Re: [ccp4bb] cannot reproduce crystals
You may unleash a deluge of anecdotes and horror stories, but this is quite common. I have experienced this many times, and you just need to step back and ask yourself what is being done differently: 1. Are all materials used in the preparation of the protein the same (suppliers, sources, expression hosts, purification media, etc.)? I have had this happen several times when I have moved. If you have not rescreened with a broader condition matrix, do so. Slight changes in protein purity and quality (and they are not the same) can radically shift the crystallization behavior. 2. Did you or someone else improve the purification? Ultrapure pure protein sometimes crystallizes less well than less pure protein, for a number of reasons. One is that an extra step adding in could remove something critical. In number of cases in my lab, we have solved structures we thought were in the apo-form, but saw that ligands were bound. Making the true apo-form led to getting no crystals. 3. Are you trying to reproduce your work or work of others? If it is the latter, talk to the original people and don't rely on an old notebook. Very subtle differences crop up when doing lab work that impact crystallization, from how you make your PEG or buffer stocks (compared to those acquired in kits) to how someone sets up crystals. One case I know of, the group lost their crystals after a technician left. After some investigating, the slight pH difference caused by making up the buffer for a hanging drop reservoir differently was the source of the problem. Hope this helps, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 14, 2014, at 7:44 AM, dusky dew duskyde...@gmail.com wrote: Dear all, I am trying to reproduce some protein crystals. The protein I am getting after cutting the his tag is very pure. I am using the reported protein concentration. The cofactor and EDTA needs to be added externally. The condition has calcium acetate, peg 4k and sodium acetate buffer. Unfortunately I am getting oil separation and light ppt. I have no clue what is wrong. Please help!
Re: [ccp4bb] How to store PEG screens
Jerome, Does anyone know the best way to store crystallization screening blocks that contain PEG 3350? I would recommend storing them in a fridge or a clean coldroom (mold-free). Lower temperature and low light does help. Is it a good idea to freeze the PEG solutions at -80°C and thaw them before use? Good idea, if they were pre-aliquoted into useful volumes. We do that occasionally. However, -20°C is just as good. Would the freeze-thaw process considerably alter the PEG chain lengths? No, the real issue is the generation of oxygen reactive species that cause aldehyde and peroxide formation, which in turn can modify your protein. It also causes cross-linked polymer formation. Also avoid metal ion contamination. Like lipids, plain PEG solutions in water and most detergents with PEG head groups (C12E8, octyl-POE, Brij, Triton, Tween, etc.) should be stored under argon and at -20°C. So, if you are tempted to use the 5-year old bottle of Triton X-100 or old 50% stock of PEG on the bench top, caveat emptor. Take a look at an old paper by Fran Jurnak (J. Cryst, Growth, 76, 577-582, 1986) for the trials and tribulations of working with PEG from different manufacturers of PEG (as well as how the purify it if you really get worried). Regards, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 14, 2014, at 11:33 AM, Jerome Nwachukwu jnwac...@scripps.edu wrote: Dear all, I have 3 short questions about PEG solutions: Does anyone know the best way to store crystallization screening blocks that contain PEG 3350? Is it a good idea to freeze the PEG solutions at -80°C and thaw them before use? Would the freeze-thaw process considerably alter the PEG chain lengths? Thank you, -Jerome Jerome Nwachukwu
Re: [ccp4bb] HisTrap Trap
Bernhard, We have had similar, but not identical issues with some insect cell media, as well as column interference by lipid. If we see this, we tend to run all the material with the protein (cell lysate or media) through step-elution ion exchange (quick on, quick off). While the purification is minimal, the protein is more concentrated and performs better on Ni-columns. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 19, 2014, at 10:13 AM, Bernhard Rupp hofkristall...@gmail.com wrote: Hi Fellows, my lab mates successfully expressed a glycoprotein in CHO cells in serum free medium, and the protein captures nicely on HisTrap Excel 1ml columns (obviously, high yield is not my problem…). We load ca 1L supernatant at 0.5 ml/min, and eluate with a steep imidazole gradient. 20mM Imidazole buffer for regeneration. Works fine (and often…see yield remark). Overcome by common crystallographers’ greed (nor creed), we switched to stable xfected HEK293, and cell free medium Gibco CD 293. The first run gave high final yields cheers. The second run less of either, because the small HisTrap column essentially dissolved – the medium collapsed, Ni leaches out, kaput as kaput goes. A 3rd run on a similar previously working column lead to the same result. Only thing changed was the cells and medium. Same buffers, same gradients, same Akta equipment, same lab techs. Before I improve the statistics by ruining further columns, has anybody experienced such a calamity that might be blamable on secret media components or similar? There is a mysterious ‘proprietary dispersant’ preventing cell adhesion quoted…. Best wishes, BR Bernhard Rupp b...@ruppweb.org b...@hofkristallamt.org http://www.ruppweb.org/ ---
Re: [ccp4bb] HisTrap Trap
Bernhard, One other point, after rereading your email. For step-elution ion exchange (quick on, quick off), you can use some very cheap and quite robust media the will resist a lot of stuff. Have them pour their own columns and toss it away when it gets too dirty. You don't have to use the high-end stuff for this step (CM-sephedex, Dowex, CM-cellulose, etc.). Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 19, 2014, at 10:13 AM, Bernhard Rupp hofkristall...@gmail.com wrote: Hi Fellows, my lab mates successfully expressed a glycoprotein in CHO cells in serum free medium, and the protein captures nicely on HisTrap Excel 1ml columns (obviously, high yield is not my problem…). We load ca 1L supernatant at 0.5 ml/min, and eluate with a steep imidazole gradient. 20mM Imidazole buffer for regeneration. Works fine (and often…see yield remark). Overcome by common crystallographers’ greed (nor creed), we switched to stable xfected HEK293, and cell free medium Gibco CD 293. The first run gave high final yields cheers. The second run less of either, because the small HisTrap column essentially dissolved – the medium collapsed, Ni leaches out, kaput as kaput goes. A 3rd run on a similar previously working column lead to the same result. Only thing changed was the cells and medium. Same buffers, same gradients, same Akta equipment, same lab techs. Before I improve the statistics by ruining further columns, has anybody experienced such a calamity that might be blamable on secret media components or similar? There is a mysterious ‘proprietary dispersant’ preventing cell adhesion quoted…. Best wishes, BR Bernhard Rupp b...@ruppweb.org b...@hofkristallamt.org http://www.ruppweb.org/ ---
Re: [ccp4bb] Issue with Molecules per Asymmetric Unit for Molecular Replacement
Matt, In addition to the suggestions of the others, have you done a simple self rotation function? It can tell you quite a bit about how things are packed and give you strict criteria for choosing one solution over another. As Roger said, choosing an even number of monomers in the ASU is a good strategy, particularly if the self rotation function shows NCS 2-folds. Also, a calculated Matthews coefficient is NEVER correct, it is a mere guideline; it only has validity for any particularly crystal form AFTER the fact. Let the number of monomers in the ASU vary from 6-10; I have had MR cases that have had as little as 40% solvent to 70% solvent, where the calculated Matthews coefficient was quite wrong (i.e., the most common value observed in OTHER crystals). Two things to watch out for are: (1) An odd number of monomers in the ASU. I have had 1 1/2 dimers in an ASU (the 1/2 dimer is paired with another in a neighboring ASU). It is sometimes confusing to people and occasionally difficult to solve with some MR programs due to clashes. (2) Translation symmetry, which still can confuse some programs (but they are are getting better at detecting it). Finally, as Herman pointed out, look at the packing of any solution you are considering. It is surprising how a correct solution looks correct: nice intermolecular contacts and a pleasing distribution of mass throughout the unit cell (meaning expand out to at least a unit cell volume, which is easy in Pymol). Any unexplained gaps (meaning not caused by a missing domain) should be viewed critically. Regards and Good Luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 15, 2014, at 6:50 PM, Matthew Bratkowski mab...@cornell.edu wrote: Hello all, I am working on the structure of a small protein in space group P212121. The protein is monomeric in solution based on gel filtration analysis. The Matthews Coefficeint program indicates that 9-10 molecules per asymmetric unit results in ~50% solvent content, while 1 molecule per asymmetric unit results in ~95% solvent. I tried molecular replacement with a search model which is essentially identical in sequence to my protein, and searched for 9 or 10 molecules/asu. Using MolRep with 9 or 10 molecules/asu, I get poor contrast scores around 1-1.5. However, when using Phaser, I get a solution with one molecules/asu. Likewise, when I went back and tried MolRep with 1 molecule/asu, I got a contrast score of 3.12. This model still has some issues, but looks more correct compaired to models created with 9 or 10 molecules/asu. It seems highly unlikely that a crystal would contain 95% solvent, but is there any possiblility that this could be the case? Assuming that the Matthews coefficient is correct, does anyone have an idea why MR seems to work better for 1 molecule/asu with 95% solvent content compared to 9-10 molecules with 50% solvent content? Alternatively, is there any reason why the Matthews coefficient could be calculating incorrectly? Any suggestions would be helpful. Thanks, Matt
[ccp4bb] Helix alignment and movement
Dear Eugene and other CCP4ers The recent discussion about superposition has prompted me to ask about a different kind of superposition problem. We are working on a small dimeric protein that is entirely made up of helices. Instead of large, concerted domain movements, which I am quite familiar with, we have 3 structures for the dimer that display slightly, but significantly different helical conformations. I have been able to find the minimal substructure that allows the best superposition (lowest RSMD and maximum number of aligned/superimposed C-alphas). The problem is that none of the superposition programs available outputs a list of residue by residue deviations OUTSIDE of the alignment set (for good reason as there may not be 1 to 1 correspondence outside of this set). I can do some of this in a piecemeal fashion with Moleman2, but not everything I want. My question is, after creating an optimal structural alignment, are there newer programs that: (1) Create a list of residue by residue deviations over different subsets of the structure, particularly OUTSIDE of an alignment set? (2) Localize and measure movement of secondary structure (helix tilt or bending)? I can't seem to find what I need, but I may not be searching with the right key words. Thanks, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 11, 2014, at 7:14 AM, Eugene Krissinel eugene.krissi...@stfc.ac.uk wrote: My guess is that only atom pairs that are superposed to some measure of distance between them, are output. Can't say that I checked lsqkab code this weekend, but documentation does not suggest anything like that. Is this a problem for you? note that you can use other aligners/superposers in CCP4, ssm or Gesamt which will output all coordinates. Eugene
Re: [ccp4bb] maltose binding protein
Rana, It is hard to answer you question without more details (MW and pI of your target protein). MBP binds very well to amylose resins and is usually quite easily bound to anion exchange resins. Did you just run a standard ion exchange protocol or try different pH regimes? However, you did mention you have used a detergent. Why do you do that? MBP binding to amylose resins can be markedly disturbed in the presence of several different detergents, which is a particularly bad thing for a membrane protein fusion. That is why all our MBP fusion constructs have an additional His-tag. If you really don't need the detergent, leave it out, then try the amylose resin again. One other point is how old is your amylose resin and, if you express in E. coli, do you regularly add a little glucose to the media? Amylose resin is degraded by E. coli amylases, which I believe is suppressed when glucose is in the medium. However, inferring from your email, I would suppose that you initially purified the MBP fusion using the amylose resin. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Mar 27, 2014, at 6:49 AM, rana ibd rna19792...@yahoo.com wrote: Dear Mark Thank you for yor reply, and yes I have tried adding it to the maltose resin after cleavage but the MBP runs through with my protein, I have also tried 1M NaCl but with no luck and I also apply detergent after cleavage to the dialysis buffer because I usually dialyze after cleavage , is there anything that could maybe precipitate the MBP Best Regards Rana From: Mark J van Raaij mjvanra...@cnb.csic.es To: rana ibd rna19792...@yahoo.com Sent: Thursday, March 27, 2014 11:35 AM Subject: Re: [ccp4bb] maltose binding protein did you try the maltose-resin? in principle it should bind MPB but not your protein. You can try to add salt or detergents to disturb interaction between MBP and your protein (also in gel filtration). No guarantee of success, unfortunately not all protein behave nicely. Mark J van Raaij Lab 20B Dpto de Estructura de Macromoleculas Centro Nacional de Biotecnologia - CSIC c/Darwin 3 E-28049 Madrid, Spain tel. (+34) 91 585 4616 http://www.cnb.csic.es/~mjvanraaij On 27 Mar 2014, at 11:26, rana ibd wrote: Dear CCP4 Does anyone know how to remove the maltose binding protein after cleavage from the target protein; I have tried gel filtration and ion exchange but with no luck, my protein is interacting with the MBP even after complete cleavage. I would be grateful for any help or suggestions Best Regards Rana
Re: [ccp4bb] off-topic: protein losing FAD during purification
Stefano, Before you address the problem, you need to ask yourself a couple of things. You say that on the gel filtration we clearly see two bands corresponding to holoprotein and free FAD. That is not too odd, but have you ask the question is all the protein good protein. Is this an enzyme? If so, assay samples before and after the gel filtration column without adding extra FAD and compare to assays with a slight excess FAD. If the specific activities are different (slight excess FAD before gel filtration no added FAD after the gel filtration no added FAD), I would follow the good advice of the others. You have just removed exchangeable FAD and are crystallizing a mixed system. Is this a recombinant protein partially purified using a His-tag or such? If the specific activities are the same, I would suspect that not all the protein is in good shape. Try ion exchange chromatography to see if two different protein variants can be separated. With luck that can be (1) active holoprotein and poorly-folded protein (always a problem with His-tag purified recombinant protein) or (2) active holoprotein and apo-protein. If it is not an enzyme, or it is not an easy assay, take the spectrum of the gel filtration-purified holoprotein and see if it differs from free FAD, then add FAD to a near stoichiometric level to see if you can saturate without a spectral change (e.g., a blue-shift of the absorption peak). That could tell you if it is an active holoprotein and apo-protein problem or active holoprotein and poorly-folded protein problem. Sometimes poorly-folded protein weakly binds the cofactors, but not in the native way. If you are lucky, supplementing with FAD, either before, during, or after purification will work fine. In some cases, I have found that it is better to purify the apo-protein, then reconstitute the holo-protein, if the former is stable enough. However, some heme, NAD(P)(H), and FAD binding protein allow removal of the cofactors but not its reconstitution. It depends are your protein system. A little more biochemistry is needed, but find a nice assay system to verify what you are doing. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Mar 13, 2014, at 6:40 PM, Benini Stefano (P) stefano.ben...@unibz.it wrote: Dear All (those dealing with wetlab stuff..), While purifying a FAD containing protein we lose part of the FAD (on the gel filtration we clearly see two bands corresponding to holoprotein and free FAD). We obtain crystals but diffracting to only about 4 A despite their beautiful look. Our hypothesis is that the crystals contain a population of molecules with and without FAD (?). The questions are: 1) how to keep FAD bound to the protein during purification and crystallization? 2) how to completely remove FAD from the protein? Thank you very much for any help provided! Best regards Stefano (part-time wetlab person) Dr Stefano Benini, Ph.D. Assistant Professor First International workshop: Molecular Basis of Fire Blight, Bolzano 15.10.2014 Laboratory homepage: http://pro.unibz.it/staff2/sbenini/B2Cl.htm Personal homepage http://pro.unibz.it/staff2/sbenini/ I don't like anything that's fake and I hate pretenders! * Bioorganic chemistry and Bio-Crystallography laboratory (B2Cl) Faculty of Science and Technology Free University of Bolzano Piazza Università, 5 39100 Bolzano, Italy Office (room K2.14): +39 0471 017128 Laboratory (room E.021): +39 0471 017910 Fax: +39 0471 017009 ogni giorno in più è un giorno in meno.
Re: [ccp4bb] Determining concentration of membrane protein
Roger, While I agree with your list, the BCA assay does not use molybdate (as we make it from scratch with bicinchoninic acid, sodium carbonate, sodium bicarbonate, sodium tartrate, and cupric sulfate pentahydrate). For membrane proteins, I prefer the BCA assay until the protein is pure enough to use A280. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Feb 13, 2014, at 10:39 AM, Roger Rowlett rrowl...@colgate.edu wrote: Your basic choices for protein assays are: Alkaline copper methods (e.g., Biuret and micro-biuret) alkaline copper + molybdate methods (e.g., Lowry, BCA assays) Hydrophobic dye methods (e.g. Bradford) UV methods (e.g., A280, A230, A210, etc.) Method 1 is least sensitive to amino acid composition, but is also has highest detection limits. Thiols interfere. Method 2 is very idiosyncratic with amino acid composition, and also subject to interference by thiols. Method 3 is not usable in detergent solutions. Method 4 has many inteferences as most everything absorbs in the far UV region. If you have some special protein cofactors, metals, chromophores, etc. these can be exploited for better measurements. For ecample metalloproteins are easy to quantify by ICP-OES or TXRF if they are reasonably pure. Cheers, ___ Roger S. Rowlett Gordon Dorothy Kline Professor Department of Chemistry Colgate University 13 Oak Drive Hamilton, NY 13346 tel: (315)-228-7245 ofc: (315)-228-7395 fax: (315)-228-7935 email: rrowl...@colgate.edu On 2/13/2014 10:06 AM, Raji Edayathumangalam wrote: Dear CC4BBers, I am trying to figure out what is the best way to determine the protein concentration of my membrane protein. My purified membrane protein is in 20mM Tris pH 7, 150mM NaCl and 0.02% DDM (CMC of DDM=0.0076%). After reading the friendly manuals and searching online, I've learned that detergents interferes with assays like Bradford but can't find good descriptions of what works best. For now, I am trying to estimate concentration from absorbance at 280nm and using molar extinction coefficients based on aromatic amino acids, but again suspect detergent interference. I would like to know what other folks working on membrane proteins are doing. Thanks very much. Raji -- Raji Edayathumangalam Instructor in Neurology, Harvard Medical School Research Associate, Brigham and Women's Hospital Visiting Research Scholar, Brandeis University
Re: [ccp4bb] largest protein crystal ever grown?
Felix, Although I have passed through George's lab at UCSD several times, I have not seen that crystal, but it should be pointed out that George's interests did extend beyond photosynthesis. Some of us older folks remember George's contributions to protein crystal growth, both theoretical and experimental (see Z. Kam, H.B. Shore, and G. Feher, J. Mol. Biol. 123, 539, 1978 or his contribution in Methods in Enzymology, volume 114). So it would not be surprising to find a humungous crystal in his lab; he was interested in the mechanism(s) of crystal growth cessation, which may have led his group to see how big a crystal they could grow. Unfortunately, parts of his lab were often dark when I was visiting due to his photosynthesis experiments. Perhaps Jim Allen or former people from the Kraut and Xuong labs would know. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 25, 2013, at 11:13 AM, Felix Frolow wrote: I guess that this crystal was never tested with any X-ray source. After all George is a physicist who study photosynthesis processes by spectroscopic methods. However (unrelated but connected) I have collected once a data set from a see urchin needle which was 1 cm long, about 3 mm across (protein mass was dissolved), it was a single crystal despite a complicated and beautiful architecture, and mosaicity was about 0.5 deg on a Rigaku AFC5 diffractometer (mounted on a rotating anode with Ni filter. So I would not bet on the large crystal - big mosaicity formula. FF This remarkable hollow Dr Felix Frolow Professor of Structural Biology and Biotechnology, Department of Molecular Microbiology and Biotechnology Tel Aviv University 69978, Israel Acta Crystallographica F, co-editor e-mail: mbfro...@post.tau.ac.il Tel: ++972-3640-8723 Fax: ++972-3640-9407 Cellular: 0547 459 608 On Oct 25, 2013, at 16:54 , Derek Logan derek.lo...@biochemistry.lu.se wrote: Hi Felix, What was the mosaicity of this crystal? The absorption correction must have been challenging too... Derek On 25 Oct 2013, at 13:23, Felix Frolow mbfro...@post.tau.ac.il wrote: Well if we start recalling rumours, I have heard that in UC San Diego in the laboratory of George Feher there was (is) a tetragonal hen egg white lysozyme crystal which weighted between 0.5 - 1.0 kg. It grew suspend on a mountain boots shoelace of the read colour. I have never visited George laboratory, but maybe among the society there are some who can shed some light on that…. FF Dr Felix Frolow Professor of Structural Biology and Biotechnology, Department of Molecular Microbiology and Biotechnology Tel Aviv University 69978, Israel Acta Crystallographica F, co-editor e-mail: mbfro...@post.tau.ac.il Tel: ++972-3640-8723 Fax: ++972-3640-9407 Cellular: 0547 459 608 On Oct 25, 2013, at 12:18 , Boaz Shaanan bshaa...@bgu.ac.il wrote: Hi, Referring to the Hb crystal that Bill Scott saw in the MRC crystal growing room (by now tho old one I guess), is that the one that was sitting in the largest part of the Pasteur pipette? I recall this one and I keep telling my students about it when they ask about crystal size limits. Cheers, Boaz הודעה מקורית מאת: simon.phill...@rc-harwell.ac.uk תאריך: אל: CCP4BB@JISCMAIL.AC.UK נושא: Re: [ccp4bb] largest protein crystal ever grown? Hi Derek, That brings back memories. I am pretty certain that is the myoglobin crystal that was already on Benno's shelf at Brookhaven when I went there in 1980 to collect my oxymyoglobin neutron data. It would the metmyoglobin crystal Benno got the early neutron data from. He just kept it on the shelf because there was, of course, no degradation in the beam and a crystal is a pretty stable way to store a protein. Whenever he wanted more data he took it off the shelf and put it back on the beamline. If Benno is reading this bulletin board I am sure he could tell us more. Simon Simon E.V. Phillips Director, Research Complex at Harwell (RCaH) Rutherford Appleton Laboratory Harwell Oxford Didcot Oxon OX11 0FA United Kingdom Email: susan.jo...@rc-harwell.ac.uk Direct email: simon.phill...@rc-harwell.ac.uk Tel: +44 (0)1235 567701 (direct) +44 (0)1235 567700 (sec) +44 (0)7884 436011 (mobile) www: www.rc-harwell.ac.uk -Original Message- From: CCP4 bulletin board [mailto:CCP4BB@JISCMAIL.AC.UK] On Behalf Of Derek Logan Sent: 24 October 2013 19:08 To: ccp4bb Subject: Re: [ccp4bb] largest protein crystal ever grown? Hi,
Re: [ccp4bb] Staining Crystals with comassie
There are many caveats to using glutaraldehyde on crystals, either for fixing crystals or for staining them. First, I would not hang a 1-2uL drop of 25% glutaraldehyde in the vapour diffusion chamber, but add enough glutaraldehyde into the reservoir to make it 0.5-1.0 % (a 1:25 or 1:50 dilution; a 1% solution is 100 mM). Not only will it be just as effective, the reservoir becomes the control: if the reservoir turns yellow, you have free amines in the system (Tris, ammonium, etc.). Second, the yellow color, which is due to Schiff's base formation, is harder to see in warm light (color temperature, not the temperature of the stage) when you are looking at small or thin crystals. Use cool, white lights (like LEDs). Finally, keeps some buffer around that is suitable for solubilizing the protein. If you are not sure about the color change, just add 10 uL of buffer to the crystals and watch if they dissolve. If they don't when treated with glutaraldehyde, they are protein crystals. As Zhijie said, be careful with handling glutaraldehyde. It is highly volatile and dangerous. If you smell it (and the sweet smell will be obvious), it is fixing you. Keep a waste bottle half-full with 1 M ammonium sulfate, not glycine (too expensive), then just dump any glutaraldehyde waste into it. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 16, 2013, at 6:45 AM, Zhijie Li wrote: Hi Danilo and all, A little trick for the glutaraldehyde staining: you can hang a 1-2uL drop of 25% glutaraldehyde (or the most concentrated stock solution you can find) besides your crystal drop in the vapour diffusion chamber. The glutaraldehyde will get into the crystal drop via vapour diffusion. The color will normally show within 2hrs and become very intense overnight. It is also a gentle way of crosslinking the crystals (http://scripts.iucr.org/cgi-bin/paper?wb0066, and http://hamptonresearch.com/tip_detail.aspx?id=74, ). Care should be taken when handling aldehyde concentrates: do not breathe it, and do not let the vapour get in touch with your eyes. Waste can be inactivated by concentrated glycine solution. BTW, the acetic acid in the coommasie blue solution seems unnecessary in a crystal staining solution. The solution recipe seems to be taken from a gel staining solution. When staining polyacrylamide gels, the acid (oringinally HCl) is supposed to denature the proteins so that they do not diffuse in the gel. The MeOH is for solubilizing the commonly used coommassie R250. (Another thing: I strongly suggest to substitute the MeOH in PAGE staining and de-staining solutions with EtOH. EtOH works perfectly fine, without MeOH's poisonous effect on human. Our staining solution contains 20% EtOH and 20%HAc.) For staining crystals, we do not need to add the acetic acid. Also coommassie G250 is more soluble in water than the R250 by having methyl groups instead of ethyl groups. 0.5% coommassie G250 can be readily made in DMSO or 95% EtOH. Then this stock solution can be diluted with water or the mother liquor 10x-100x for the staining. Many crystals can tolerate up to 10% DMSO. Zhijie -Original Message- From: Danilo Belviso Sent: Wednesday, October 16, 2013 3:53 AM To: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] Staining Crystals with comassie Dear All, izit dye is a solution containing methylene blue that you could prepare in your lab. I usually prepare a solution of 0.05%w/v of dye in water and then I add a volume of dye solution equals to 10% of the volume of the drop containing the crystal to test. I prefer to add the dye solution in small portions (if the volume permits) every 2-3h in order to limit the shock due to the new solution on the crystal. You should remember that this test is not definitive: the dye is a cationic dye, that needs of anion counter part to bind the protein. Therefore, the dye is not able to colour all protein crystals: in addition, colouration is affect by pH of the crystallization condition, since low pH could increase the positive charge on the protein reducing its ability to bind the dye. You could try also glutaraldehyde as alternative. In order to perform this test, you should put the crystal into a low ionic strength buffered solution containing up to 2% glutaraldehyde. In this condition, formation of Schiff bases with the lysines and N-term residues occurs and the crystal become a yellow gel, while salt crystals dissolve and should not be coloured. To perform comassie crystal staining you should prepare a solution
Re: [ccp4bb] limited proteolysis / ms
Gloria, Here is how we are now doing it, courtesy of Yu-Jing's and Merlin's fine development work. We have also tried it to quantitatively remove tags by TEV cleavage, but we haven't solved that problem yet. Regards, Michael Tan, Yu-Jing, Wei-Han Wang, Yi Zheng, Jinlan Dong, Giovanni Stefano, Federica Brandizzi, R. Michael Garavito, Gavin E. Reid, and Merlin L. Bruening (2012) Limited Proteolysis Via Millisecond Digestions in Protease-Modified Membranes. Analytical Chemistry 84(19), 8357-63 R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 8, 2013, at 3:34 PM, Gloria Borgstahl wrote: We are trying to apply this approach for the first time to define a crystallizable domain of our protein of interest. Can anyone send a protocol that includes exactly how to do the mass spectrometry measurement? Our core lab here doesn't know and I'm just not that gifted. Happy October, G
Re: [ccp4bb] Identity of a Bacterial lipid
Dear Andre, It always impressive to see a near atomic resolution structure with a bound lipid. Congratulations. However, to identify what lipid you have requires a bit more analysis than just looking in databases. First, what is the bacterium you are using as the host? If it is E. coli, then the known lipids are very well characterized. Also, VERY FEW E. coli lipids have sites of unsaturation, and virtually polyunsaturated fatty acids (PUFAs) have one sp3 carbon in between the double bonds (arising from the mechanisms of biosynthesis). So your proposed structure doesn't seem right from a biological sense, which makes looking into databases unproductive. However, since you solved the structure, do your produce enough protein to isolate the putative lipid by simple TLC? With the help of a good lipid lab you should be able to tell what it with greater certainty. Try to get a copy of Techniques of Lipidology by Morris Kates from Elsevier. Although it is old school stuff, it will help you isolate enough for mass spec and NMR analysis. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 3, 2013, at 8:42 AM, Andre Luis Berteli Ambrosio wrote: Dear colleagues, We have just determined the crystal structure (at 1.1 A max resolution) of a recombinant protein that crystallized in complex with a leftover bacterial lipid, the full identity of which we are currently struggling to identify. Please see attached (3 views of the molecule). The map strongly suggests an 18-carbon long polyunsaturated fatty acid, with 5 conjugated unsaturations (at positions 5, 7, 9, 11 and 13, all cis), covalently bound to some extra chemical group at is polar head. This extra group seems to be comprised of 4 (5?) atoms, though I am afraid cannot tell if it extends further into not-so-well-structured atoms. Myself and a student have spent the last two days searching on the web for possible matches for this ligand without any success. For instance, we have generated a SMILES formula for the aliphatic tail comprising the unsaturations and browsed for similar compounds at PubChem with different similarity cutoffs, but nothing seemed to resemble the complete molecule. We would appreciate if you could make any comments on the nature of this ligand or perhaps suggest your favorite computational tools. We will perform mass spec on it soon. Thank you beforehand. Andre LB Ambrosio, DSc Laboratório Nacional de Biociências - LNBio CNPEM, Brazil FA-density.png
Re: [ccp4bb] membrane protein and phase separation
Pascal, The reason this phenomenon looks odd to you is that detergent phase separation is not a micelle size phenomenon, but a micelle surface phenomenon. Like any colloidal solution, even protein, there can be conditions where the colloidal particles aggregate (the cloud point), creating a mixture of solubilized colloidal particle L' and a second phase of aggregated particles L''. The second phase of aggregated particles (which can appear to be a viscous liquid) can be micelles, PDCs, or simply protein when working with only soluble proteins; for the latter two, this can lead to precipitation or crystals. However, at the concentrations of detergent/lipid we typically use, we tend to see only aggregates of micelles or PDCs in the second phase, rather than the other surfactant phases (bicubic, lamellar, hexagonal, etc.). Nor is the second phase formation an indication of micelle fusion (which was a hot topic of discussion in the 1980's). The physical conditions that cause phase separation are the same as for crystallization (the solvent environment variables of pH, salt, polymer solutes, etc.). It can also be a unique phenomenon of the PDCs you make with your protein and the chosen detergent. One quick experiment you can try is to add glutaraldehyde to the reservoir and fix a drop with phase separation (glutaraldehyde is quite volitile). CAVEAT: you can't do this in the presence of Tris, free amine solutes like ammonium sulfate, or lipids/detergents with a free amine. If the droplets are fixed into a gel, then the predominant species in the second phase is not pure detergent micelles, but your PDCs at very high concentration (50 mg/mL). Hence, your PDCs want to phase out, which is not always a bad thing. The trick then is to encourage crystallize. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 1, 2013, at 6:23 PM, Pascal Egea wrote: Dear All, I have a question tailored for the membrane protein and detergent folks. We are purifying a membrane protein that associates into an homoligomeric pore and we have been successfully preparing it in two detergents: FC-12 or a mild lipid. The two Protein Detergent Complexes look very homogenous by SEC and can be concentrated without protein loss on membranes with MW cutoffs (100kDal) way larger that the expected their respective free detergent micelles. Everything looks good so far... until we get to the crystallization stage. While the PDC in FC12 does not tend to form too much phase separation, the PDC in the lipid does. This looks a bit odd to me since these lipid micelles are supposed to be a bit smaller than the FC-12 micelles. We are working at twice the CMC and besides lowering the detergent concentration, I am a bit perplex about what I am observing. Intuitively I would have expected to observe the reverse behavior: worst in FC-12 than the lipid. This lipid is a very mild solubilizing/reconstituting agent that has already been successfully used for structure determination. Any advice or thoughts will be greatly appreciated. Is this something that some of you have already observed? Many thanks in advance, -- Pascal F. Egea, PhD Assistant Professor UCLA, David Geffen School of Medicine Department of Biological Chemistry Boyer Hall room 356 611 Charles E Young Drive East Los Angeles CA 90095 office (310)-983-3515 lab (310)-983-3516 email pe...@mednet.ucla.edu
Re: [ccp4bb] Dose anyone see this ligand before?
Wei I heartily second Dale's comment. Since you do know what has been in contact with your protein, you should be able to make a list of ALL POSSIBLE compounds your protein has been exposed to. However, don't go off the deep end. You say that We guess that the molecular formula should be C8H18O2. So we search this formula in google and find two candidate molecules, but how did you determine C8H18O2 when you can't see hydrogen, and guessing the elemental composition of a ligand may be possible at high resolution IF you know your occupancy. Two observations I have are: 1. Anything you exposed your protein to could bind to it tightly regardless of the number of purification steps the protein went through. But as Dale said, the ligand is most likely biologically derived. 2. As an example of the exception proves the rule circumstance, I believe it was the monoamine oxidase B structure that had one of the plasticizers from the plastic trays bound in the active site. Something to think about as we drink from our water bottles and mull about neurochemistry. Nonetheless, it was not a wild, off-the-shelf ligand, but some thing the protein had been in contact with. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 17, 2013, at 2:20 PM, Dale Tronrud wrote: Do you have any reason to expect either of these molecules would be in your crystal? The model you build has to fit the density, be consistent with the surrounding environment (which you haven't shared with us) and you have to have some story for how that molecule got in your crystal. Personally I would steer away from industrial compounds and focus more on biological molecules and common additives used in purification and crystallization. The environment is critical to identifying this molecule. What hydrogen bonds does this molecule make? What charges are near by? Certainly the presence or absence of hydrogen bonds will distinguish between these two compounds before you go to the trouble to build a model of either. Dale Tronrud On 7/17/2013 6:35 AM, Wei Feng wrote: Dear all, Thank you for your advices. I had tried to use MPD and pyrophosphate etc to fix the density map but all of them were too small. We guess that the molecular formula should be C8H18O2. So we search this formula in google and find two candidate molecules 1: http://flyingexport.en.ecplaza.net/dhad-99-5--137042-689140.html 2: http://en.m.wikipedia.org/wiki/Di-tert-butyl_peroxide Could you tell me how to get the coordinate of these molecules? Thank you for your time! Wei
Re: [ccp4bb] Concentrating purified membrane protein
Toufic, That is my point. Most non-surfactant chemists (i.e., us) seem to feel that detergents have a fairly homogeneous micelle size (DDM micelle ~65-70 kDa), which is not true. You are talking about average micellar size, not the distribution. Micelles readily and rapidly disaggregate into free micelles, into partial micelles, they deform, or they cluster into aggregates, all in a manner that is dependent on the solute conditions. For some detergents (but not all), their effective CMC can change with local concentrations (as can occur at high salt or protein concentrations or with high lipid content). The real point is defining EXCESS free detergent, not the total concentration or the amount bound to the protein. There is no assay that determines protein-free detergent concentration, only the total detergent concentration. A 100 KDa protein at 10 mg/mL is at 0.1 mM. Given your examples, the effective concentration FOR ONLY the protein-bound detergent is 40-100 times that. For DDM, the protein-bound detergent concentration is already ~23 -110 times its CMC, and for DM, that can be ~2.3 - 11 times its CMC. Now, we have to consider that this protein-bound detergent is in rapid equilibrium with an unknown amount of free detergent in both micellar and monomer forms. At the moment, we cannot rapidly determine all the parameters needed to find an analytical method to the problem of controlling precisely the detergent concentration, so we are stuck with an empirical one (try different methods till one works, i.e. you reproducibly get crystals) and off-the-cuff calculations (like above). But the real take-home message is that crystals WERE NEVER GROWN in drops containing 0.1 or 0.05% detergent, but 0.1 or 0.05% detergent PLUS whatever is bound to the protein. For some cases, that will be 2-5X the nominal detergent concentration. Finding an optimal detergent environment for membrane protein crystal growth depends on finding the sweet spot in this complex detergent equilibrium. This means distinguishing between the nominal detergent concentration (what you add to your buffers) and what you actually need to have in your protein solution to allow crystal growth. I think that much of my gray hair came from dealing with these %*($# detergents. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 15, 2013, at 10:52 AM, El Arnaout, Toufic wrote: Hi, That is true, Michael.. can't remember the reference, but in some study they found that even if you use a higher MWCO (for example 100 kDa) with DDM (micelle 65-70 kDa), there is still 70-80 % retained detergent in the protein sample. Raji, another thing I would like to say is that the right MW cut-off may depend on the protein, the detergent micelle, but also the bound detergent molecules per protein. It could be 40-50 molecules of DM let's say for protein A, but 80-90 molecules for protein B. You can quickly evaluate the concentration step for your protein using SDS-PAGE or UV abs, and for the detergent there are many methods like colorimetric assays, TLC.. Best wishes t toufic el arnaout School of Medicine - 660 S Euclid Ave Washington University in St. Louis St Louis, MO 63110, USA From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of R. M. Garavito [rmgarav...@gmail.com] Sent: Sunday, July 14, 2013 11:17 AM To: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] Concentrating purified membrane protein Raji, One point the most people forget about is that whenever you concentrate any detergent-solubilized membrane protein is that you will ALWAYS concentrate the detergent. So regardless of the MWCO, if the protein-detergent complex concentrates, the overall detergent concentration also increases. What you want to shoot for is balancing protein loss with obtaining a sample having minimal EXCESS free detergent. While a concentration step with a Ni-column followed by dialysis will work, so will a wise choice of concentrator. One trick to moderate high excess detergent is just to avoid the need for a many fold concentration where you will really concentrate the free detergent. Nonetheless, we have used concentrators effectively, and a Ni-column followed by dialysis as well, but if you still have too much free detergent, you can always use a spin desalting column with G-25 sephedex to bring remove detergent down to a nominal concentration. In all cases, you will lose some protein. Good luck, Michael R. Michael Garavito
Re: [ccp4bb] Concentrating purified membrane protein
Raji, One point the most people forget about is that whenever you concentrate any detergent-solubilized membrane protein is that you will ALWAYS concentrate the detergent. So regardless of the MWCO, if the protein-detergent complex concentrates, the overall detergent concentration also increases. What you want to shoot for is balancing protein loss with obtaining a sample having minimal EXCESS free detergent. While a concentration step with a Ni-column followed by dialysis will work, so will a wise choice of concentrator. One trick to moderate high excess detergent is just to avoid the need for a many fold concentration where you will really concentrate the free detergent. Nonetheless, we have used concentrators effectively, and a Ni-column followed by dialysis as well, but if you still have too much free detergent, you can always use a spin desalting column with G-25 sephedex to bring remove detergent down to a nominal concentration. In all cases, you will lose some protein. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 14, 2013, at 10:29 AM, Raji Edayathumangalam wrote: Thanks everyone for your responses. I definitely plan to save the flowthrough so we'll see what happens. My protein has a His tag and I did consider doing an affinity step for concentration except I do not want to have imidazole for some functional assays that I need to carry out with the protein. Just occurred to me that I could simply dialyze out the imidazole after the affinity step. Thanks again! Raji On Sat, Jul 13, 2013 at 8:47 PM, Raji Edayathumangalam r...@brandeis.edu wrote: Hi Folks, Sorry for the non-ccp4 post. I have purified an 18kDa membrane protein and want to concentrate the protein from gel filtration fractions, which are in buffer containing 0.05% DDM (well above the CMC for DDM). My colleague was able to concentrate a 30kDa membrane protein using a 100kDa MWCO concentrator but I am not sure if I can do the same without losing protein in the flowthrough. On the other hand, if use too low a MWCO for the concentrator, then I'm concerned that I may end up concentrating the DDM and end up with too much detergent in the final sample. Any tips about how to concentrate my low MW protein without concentrating the DDM? Many thanks. Raji -- Raji Edayathumangalam Instructor in Neurology, Harvard Medical School Research Associate, Brigham and Women's Hospital Visiting Research Scholar, Brandeis University -- Raji Edayathumangalam Instructor in Neurology, Harvard Medical School Research Associate, Brigham and Women's Hospital Visiting Research Scholar, Brandeis University
Re: [ccp4bb] Harmful effect of X-ray
Most modern textbooks have sections on the proper protections and measures to take, although the information may be dated. See chapter 6 in Volume III of the International Tables for X-ray Crystallography. With the modern equipment and regulatory measures in most countries, you really have to work hard to be exposed at dangerous levels (which can lead to skin lesions and burns). However, you can get yourself exposed if you intentionally circumvent the safety measures and interlocks. In my experience, X-ray exposure in crystallography labs is very low and not dangerous. Our radiation safety people find our labs to be very clean with respect to scattered radiation around the sample compared to medical X-ray labs. Talk to your institute's safety people for advice. For in-house equipment, you are most at risk of exposure during aligning the equipment. If you talk to the old crystallographers (or their students who are now +50 year old), you might hear stories of aligning collimators and cameras by the tingle on your eye as you look into the beam. By the time protein crystallography came around (50s-60s), phosphors and film were used for alignment so the danger comes mostly from scattered radiation and poor shielding. In all the years I have worked with X-rays without protection (I only wore a lab coat to prevent film developer from staining my clothes), neither I nor my colleagues have ever had X-ray exposures above background as determined by film badges and ring badges. In fact, we once exposed a film badge intentionally to see if anyone cared. We caught hell for doing that. For synchrotron sources, chances of being exposed as a general user are now nil unless you really work hard to subvert the safety measures (which will get you kicked out). Hope this helps, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 12, 2013, at 11:14 AM, diptimayee mishra wrote: Dear All, Can anyone please tell me regarding the harmful effects of X-ray , we are using for protein crystallography, on human being and what are the precautions we should take. Thanks
Re: [ccp4bb] Heterogeneity during purification
Dear Teresa, In addition to Bert's excellent list, I have found that many membrane proteins aggregate in regular SDS-PAGE sample buffer, particularly when heated. In the best case, we always see dimers or, in the worst case, a ladder or smear of aggregates. If you heat your SDS-PAGE samples, try not heating them. Secondly, your observation that you have protein eluting near the void volume on Superdex 200 may also suggests that you have either incomplete solubilization (i.e., big lipid-protein complexes) or, as Bert remarks, an unhappy protein that aggregates in the detergent you have chosen. However, before you consider this a problem, ask yourself if this is an expected result due to the heterologous overexpression of a recombinant protein. Unlike purifying a native protein, overexpression of a recombinant protein can led to a considerable amount of misfolded or misassembled protein mixed with good protein. As you might be overloading the membrane protein insertion system, you may need to dial back the induction. Finally, were you measuring the amount of high order aggregates in SEC by A280 or by another means? The proportion of high order aggregates observed near the void volume is always skewed due to light-scattering of the much larger particles. Because of this, a large void volume peak (~7.5-8 mL on a Superdex 200 10/300 column) may contain a lot less protein than you think. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 9, 2013, at 4:47 AM, Bert Van-Den-Berg wrote: since your protein aggregates even in a mild detergent you may have to find an ortholog that is more stable. however, there are a few things you can try before moving on (in arbitrary order): 1. add glycerol during purification (5-20%) 2. get rid of the imidazole as fast as possible after Ni. Many proteins do not like imidazole at all. 3. explore different buffer conditions during purification, especially regarding ionic strength and pH. 4. adding reducing agents may help, but given that your aggregates are large I expect this may not help that much. 5. add lipids and/or substrate/inhibitor etc during purification. Good luck, Bert From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of Theresa Hsu [theresah...@live.com] Sent: Tuesday, July 09, 2013 5:01 AM To: CCP4BB@JISCMAIL.AC.UK Subject: [ccp4bb] Heterogeneity during purification Dear all I am working on a 30 kDa membrane protein which forms a functional dimer. The protein is His-tagged at N-terminal. In small scale expression screening from whole cells, there is only a single band on Western blot at 30 kDa. But, after purification, additional bands appear at 60 and 120 kDa on SDS-PAGE and Western blot. On size exclusion with Superdex 200, a large proportion elute near the void volume (8 ml). Detail purification For small scale screening, I lysed cells in 20 mM Tris pH 8, 100 mM NaCl, 1 mg/ml lysozyme, 1 % DDM and DNAse for 2 hours and then centrifuged at 16000 g. I then checked the supernatant on SDS-PAGE and scale it up for purification. For purification, I use the buffer 50 mM Tris pH 8, 300 mM NaCl, 20 mM imidazole, 0.05 % DDM (two times CMC of DDM). Is there suggestion to get homogeneous protein? Thank you. Theresa
Re: [ccp4bb] Puzzling observation about size exclusion chromatography
Dear Zhen, I should also point out that the statement Matt made (Superdex is known to have some ion-exchange characteristics, so that it can weakly interact with some proteins.) is not completely correct. Superdex and all chromatographic media made from carbohydrates (dextran, agarose, etc.) are also quite hydrophobic (which is surprising to some). The observation Zhen made is the classic behavior of hydrophobic interaction with the gel filtration media, which led to the development of hydrophobic interaction chromatographic (HIC). For HIC, you load the protein in high salt, and elute with low salt or a chaotrope (LiCl). So when you redo you experiment, Zhen, try with AND without salt. As Matt said, if it is an ion-exchange interaction, extra salt will make it elute more normally. But if it gets worse, then the extra salt is increasing the hydrophobic interactions, and you should run the column in lower salt (~50 mM NaCl) or with a bit of detergent. Given that you are refolding your protein, a partially folded protein may have have more hydrophobic patches. As my lab is routinely refolding our target proteins, we are always watching for this behavior, including on our analytical Superdex 200 10/300 columns, which is one of our favorites. Excessive hydrophobic interactions can also lead to clogged columns, which is not what you want for this expensive column. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jun 20, 2013, at 5:38 PM, Zhang, Zhen wrote: Hi Matthew, Thanks a lot. That is a great idea. I will try the high salt and worry about the crystallization later. Zhen -Original Message- From: Matthew Franklin [mailto:mfrank...@nysbc.org] Sent: Thursday, June 20, 2013 4:34 PM To: Zhang, Zhen Cc: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] Puzzling observation about size exclusion chromatography Hi Zhen - Superdex is known to have some ion-exchange characteristics, so that it can weakly interact with some proteins. This is why the manufacturer recommends including a certain amount of salt in the running buffer. I have had the same experience with a few proteins, including one that came off the column well after the salt peak! (The protein was very clean after this step; all other proteins had eluted earlier.) As others have said, you can't rely on molecular weight calibrations in this case, but this behavior alone is no reason to think that the protein is misfolded or otherwise badly behaved. If you don't like the late elution, try increasing the salt concentration of your running buffer to 250 or even 500 mM. You'll probably need to exchange the eluted protein back into a low-salt buffer for your next steps (e.g. crystallization) if you do this. - Matt On 6/20/13 3:09 PM, Zhang, Zhen wrote: Dear all, I just observed a puzzling phenomenon when purifying a refolded protein with size exclusion chromatography. The protein was solubilized by 8M Urea and refolded by dialysis against 500mM Arginine in PBS. The protein is 40KDal and is expected to be a trimer. The puzzling part is the protein after refolding always eluted at 18ml from the superdex S200 column (10/300), which is calculated to be 5KDal by standard. However, the fractions appear to be at 40KDal with SDS PAGE and the protein is functional in term of in vitro binding to the protein-specific monoclonal antibody. I could not explain the observation and I am wondering if anyone has the similar experience or has an opinion on this. Any comments are welcome. Thanks. Zhen The information in this e-mail is intended only for the person to whom it is addressed. If you believe this e-mail was sent to you in error and the e-mail contains patient information, please contact the Partners Compliance HelpLine at http://www.partners.org/complianceline . If the e-mail was sent to you in error but does not contain patient information, please contact the sender and properly dispose of the e-mail. -- Matthew Franklin, Ph. D. Senior Scientist New York Structural Biology Center 89 Convent Avenue, New York, NY 10027 (212) 939-0660 ext. 9374
Re: [ccp4bb] Self rotation function interpretation
Emma, You need to provide more information. Luckily, I222 is orthorhombic, so a* is along a, b* is along b, and c* is along c. This makes things so much easier to interpret. I hope the plotting conventions you used are typical: b along +y (North pole), a along +x (to the right), and c perpendicular to the page. Let us also assume that you have contours at every 10% of the origin peak height. If so, my quick assessment is that there is not much. At each of the crystallographic axes in the chi=180 map, you see an origin peak arising from the crystallographic 2-fold symmetry; the mirror symmetry is expected with orthorhombic space groups. At ~45 degrees in the ac-plane there is a small peak that is ~40% of the origin peak. An interpretation is that there may be 2-fold NCS, with the axis ~45 away from the crystallographic a- and c-axes, but the noise across the map is kind of high. However, without more details about how you set the SRF up (data completeness, resolution, etc.), matthew's coefficient, and what you expect to see, little more can be done. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Feb 9, 2013, at 11:00 PM, Emma Littlejohn wrote: Hi all, I have generated a self rotation function using MOLREP for a low res data set which indexed in space group I222 (attached). I am hoping it may provide some information on the oligomeric state of the protein. I am new to analysing SF and was wondering if anyone can help with the interpretation or has any tips that might be able to help me shed some light on what the SF is revealing? Thanks in advance. Emma SF.jpg
Re: [ccp4bb] protein crystals or salt crystals
Ed, Protein crystals are fragile but not soft. If your crystals are like gelatine it's unusual. I hate to disagree with the disagreement, but there are many exceptions to this rule. I have seen many protein crystals that are quite malleable and bendable. One protein produced rod-shaped crystals (150x40x40 um) that I could bend by almost 60 degrees and it would slowly snap back. Mounting it old school was a real pain, and their diffraction was mediocre. However, the majority of the crystals I have worked with adhere to the general rule you describe. Where the crystal physical behavior is anomalous, it is often when PEG is used and/or there are multiple components that contribute to the crystal's integrity (as in the case of membrane protein crystals with detergent). In the former case, crystals that sit in PEG solutions too long tend to be cross-linked (most likely due to the aldehydes that can exist in some batches of PEG). One could argue that the crosslinking adds long-range elasticity and a resistance to fracturing. In the latter case, I have observed large beautiful crystals of membrane proteins that have the consistency and malleability of warm butter. Sometimes optimization improved their integrity, and other times a new crystal form is needed. Regards, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Feb 8, 2013, at 9:23 AM, Ed Pozharski wrote: On Fri, 2013-02-08 at 14:53 +0400, Evgeny Osipov wrote: Protein crystals behave rather as gelatine and not as solid I'd have to disagree on that. Protein crystals are fragile but not soft. If your crystals are like gelatine it's unusual. It has been demonstrated that elastic properties of protein crystals are similar to organic solids. Cheers, Ed. -- I'd jump in myself, if I weren't so good at whistling. Julian, King of Lemurs
Re: [ccp4bb] vitrification vs freezing
Actually, to echo Ron, many low-temperature/freezing/vitrification crystal experiments were done in the 1970's, some by Tsernoglou and Petsko, when they were both at Wayne State, I believe. However, the direction Jacques Dubochet was looking at was an extension of work from the early 1960's. EM researchers were looking at freezing of tissues for freeze-fracture imaging. They have tomes about freezing and the different zones of ice crystal formation and vitrification. In fact to bring it full circle, these electron microscopists cite Kathleen Lonsdale (of the International Tables fame) and here work on ice crystal diffraction in the 1950's. While I was trying to stay out of this discussion, I am in favor of flash-freezing or flash-cooling and have no problem with the word frozen to describe a crystal in liquid N2, regardless of the crystallinity (or lack thereof) of the ice. This is the exact opinion of the electron microscopists doing freeze-fracture imaging for over 50 years: the tissue was frozen with a cryogen (e.g., freon), and you looked for the regions that were vitrified. My 2 cents. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Nov 16, 2012, at 1:26 PM, Ronald E Stenkamp wrote: I'm a little confused. Petsko and others were doing low-temperature/freezing/vitrification crystal experiments in the 1970s, right? (J. Mol. Biol., 96(3) 381, 1975). Is there a big difference between what they were doing and what's done now. Ron On Fri, 16 Nov 2012, Gerard Bricogne wrote: Dear all, I think we are perhaps being a little bit insular, or blinkered, in this discussion. The breakthrough we are talking about, and don't know how to call, first occurred not in crystallography but in electron microscopy, in the hands of Jacques Dubochet at EMBL Heidelberg in the early 1980s (see for instance http://www.unil.ch/dee/page53292.html). It made possible the direct imaging of molecules in vitrified or vitreous ice and to achieve higher resolution than the previous technique of negative staining. In that context it is obvious that the vitreous state refers to water, not to the macromolecular species embedded in it: the risk of a potential oxymoron in the crystallographic case arises from trying to choose a single adjective to qualify a two-component sample in which those components behave differently under sudden cooling. I have always found that an expression like flash-frozen has a lot going for it: it means that the sample was cooled very quickly, so it describes a process rather than a final state. The fact that this final state preserves the crystalline arrangement of the macromolecule(s), but causes the solvent to go into a vitreous phase, is just part of what every competent reviewer of a crystallographic paper should know, and that ought to avoid the kind of arguments that started this thread. With best wishes, Gerard. -- On Thu, Nov 15, 2012 at 11:35:46PM -0700, Javier Gonzalez wrote: Hi Sebastiano, I think the term vitrified crystal could be understood as a very nice oxymoron (http://www.oxymoronlist.com/), but it is essentially self-contradictory and not technically correct. As Ethan said, vitrify means turn into glass. Now, a glass state is a disordered solid state by definition, then it can't be a crystal. A vitrified crystal would be a crystal which has lost all three-dimensional ordering, pretty much like the material one gets when using the wrong cryo-protectant. What one usually does is to soak the crystal in a cryo-protectant and then flash-freeze the resulting material, hoping that the crystal structure will be preserved, while the rest remains disordered in a solid state (vitrified), so that it won't produce a diffraction pattern by itself, and will hold the crystal in a fixed position (very convenient for data collection). Moreover, I would say that clarifying a material is vitrified when subjected to liquid N2 temperatures would be required only if you were working with some liquid solvent which might remain in the liquid phase at that temperature, instead of the usual solid disordered state, but this is never the case with protein crystals. So, I vote for frozen crystal.- Javier PS: that comment by James Stroud I forgot to mention that if any dictionary is an authority on the very cold, it would be the Penguin dictionary., is hilarious, we need a Like button in the CCP4bb list! -- Javier M. Gonzalez Protein Crystallography Station Bioscience Division Los Alamos National Laboratory TA-43,
Re: [ccp4bb] Reservoir buffer
Theresa, In addition to the comments provided, you do need to consider the vapor diffusion experiment process as a whole. The primary reasons why we put the precipitant mixture in the reservoir, aside from being lazy, is (1) it provides a straight forward and partially accurate starting point for making artificial mother liquors for handling and soaking soaking crystals and, more importantly (2) it ensures that ALL VOLATILE components come to the expected (assumed) equilibrium values. When I speak of volatile components, I mean not only organics (isopropanol, ethanol, etc.), but also components like ammonia from ammonium sulfate. Since the reservoir volume is often ~50x greater than that of the protein drop, mass action will significantly change the concentrations of the volatile components in the protein drop, which can markedly effect even pH. I know of one group where pH drifting (due to the reservoir solution having a different pH than the drop) had them losing the crystals until they figured that out. This also underscores the importance of having your students and tech keep good notebooks even for buffer making. While we have tried to use different concentrations of a standard PEG solution or LiCl solution as reservoirs, it turned out that we had to adapt it individually to any condition with a volatile component in the protein drop. In the end, inertia and laziness ended up with us returning to the old method. You may have more will power, but you also need to ensure reproducibility. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On 13/11/2012 07:03, Theresa Hsu wrote: Dear all In *initial screening* using vapor diffusion crystallization, does it matter whether the reservoir buffer is also the precipitant in the drop or just a high salt solution like 5 M NaCl? Thank you. Theresa
Re: [ccp4bb] protein cleavage
Dear Rana, I think you need to clear up some confusion about this experiment. MBP fusions suffer from a number of drawbacks depending on what you are doing. First, did you use the MPB domain to purify the fusion protein (with an amylose column)? If so, you also purified native MBP from the E. coli as well (and there is a good amount in the periplasm). Therefore you should expect to see MBP before and after TEV cleavage, regardless of whether you have a fusion or not. Second, did you see the MBP fusion on SDS PAGE, particularly on Westerns with anti-MBP? Depending on your answers, we can troubleshoot your situation. The MBP fusion vectors we have made incorporate an N-terminal His tag, followed by MBP and TEV, so we can purify the fusion either by Ni-chelation or amylose column chromatography (or both). Also we have experienced cases where, despite our best efforts, MBP fusion either is a truncated expression fragment (mostly MBP) or has a relatively inaccessible TEV site. For example, does DHBx dimerize? This could block access to the TEV site. But first, does the MBP-DHBx fusion exist and did you purify the fusion with an amylose column? Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Nov 4, 2012, at 10:24 AM, rana ibd wrote: Dear CCP4 I am having a problem with cleaving my fusion protein and I would be grateful if you advice me regarding this situation, I have an MBP-DHBx fusion protein and I am trying to cleave it using TEV protease, I have tried different ratios and different temperatures with different incubation time but still it will not cleave, all I observe on the gel is the bands of the fusion protein which is 59kDa and the MBP which is 42kDa and the TEV protease which is 27kDa and no sign of the DHBx which is 17kDa,I have also checked the sequence if there was any problem but I could not find anything unusual the sequence was fine , so if you have any suggestions regarding this situation I will be thankful Best Regards Rana
Re: [ccp4bb] off-topic: detergents for the stabilisation of water-soluble proteins
Vitali, Echoing what Dan said, I am not sure why you have chosen detergents first, as there are many other agents which stabilize proteins. Is the instability due to hydrophobic surfaces (e.g., made worse at higher salt) or not. Some non-detergent suggestions are: 1) diols like MPD (see work from Gil Prive's group) 2) non-detergent sulfobetaines (NDSBs), which is the head group in the zwittergent class of deterents. 3) Trimethylamine oxide, which is the head group in the amine oxide class of deterents (LDAO). 4) 200-500 mM L-Arginine 5) Also try 200-500 mM LiCl. The recommendations you received for detergents are very good ones, but remember that many of these detergents are quite dirty, as well as being chemically heterogeneous. Tween 80, Tween 20 and Nonidet P-40 are generally sold at industrial purity (i.e., they are delivered in tanker cars), so either clean them up yourself or buy purified samples (like the Pierce Surfact-Amps brand). Also, be aware that they may be hard to get rid of when you come to setting up crystals. Also look up Sam Gellman's work on detergent-assisted refolding for other stabilizing detergents. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 12, 2012, at 12:42 PM, Vitali Stanevich wrote: Hi, Sorry for off-topic question. Does anyone have experience of the stabilisation of water-soluble proteins by detergents? Protein I'm working with is definitely water-soluble and has high yield, but, unfortunately, not very stable. Especially during concentration. So, we thought that adding some detergents may one of the ways to stabilise protein. So, did anyone do it before or may be know published examples? Any suggestions on the detergent type/concentration would be welcome. Thanks, Vitali
Re: [ccp4bb] sarkosyl
Rajan, This is old news. Ron Kaplan published a lot about sarcosyl extraction of native mitochondrial carrier proteins from inclusion bodies in the late 1990's, but there is an equally robust literature stating that sarcosyl is a denaturant. It is very much dependent on a protein's characteristics as to whether it is sensitive to sarcosyl or not. One has to just try the experiment. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Sep 20, 2012, at 10:33 AM, rajan kumar wrote: here by i am sending you a paper it may help you. regards rajan ACTREC chaps and triton.pdf
Re: [ccp4bb] Process multiple data sets
Uma, Before this discussion goes much further, you need to provide more details: 1) putative space group? 2) observed resolution and diffraction anisotropy? 3) how big was the crystal and what was its shape? Was the crystal split? 4) were the data sets taken at different points on the crystal? Is radiation damage a factor? 5) did you just rotate around phi (or omega) to collect the different data sets or did you change the other angular settings? 6) are all your data sets indexed in exactly the same way (a tricky and non-obvious factor for a novice to appreciate). Using pointless on unmerged data sets helps with this. You have a number of unknowns here, and your problem in merging the data sets may be due to radiation damage, non-uniformity in a large crystal, index refinement problems due to diffraction anisotropy, etc. We routinely merge different data sets from a single crystal, which has been translated and rotated about one axis only. We try to index and process the data sets using a common setting matrix (which is easy with XDS). However, sometimes it just does work, but merging pairs of data sets often allowed us to discard the worst offender(s). Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 1, 2012, at 4:37 PM, Uma Ratu wrote: I notice one thing with my data sets. The unit cells is slightly different from each other. For example, one has a/b/c @ 79/126/83. The other has a/b/c @ 84/127/90. Although they are collected from the same crystal. Is this the reason that I can't index both with same parameter in HKL? And subsequently, can't integrate and scala together. If so, is there a way that I can fix it? Thank you for your advice Uma On Wed, Aug 1, 2012 at 8:50 AM, Uma Ratu rosiso2...@gmail.com wrote: Dear All: I collected 5 data sets from one crystal and would like to process them together. Here is how I did: In HKL2000, load the all data sets. Index each set. When I try Intergrate, the program automatically go through the whole data sets there, and do not go through. I then process data sets by loading one at each time. Index, intergrate and scale all go through very smoothly. But when I put them together, the program just goes crazy. Thank you for advice Uma
Re: [ccp4bb] Somewhat OT: question of professional courtesy
Evette, I think the primary issue is what kind of analysis was being reported on. That is what I look for when I review a manuscript. If the authors are doing a broad structural analysis (homology of TIM barrels, X-ray refinement protocols, etc.), I wouldn't expect citations beyond stating the PDB entries used. However, if this was a primary structural analysis of a macromolecule, I would expect that a discussion of the structural comparison would include references to earlier work(s) on related molecules, but I have seen this happen where a group reinvents the wheel (sometimes rather badly) because they don't take the time to look at the literature, just a DALI run and a PDB search. It is just bad science not to discuss what earlier researchers have done to put your work in context. Just my 2 cents worth, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 25, 2012, at 9:15 AM, Tim Gruene wrote: -BEGIN PGP SIGNED MESSAGE- Hash: SHA1 Dear Evette, the PDB lists the citation when you enter the PDB-ID in the search mask of any of the web-interfaces, which is much easier for the reader than typing the information from the list of references, i.e. all information is in the article by mentioning the PDB-ID. Why do you consider it a matter of courtesy to re-cite the structural work? Cheers, Tim On 07/25/12 14:33, Radisky, Evette S., Ph.D. wrote: Dear bb, This morning as I scanned an accepted manuscript from a well-respected-but-not-particularly-glamorous journal that publishes many macromolecular structures, I came across a brief mention of homology and rmsd with a published structure listed by PDB accession number, but no citation of the primary reference for this structure. (OK, so I wouldn't have noticed or cared had it not been one of mine.) The paper did not have a lot of references, so it was not due to limitation in the number of refs permitted. I have always thought it a matter of professional courtesy to cite the appropriate reference when one uses and mentions a structure from the PDB, but as I think back, I realize no one explicitly told me this-- it is just an assumption that I made. Maybe I am the one with unrealistic expectations here? Is there a general consensus among crystallographers on this practice? Thanks! Evette Evette S. Radisky, Ph.D. Assistant Professor Mayo Clinic Cancer Center Griffin Cancer Research Building, Rm 310 4500 San Pablo Road Jacksonville, FL 32224 (904) 953-6372 - -- - -- Dr Tim Gruene Institut fuer anorganische Chemie Tammannstr. 4 D-37077 Goettingen GPG Key ID = A46BEE1A -BEGIN PGP SIGNATURE- Version: GnuPG v1.4.12 (GNU/Linux) Comment: Using GnuPG with Mozilla - http://enigmail.mozdev.org/ iD8DBQFQD/FSUxlJ7aRr7hoRAsCRAKDBB5CprXaR1v2QtA57n+3LmVPbAACfegbW I/IlD77jIjoUXgFCiMo9tdI= =xqVY -END PGP SIGNATURE-
Re: [ccp4bb] cryo for high salt crystal
Roger's note reminded me of some older literature (old in the sense that this problem extends back into the mid-1970's). Dealing with cryopreservation of crystals grown in high salt can be a real problem, but as many people have pointed out, the normal cryoprotectants can work, although many salts work just as well (not only malonate, but ammonium formate, lithium citrate, etc.). I don't consider 1.6-2 M ammonium sulfate as really high salt; try working with 3 M ammonium sulfate. However, the trick is not simply the cryoprotectant, but also the exchange method, as Roger mentions. While you can make artificial mother liquors (I love these old terms) with high concentrations of salt and nonionic cryoprotectants (sugars, glycerol, ethylene glycol, etc.), that does not mean they will readily exchange with the crystal, even after soaking for hours. Bill Ray, a classical enzymologist and physical biochemist from Purdue, really wanted to determine the crystal structure of phosphoglucomutase with ligands, but there were many difficulties. Using crystals grown in 2.1 M ammonium sulfate was one of these problems. He realized that the phase interactions between the bulk solution (i.e., the mother liquor) and the interstitial salt-rich solvent was a major obstacle in the proper solvent exchange and good subsequent cryopreservation. See how he solved the problem: W. J. Ray, Jr., et al. Removal of salt from a salt-induced protein crystal without cross-linking. Preliminary examination of desalted crystals of phosphoglucomutase by X-ray crystallography at low temperature. Biochemistry. 1991 Jul 16;30(28):6866-75. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jul 12, 2012, at 2:10 PM, Roger Rowlett wrote: We frequently crystallize one of our proteins and variants of it in 1.6-1.8 M ammonium sulfate solutions. Cryoprotection with 25-30% glycerol or 25-30% glucose does not cause precipitation of salts. Both KCl (4.6 M) and ammonium sulfate (5.6 M) have enormous solubilities in water, so I would not expect cryoprotectant concentrations of glycerol or glucose to cause precipitation (We can save cryoprotectant solutions of at least 2 M ammonium sulfate indefinitely). How are you introducing cryprotectant? We use one of two methods: Fish the crystal out of the mother liquor and place into artificial mother liquor with the same composition as the well solution + cryoprotectant. For glycerol or other liquids, you have to make this from scratch. For glucose, we just weigh out 300 mg of glucose in a microcentrifuge tube and make to the 1.0 mL mark with well solution. (Mix well of course before use. Gentle heating in a block or sonication will help dissolve the glucose. Add 4 volumes of artificial mother liquor + 37.5% cryoprotectant to the drop the crystals are in. You can do this all at once, or in stages, keeping the drop hydrated by placing the hanging drop back in the well between additions. If your drops are drying out during crystal harvesting (very possible in dry conditions), you might try harvesting in the cold room, where evaporation is slower. We often have problems with crystal cracking and drop-drying in the winter months when the humidity is very low indoors. The cold room is usually humid enough and cold enough to slow evaporation to allow crystal harvesting. (I hate working in the meat locker, though.) Cheers, ___ Roger S. Rowlett Gordon Dorothy Kline Professor Department of Chemistry Colgate University 13 Oak Drive Hamilton, NY 13346 tel: (315)-228-7245 ofc: (315)-228-7395 fax: (315)-228-7935 email: rrowl...@colgate.edu On 7/12/2012 12:55 PM, m zhang wrote: Hi Jim, 25% is w/v. Thanks for the information. Will check the webinar. Thanks, Min From: jim.pflugr...@rigaku.com To: mzhang...@hotmail.com; CCP4BB@JISCMAIL.AC.UK Subject: RE: [ccp4bb] cryo for high salt crystal Date: Tue, 10 Jul 2012 17:39:56 + Sucrose, sorbitol, Splenda, trehalose, etc, but instead of 25% (is that w/v or v/w?), try using 100% saturated in reservoir, 75% saturated in reservoir, or 50% saturated in reservoir. You will have to TEST these. See also this webinar on cryocrystallography which shows how to make these solutions: http://www.rigaku.com/node/1388 You could also try high salt solutions with similar technique. Good luck! Jim From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of m zhang [mzhang...@hotmail.com] Sent: Tuesday, July 10, 2012 11:28 AM To: CCP4BB@JISCMAIL.AC.UK Subject:
Re: [ccp4bb] The effect of His-tag location on crystallization
Most of the comments you will get will be anecdotal in that people will report the successful results and do not take the time or effort to characterize the less successful results. This often occurs because the tagged portion of the protein is most often disordered, even in the best crystals. Thus, other than saying tagging on this end works, but tagging on that end doesn't, there is little more you can say. Each case will be different, and it is almost impossible to arrive at any generalized conclusion. We prefer C-terminal tagged proteins for a number of reasons, but if an N-terminally tagged protein crystallizes well, so be it. Of the dozens of N- and C-tagged protein structures we have solved in my lab and with collaborators, I have only seen one case of an ordered His-tag: the His residues had coordinated Cd ions, which proved essential for getting good crystals. However, beyond that there was not much more to say. For your protein and the resulting crystals, an N-terminally tagged protein crystallized well. Whether you can draw any more conclusions from these results depends on characterizing crystals of both N- and C-tagged proteins. Just assuming that the C-tagged protein is trying to crystallize in the same or related crystal form as the N-tagged protein is an unwarranted assumption without experimental evidence to back it up. That is why most groups just run with the winner. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jun 26, 2012, at 9:06 PM, weliu wrote: Dear all, We crystallized a protein and found that crystal quality greatly depended on the location of His-tag. When a His-tag was added at the C-terminus, only crystalline precipitate or spherical quasi crystals were grown. However, when the His-tag was moved to the N-terminus, single crystals were grown under a number of conditions, and the best one diffracted to 1.7 angstrom after optimization. I was wondering if there were published reports describing similar cases. Thank you in advance Wei Liu
Re: [ccp4bb] help regarding structure solution
Dear Sonali, I think that first item on your possible to-do list is to verify that you have indeed crystallized the protein you purified. We, too, got great crystals once with protein X (100 kD) and noticed that 1) the lattice constants, space group symmetry, and Matthew's coefficient were within expected values (100 kD monomer in the ASU with moderate solvent content) and 2) the protein seemed to be cleaved into 50 kD fragments in the drop and in the crystal (as expected). However, it was a totally different protein that crystallized, which was why MR didn't work at all. After solving the structure by MIR, we found that a 50 kD minor contaminant ( 1%) crystallized, while our target protein did not. While there is still a good chance that the crystals you looked at contains at least a fragment of your target protein, make sure first, if you can. As Matt recommended, analyze your protein by mass spec and/or N-terminal sequencing to verify that it is what you think it is. Then, as he recommended, try cloning and expressing a truncated variant. Careful limited proteolysis of the full-length protein would also be worthwhile in getting crystals faster. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jun 20, 2012, at 2:13 PM, sonali dhindwal wrote: Dear All, I am working on a protein for last so many years and for which i have got crystal now in a tray which i kept 1 years ago. It diffracts well and resolution is 2.2A, which is good. I indexed in HKL2000, mosflm and automar and it shows P21 space group in all data reduction packages. But when I tried using molrep or phaser then I do not get any solution. The sequence of my protein is having 46% identity with other available crystal structure. Also when I tried to get matthews coffecient, it calculates its molecular mass less ( about 35 kDa) than which should be (original 54kDa) with solvent content 47%. I have also run the silver staining gel of the protein which contained crystal that shows about 45 kD protein band which is 10 less than the original. Also I tried to run gel on crystal but it did not give anything as it was a small crystal. I have tried all combinations of the search model and tried to break available pdb many ways to make different search models but have not got any good solution. Molrep gives contrast even 10 or more but no good electron density map yet. Free R and figure of merit becomes 52% and 42% respectively in Refmac with all the solutions. I will highly appreciate all the suggestions for this kind of problem. Thanks and regards -- Sonali
Re: [ccp4bb] how to get phase of huge complex
Lisa, As others have said, using careful data collection and the modern program suites available (SHARP, Phenix, etc.), a 300 KD complex with 111 Se-Met residues should be quite solvable. But you didn't state is what is in the asymmetric unit (the important figure): one complex with 111 Se-Met residues or 111 Se-Met residues and 300KD in the ASU. We recently solved a structure by SAD with 750 KD of protein and 114 Se-Met residues in the ASU. The anomalous signal at 3.1 A was strong enough to find 109 Se-Met residues and trace about 70% of the chain after the first round of phasing. While we had the potential of NCS to work with, conformational changes between the different monomers meant that NCS methods could not be used in the initial phasing. Trying also James' suggestion of differential labeling and/or including MIRAS methods (Hg, Pt, Sm, etc.) with Se-Met protein should increase your chances. With modern beamlines, you can tune to the most optimal wavelengths for data collection. Good hunting, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Jun 12, 2012, at 10:46 PM, LISA wrote: Hi all, My work is to solve huge complex containing 4 different proteins and total molecular weight is about 300 KD. I can purify the complex by co-expression them in E.coli. This complex contains 8 protein A, 2 protein B and 1 protein C and D. protein B and protein C have homology structures deposited in PDB database. No homology structure available for protein A and D, which contribute 60% of the whole molecular weight for the complex. Now I am trying to find a way to solve the phase of this complex. I am thinking of use sad or mad with se-Met. There total 111 Met residues in this complex. Is it possible to solve this complex by se-Met? Does someone have experience to solve huge complex structure with se-met? It is also very welcome for all the suggestion. Thank you. All the best, Lisa
Re: [ccp4bb] saxs on xtals
Dear Anna, I know that you already have gotten replies from some top experts, but your intriguing problem brought up some issues I have run across in the past. First, from you experience with single crystal diffraction, your results are not that much different from those seen in virus structures where the nucleic acid structure is averaged out. As the nucleic acid doesn't (and mostly can't) adopt the symmetry of the protein shell, the crystallization process alone does the averaging. Just because that ferritin and magnetite have cubic symmetry elements, if they don't line up, the magnetite structure can be averaged out upon crystallization. So, working at lower symmetry may not help, unless there is some directional correlation of the magnetite symmetry and position with the crystal axes. But try P1 and see what happens. A second comment is why not try neutron scattering (SANS or single crystal neutron diffraction), particularly as you can match out the protein with D2O and see just the magnetite. While the same concerns apply for single crystal neutron diffraction, you see more clearly regions of higher average density inside the protein shell. And lastly, have you tried crystallizing your ferritin/nanoparticle complexes in the presence of a magnetic field? It would be a neat trick, and people have tried such things in the past, such as for orienting biomolecules. Some even used old NMR magnets. Would be wild, if it worked. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 7, 2012, at 12:30 PM, anna anna wrote: Dear all, I'd like some suggestions/opinions about the sense of an experiment proposed by a collaborator expert in saxs. In few words, he wants to collect SAXS data on a suspension of protein xtals to investigate low resolution periodicity of the xtal (more details below). The experiment requires a very huge number of xtals to obtain the circles typical of saxs and it is very time-consuming to me (I know nothing about saxs, I have only to prepare the sample). I proposed to measure a single rotating xtal (like in XRD) but he told they don't have a goniometer on saxs beamline. Here is my concern: does it make sense to measure many xtals together? Don't we lose information with respect to single xtal? And, most of all, what can I see by saxs that I can't see by waxs?? Sorry for the almost off-topic question but I think that only someone who knows both the techniques can help me!! Some detail for who is intrigued by my story: we prepared doped magnetite nanoparticles using ferritin as bioreactor. I crystallized this spheres filled with metal and solved the structure at 3.7A but I can see only the protein shell while there is no density inside, even if I know that the nanoparticles are there. A simple explanation is that the particles are free to move in the cavity(note that the diameter of the nanoparticle is shorter then the inner diameter of the protein shell), ie are disordered, and do not contribute to diffraction, in fact, to my knowledge, nobody have ever seen the metal core inside ferritin or dps proteins. However, since they are magnetic particles they must see each other through the protein wall, ie they can't be completely free to move in the cavity. Maybe, but this is just my opinion, I don't see the particle because the period of the particle in the xtal is different/longer than the period of the protein shell. Anyway, we are interested in the relative distance between the magnetic particles in the xtal to study the effects of magnetostatic interactions in nanoparticles 3D arrays. We are going to do this by saxs since, they told me, lower resolution is useful in studying this long range periodicity (the diameter of ferritin is about 120A) but it seems fool to me using a suspension of so many xtals to obtain a scattering curve while I could collect diffraction images from a single xtal!!! I know that saxs is used when you don't have xtals but if you have xtals, ie your system is ordered, xtallography is much more powerful!! Another question: how can I handle my diffraction data at 3.7A resolution to look for nanoparticles? Should I try a lower symmetry? Maybe the anomalous signal? Have you any reference for a similar case? Thank you very much!! anna
Re: [ccp4bb] detergent or protein
Anita, To answer a lot of your questions, you need to provide some more information about your protein, at least some of its biophysical characteristics (size, oligomeric state, an integral membrane protein, etc.). Just because you are using detergents does not mean all your problems are detergent related. Also you need to provide a bit more information about the conditions where you see crystals, particularly about the presence of divalent cations and/or phosphate and the detergent concentrations. Lots of things will crystallize: contaminating proteins, small molecules, and (although not as often) detergents. The fact that you see crystals under conditions with quite different detergents suggests that the crystals are not detergents (unless you are very unlucky). For example, C10E6 is a well characterized and studied detergent that has no apparent crystalline phase in aqueous solution. Thus, your hexagonal crystals are interesting. My own bias is that I don't trust what the Izit dye suggests, because its partitioning is not exclusively dependent on protein. If your conditions don't have ammonium ions or free amines (i.e., no Tris buffer, ammonium sulfate, etc.), try adding a microliter droplet of glutaraldehyde next to your protein drop. The glutaraldehyde will vapor diffuse into the protein droplet and protein crystals will become fixed via the lysines. They won't dissolve in buffer; they will also turn a bit yellow from the Schiff's bases being formed. Works great on membrane proteins. It is possible that you can get different crystal habits using different detergents, but providing more information about the conditions of the hits and the detergent concentrations would make it easier to give you suggestions. Were you actually using 5% DDM in the drop or using a 5% DDM stock? With many precipitants, high detergent concentrations results in substantial phase separation. Do you see this? When evaluated suspected hits from screens on a membrane protein, I first think of inorganic compounds, particularly the dreaded struvite (NH4MgPO4), which occurs at very low [PO4], but high ammonium and magnesium concentrations. I next consider the quality of the protein I am producing and the reproducibility of the purification scheme. Sometimes a too pure protein prep won't crystallize, while a less pure prep does fine. Contaminating proteins, particularly those that copurify with Ni-chelation resins (see Bolanos-Garcia and Davies, Biochimica et Biophysica Acta 1760 (2006) 1304–1313 and Structural Genomics Consortium NATURE METHODS 5(2) 135, 2008) can also crystallize at concentrations below 1 mg/mL. Your protein could appear to be 95% pure, but but a 1-2% contaminant could be crystallizing, albeit poorly. A strategy might be 1) to repeat with different protein preps to demonstrate reproducibility and 2) to tweak some of your conditions to explore a wider environmental space. Then evaluate whether your protein preparation protocol needs optimization. While a problem, don't focus to much on the detergent a sole source of your problems. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 6, 2012, at 7:45 AM, anita p wrote: Hi All, Thanks for your advices. I tried staining with izit dye, please have a look at the attached image. The granules are all stained. The detergent used here is Chaps (as an additive to crystallization condition) Does it look like protein ?? If yes, please do advice on methods to improve this. I found granules in some other conditions as well and they didn't stain with izit. So probably they were of detergent... Plz suggest Thanks in advance for suggestions regards Anita On Fri, May 4, 2012 at 11:43 PM, John K Lee kw...@msg.ucsf.edu wrote: Hi Anita, One question that comes up immediately is why you are using DDM @ 5% concentration. If it's a typo, understandable. I figure most of us use DDM at 0.05-0.1% concentration, and I've used as low as 0.015%. Also, as others have pointed out, until you shoot it, you don't know. But if it's hexaganol with very smooth/non-sharp edges, many of us have seen it using DDM. -john On May 4, 2012, at 7:48 AM, anita p wrote: Hi All, I would like to have your expert advice on crystals. I am using detergents as 5% (w/v) of DDM 0.4ul in a 4ul (protein + condition + detergent). The precipitant is 28% peg 20K After 1 day I am able to see little plates of irregular shape . I am able to see some needles if I change into MEGA-10. I am able to see some hexagonal 3D crystals when
Re: [ccp4bb] detergent crystal?
Hongjun, I am in agreement with Bert as DDM is exceedingly difficult to crystallize, even in organic solvents. This is one of the reasons it is so expensive. However, you can produce a lot of quasi-crystals that do show low resolution diffraction. As Bert said, you may have protein/detergent crystals that are just poorly ordered. I would disagree with Bert only slightly concerning these crystals in that while you might suppose that most or all of the lattice contacts are mediated by detergent and not by protein. Instead, you might also be observing protein-protein contacts being disturbed by less than optimal detergent contacts (either detergent-detergent or detergent-protein). Try changing the detergent to decyl-maltoside (DM) to see if you get similar results. It was the change from DDM to DM that really gave great crystals for the 13-subunit bovine cytochrome c oxidase. Another thing to watch out for is the dreaded contamination factor, either by protein or detergent. I have seen cases where crystals were from a contaminating protein (such as those which may bind to Ni-affinity resin and are not separated by gel filtration) at as low as 1% (by weight) contamination. More insidious is detergent contamination. DDM is is the beta anomer, but all batches are contaminated with varying amounts of the alpha anomer. The alpha anomers of alkyl glycoside detergents tend to crystallize much more readily than the beta anomers. Despite their best efforts, manufacturers occasionally produce batches with a high level of alpha anomer contamination. I have personally tested a batch of beta-octyl glucoside (from a very reputable company) that did not dissolve; other batches from a different company were cloudy when making a 10% stock solution. Alpha-octyl glucoside is not soluble below ~32C and make some very nice crystals in aqueous solution at room temperature. So try a batch of DDM from another source. Best of luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 26, 2012, at 5:32 PM, Van Den Berg, Bert wrote: Generally speaking it is quite hard to crystallize DDM since it is so soluble (20% in water). You most likely have protein crystals (of course containing a lot of detergent as well) that are just not ordered, presumably because most or all of the lattice contacts are mediated by detergent and not by protein. Unfortunately this is the norm for membrane protein crystallization. Good luck, Bert From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of 于洪军 [hongju...@moon.ibp.ac.cn] Sent: Friday, April 27, 2012 6:07 AM To: CCP4BB@JISCMAIL.AC.UK Subject: [ccp4bb] detergent crystal? Hi, I am trying to screen crystals of membrane protein in DDM solutions. I got crystals and its diffraction pattern as I enclosed. Membrane protein crystallization seems quite different from soluble protein. The condition contains PEG400. I learn from other topic here that PEG400 can easily produce DDM detergent crystals. Is it detergent crystal ? Can I tell this from the diffraction pattern? Advices would be greatly appreciated. Thank you. Hongjun
Re: [ccp4bb] detergent crystal?
Jacob, The solubility of the alpha anomer of DDM is actually not bad (it's only really bad for the alkyl glucosides), just the phase behavior is different. Still getting pure beta anomer is the tough problem, which is part of the reason it is so expensive. Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 27, 2012, at 10:54 AM, Jacob Keller wrote: Wouldn't the lack of solubility of the alpha form of DDM suggest an easy purification protocol for the beta form? JPK On Fri, Apr 27, 2012 at 8:40 AM, R. M. Garavito rmgarav...@gmail.com wrote: Hongjun, I am in agreement with Bert as DDM is exceedingly difficult to crystallize, even in organic solvents. This is one of the reasons it is so expensive. However, you can produce a lot of quasi-crystals that do show low resolution diffraction. As Bert said, you may have protein/detergent crystals that are just poorly ordered. I would disagree with Bert only slightly concerning these crystals in that while you might suppose that most or all of the lattice contacts are mediated by detergent and not by protein. Instead, you might also be observing protein-protein contacts being disturbed by less than optimal detergent contacts (either detergent-detergent or detergent-protein). Try changing the detergent to decyl-maltoside (DM) to see if you get similar results. It was the change from DDM to DM that really gave great crystals for the 13-subunit bovine cytochrome c oxidase. Another thing to watch out for is the dreaded contamination factor, either by protein or detergent. I have seen cases where crystals were from a contaminating protein (such as those which may bind to Ni-affinity resin and are not separated by gel filtration) at as low as 1% (by weight) contamination. More insidious is detergent contamination. DDM is is the beta anomer, but all batches are contaminated with varying amounts of the alpha anomer. The alpha anomers of alkyl glycoside detergents tend to crystallize much more readily than the beta anomers. Despite their best efforts, manufacturers occasionally produce batches with a high level of alpha anomer contamination. I have personally tested a batch of beta-octyl glucoside (from a very reputable company) that did not dissolve; other batches from a different company were cloudy when making a 10% stock solution. Alpha-octyl glucoside is not soluble below ~32C and make some very nice crystals in aqueous solution at room temperature. So try a batch of DDM from another source. Best of luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 603 Wilson Rd., Rm. 513 Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Apr 26, 2012, at 5:32 PM, Van Den Berg, Bert wrote: Generally speaking it is quite hard to crystallize DDM since it is so soluble (20% in water). You most likely have protein crystals (of course containing a lot of detergent as well) that are just not ordered, presumably because most or all of the lattice contacts are mediated by detergent and not by protein. Unfortunately this is the norm for membrane protein crystallization. Good luck, Bert From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of 于洪军 [hongju...@moon.ibp.ac.cn] Sent: Friday, April 27, 2012 6:07 AM To: CCP4BB@JISCMAIL.AC.UK Subject: [ccp4bb] detergent crystal? Hi, I am trying to screen crystals of membrane protein in DDM solutions. I got crystals and its diffraction pattern as I enclosed. Membrane protein crystallization seems quite different from soluble protein. The condition contains PEG400. I learn from other topic here that PEG400 can easily produce DDM detergent crystals. Is it detergent crystal ? Can I tell this from the diffraction pattern? Advices would be greatly appreciated. Thank you. Hongjun -- *** Jacob Pearson Keller Northwestern University Medical Scientist Training Program email: j-kell...@northwestern.edu ***
Re: [ccp4bb] CCP4I on an imac
Rex, Can give us a bit more detail (OS version, source of CCP4i, and type of errors)? I am in the midst of reorganizing our little Mac computer cluster, and CCP4 with CCP4i installs nicely on OS 10.6.8 and 10.7.1. However, there were some permissions that need resetting to allow you to fire up CCP4i correctly (such as having write permissions on some of the files and directories to setup CCP4i for the first time). Is this the problem? This and other permission problems were discussed earlier, with potential solutions, on the CCP4BB. However, two insidious problems linger with the packages that come from the CCP4 download site. For Coot 0.6.2 to find the CIF files properly with CCP4 6.2.0, the coot startup file has to be edited to point to the CCP4 6.2.0 directory and not to the non-existing CCP4 6.1.13 directory. But this may not be a problem with Coot 0.6.2 and CCP4 6.2.0 obtained elsewhere (such as from Bill Scott's versions on Fink). While I could correct the former problem easily enough, working with Lion and Java with CCP4 6.2.0 output seems more confusing. Opening the the *.log and *.html results in a variety of browsers leads to a note that Java is not enabled for the graphs. Well, Java is installed (which must be done separately on Lion) and enabled, but the browsers still do not seem to be able to find the JLogGraph files in $CBIN. It works fine on 10.6.8, though, and the graphs work fine when using CCP4i. So it is a minor annoyance. But the bottom line is that CCP4i installs fine on most Intel Macs, from iMac to MacMini to MacPro. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Oct 10, 2011, at 6:33 AM, REX PALMER wrote: Dear CCP4'ers Can anyone help with the following problem: How do we install CCP4I on an imac. As yet we have had no success only error messages concerning working directories etc. Thanks in advance Rex Palmer http://www.bbk.ac.uk/biology/our-staff/emeritus-staff http://rexpalmer2010.homestead.com
Re: [ccp4bb] Why Does Cross-linking Mean Anything?
Jacob, One of the problems with glutaraldehyde is the its chemistry is so bizarre. It actually forms quite long transient polymers in solution. You also have to ask yourself why formaldehyde also fixes tissues. This is why glutaraldehyde works so well for tissue fixation for EM as opposed to our usual bivalent crosslinkers we use in biochemistry experiments. Check out the old EM literature about discussions of glutaraldehyde chemistry. Moreover, the Schiff's base linkage glutaraldehyde is slowly reversible. You need to reduce it to make it permanent. I think that glutaraldehyde is a very poor choice for a precise bivalent crosslinker, but as a broad spectrum crosslinker (hitting lysines and a free amino terminii that are different distances apart), glutaraldehyde is great. As it is highly volatile (its unique smell tells you you've had the bottle open too long), you can crosslink crystals by vapor diffusion in an hour. So I would be cautious about interpreting any crosslinking results using glutaraldehyde, except the obvious (i.e., oligomers may indicate the native tertiary state of a protein or complex). Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Sep 15, 2011, at 4:10 PM, Jacob Keller wrote: Dear Crystallographers and Biochemists, cross-linking, say with gluteraldehyde, is an oft-used method of demonstrating a protein's oligomeric state in solution. I have a difficulty with this, however: theoretically (and in practice!), one can tune the amount of cross-linker to get what ever result is desired, such that any protein with some exposed lysines can be cross-linked in any oligomeric state. How, then, does one evaluate the power of this evidence? Maybe one should do a gradient of gluteraldehyde concentrations, then plot the deviation of the observed cross-linked oligomerization from a theoretical null hypothesis? Seems like this could be done, but I have never seen this in the literature... Best, Jacob Keller -- *** Jacob Pearson Keller Northwestern University Medical Scientist Training Program cel: 773.608.9185 email: j-kell...@northwestern.edu ***
Re: [ccp4bb] Protein elution in Size Exclusion
Nian, Before you dump the column, clean it and run some protein standards on it. If everything looks OK, run a small sample of your protein again. If it behaves the same way, then you may have a protein with hydrophobic patches. Anomalous binding to and elution from polymeric SEC media (sepharose, superdex, etc.) occurs with hydrophobic proteins, including some membrane proteins. This is how they discovered hydrophobic chromatography. If you equilibrate your column in low salt, but your protein is in high salt (as might happen right after a Ni-chelation column), some of it will bind to the column. As the salt concentration decreases, it will release off the column, spreading the protein peak over a wider volume. However, it may still weakly bind to the column matrix and spread out even more. If this is the case, some remedies would be to vary the salt concentration (lower the better), add in detergents, or use a chaotropic salt like LiCl instead of NaCl. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 28, 2011, at 5:35 PM, Nian Huang wrote: Hi, David, What is the common cause of knackered SEC column? Will equilibrizing a buffer containing 150 mM NaCl directly into a 20% EtOH or vise versa cause the problem. There was no problem just after packing the column. Nian On Sun, Aug 28, 2011 at 9:24 AM, David Briggs drdavidcbri...@gmail.com wrote: Following on from Roger's fine suggestions: 8. Your column is knackered. Can you see fine lines or cracks in the column? Good packing is v.important for SEC columns. HTH, Dave David C. Briggs PhD Father, Structural Biologist and Sceptic University of Manchester E-mail: david.c.bri...@manchester.ac.uk http://manchester.academia.edu/DavidBriggs (v.sensible) http://xtaldave.wordpress.com/ (sensible) http://xtaldave.posterous.com/ (less sensible) Twitter: @xtaldave Skype: DocDCB On 28 August 2011 10:25, Allan Pang a.p...@qmul.ac.uk wrote: Hi there everyone, What does it mean when you have proteins eluting in almost the whole column volume of S200? I ran a gel with fractions from 8ml to 20ml and saw band for my protein all throughout. Judging peaks on chromatogram is not useful as it doesn't have any aromatic rings. Cheers, Allan -- Allan Pang PhD Student G35 Joseph Priestley Building Queen Mary University of London London E1 4NS Phone number: 02078828480
Re: [ccp4bb] Aging PEGs
Time to start digging in the archives. Try looking at work by Fran Jurnak in 1986 (J. Cryst. Growth 76, 577) and Bill Ray's work in 1985 (Analytical Biochem 146, 307), and then the works that cite them. I thought this was common knowledge, but I guess it goes in phases. Aging of poly(oligo)-oxyethylene-based compounds is well known in the surfactant field as it changes the chemical properties of common detergents (Brij, Triton, C10E6, etc.), not only by adding aldehydes and carboxylates to the system, but also by increasing metal binding. It is a sobering sight to see old PEG cross-link your 3-month old crystal: the damn thing wouldn't dissolve after 2 hours in plain buffer, but when I poked it with a glass fiber, the protein oozed out like the center of a cherry cordial, leaving a sad looking deflated shell of a crystal. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 24, 2011, at 5:21 PM, Frank von Delft wrote: And now, does anybody know of systematic data indicating how consistently all this matters? phx On 24/08/2011 21:45, Prince, D Bryan wrote: For those of us truly controlling types :), I used to make the PEG solutions and filter them over a Bio-Rad resin that filtered out all the junk added to stabilize the PEG solution. Then, of course I had to freeze all my PEG solutions in aliquots, or wrap them in foil and store at 4C in the dark. This would take several days, depending on the FW of the PEG. If you are really sensitive about what is in your PEG solutions, try GC-grade PEG's. The FW profile is much more restricted around the reported value (i.e. PEG 3350 molecular biology grade has a broad peak centered on Mr=3350. PEG 3350 GC-grade has a much tighter peak profile.) Back before you could buy Crystal Screen I, II or HT, you had to make the stock solutions, then make the screen. But at least when you did that, you had all the stocks. Now, I just buy pre-made solutions, and keep them in a drawer with a date opened written on the bottle. Isn't progress grand? :) Bryan -- Confidentiality Notice: This message is private and may contain confidential and proprietary information. If you have received this message in error, please notify us and remove it from your system and note that you must not copy, distribute or take any action in reliance on it. Any unauthorized use or disclosure of the contents of this message is not permitted and may be unlawful. -Original Message- From: CCP4 bulletin board [mailto:CCP4BB@JISCMAIL.AC.UK] On Behalf Of Jacob Keller Sent: Wednesday, August 24, 2011 3:18 PM To: CCP4BB@JISCMAIL.AC.UK Subject: [ccp4bb] Aging PEGs A while ago I measured the pH's of old and new PEGs and found them very different, and internally attributed all old vs new PEG issues to pH. Upon reflection, this seems too simplistic. Are there other known mechanisms of crystallization capacities of PEGs of various ages? Jacob Keller *** Jacob Pearson Keller Northwestern University Medical Scientist Training Program cel: 773.608.9185 email: j-kell...@northwestern.edu ***
Re: [ccp4bb] recommendation for ammonium dihydrogen phosphate cryo
Chris, As others have said, using sugars as cryoprotectants is a good first choice. However, we have run into problems freezing crystals with sugars when the primarily crystallization reagent is a salt at high concentrations (0.8-2M). Although 0.4M ammonium phosphate, is not particularly high, you might try ammonium formate or sodium malonate, sometimes even sodium citrate works (1.0-1.2 M). My worry about any crystals grown in phosphate is that the phosphate anion may be crucial to crystals growth, and its displacement by like ions (sulfate) may be detrimental. If it is not in your case, you might also try lithium sulfate. We have used mixtures of sodium citrate and lithium sulfate to freeze crystals grown in 0.8 M sodium citrate. One other point is that sometimes the crystals need to have a bit of the cryoprotectant as a component of crystallization. I have seen cases where crystals could not be transferred into glycerol, but adding 1-3% glycerol to the crystallization mix yielded crystals that could be transferred into glycerol for freezing. You have a lot more options to try, but a drop on a coverslip may not be the best way to test freezing. Proper freezing depends on having maximal heat transfer, sometimes that can be defeated by having large heat reservoirs (i.e., big drops and the big coverslip) and insulators (i.e., glass coverslips). Freezing works because the objects size (drop and crystal) is small enough to allow rapid and relatively isotropic heat transfer. We alway use a slightly larger loop to test our cryoprotectants. When using ammonium phosphate, watch out for the formation of struvite (NH4MgPO4), a type of kidney stone. They are lovely looking (often octahedral) crystals that easily grow in ammonium phosphate; although magnesium phosphate can be soluble up to ~12 mM, the presence of ammonium markedly increases the formation of struvite, even with micromolar (contaminating) concentrations of magnesium. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com -Original Message- From: CCP4 bulletin board [mailto:CCP4BB@JISCMAIL.AC.UK] On Behalf Of Chris Ulens Sent: Thursday, May 26, 2011 5:27 AM To: CCP4BB@JISCMAIL.AC.UK Subject: Re: [ccp4bb] recommendation for ammonium dihydrogen phosphate cryo Hi, I would like to get recommendations for a proper cryo solution for a crystallization hit from the Hampton crystal screen Ammonium di- hydrogen phosphate 0.4M. I tried increasing glycerol up to 30% with the same ammonium phosphate concentration or increasing glycerol up to 30% in the presence of 1.3M ammonium phosphate. Both gave iced up drops (I only tried quick and dirty tests by dipping a cover glass in liquid nitrogen). Thank you. -Chris
Re: [ccp4bb] crystal bent once open cover slip
Weikai, What you might be experiencing is a detergent effect, i.e., you are near a detergent-dependent crystallization boundary. We have been hit with this many times. Under vapor diffusion conditions, sitting or hanging drop, the protein-detergent complex crystallizes and the free/ bound detergent reaches an equilibrium, but when you open the well, the drop begins to dry out. Hence, the detergent concentration increases, which dissolves/cracks the crystals. We also found that this also happened when our stabilization/freezing buffers had too high a detergent concentration (lower didn't hurt nearly as badly). Anecdotally, we have experienced that too high detergent concentrations inhibited crystallization, perhaps by having too high a concentration of free detergent micelles, which may interfere with the crystallization of the protein-detergent complex (there is alway detergent exchange between the solvent and the crystal). The way we solved the problem is by setting up the crystals at lower initial detergent concentrations. Note that as the concentrations of salt and PEG increases (like during crystallization), the detergent CMC decreases, even for non-ionic detergents. Therefore, by dropping the detergent concentration, often to just below the apparent CMC, we could grow nice stable crystals. Hope this helps, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 24, 2011, at 1:19 PM, weikai wrote: Hi Folks, We have some membrane protein crystals that are grown in 30%PEG400, 0.1M Na Citrate pH 4.5, 0.1M LiCl. The protein is purified in DDM. The crystals are long rods and grown under room temperature in a hanging drop set up. But once we open the cover slip, we see the rods start to break and bend in a few seconds. Since it is in high PEG400, we just directly freeze the crystals. The diffraction only goes to 10 Ang at synchrotron. Have anybody had similar problem before and any suggestions? Thanks a lot, Weikai
Re: [ccp4bb] Heme Proteins
Dear Hari, You might look at Lucy Waskell's experiences with full length mammalian Cytochrome P450 2B4. While she got 50 mg of purified protein per liter of culture, many other heme proteins are much harder to express in E. coli with heme assembly proteins, chaperones, etc. You just have to try it. However, for cytochrome P450s, you might want to resort to some molecular engineering to increase expression (a la Eric Johnson and co.; look of cytochrome P450 2C9 structure for example). Saribas, Gruenke, and Waskell (2001) Overexpression and Purification of the Membrane-Bound Cytochrome P450 2B4. Protein Expression and Purification 21, 303–309. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Aug 26, 2010, at 12:56 PM, Hari Namboodiri wrote: Hi CCPers Can anyone provide insights about expressing heme containing proteins in E.col? Does E.coli need any porphyrin precursor during expression or you need special E.coli strains. I have references mentioning delta-aminolevulinic acid for one ortholog but none for another. The enzyme is CYP51. Thanks Hari
Re: [ccp4bb] Is it possible to mutate a reversible epimerase into an inreversible one?
Vinson, As Dale and Randy pointed out, you cannot change the ΔG of a reaction by mutation: enzyme, which is a catalyst, affects only the activation barrier (ΔE double-dagger). You can just make it a better (or worse) catalyst which would allow the reaction to flow faster (or slower) towards equilibrium. Nature solves this problem very elegantly by taking a readily reversible enzyme, like an epimerase or isomerase, and coupling it to a much less reversible reaction which removes product quickly. Hence, the mass action is only in one direction. An example of such an arrangement is the triose phosphate isomerase (TIM)-glyceraldehyde 3-phosphate dehydrogenase (GAPDH) reaction pair. TIM is readily reversible (DHA = G3P), but G3P is rapidly converted to 1,3-diphosphoglycerate by GAPDH. The oxidation and phosphorylation reactions of GAPDH now make TIM work in one direction. Since many epimerases are very optimized enzymes, why not consider making a fusion with a second enzyme (like a reductase) to make the system flow in one direction. Of course, this depends on what you want to do with the product. Cheers, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On May 18, 2010, at 11:54 AM, Dale Tronrud wrote: Hi, I'm more of a Fourier coefficient kind of guy, but I thought that a ΔG of zero simply corresponded to an equilibrium constant of one. You can certainly have reversible reactions with other equilibrium constants. In fact I think irreversible reactions are simply ones where the equilibrium constant is so far to one side that, in practice, the reaction always goes all the way to product. As Randy pointed out the enzyme cannot change the ΔG (or the equilibrium constant). You could drive a reaction out of equilibrium by coupling it to some other reaction which itself is way out of equilibrium (such as ATP hydrolysis in the cell) but I don't think that's a simple mutation of your enzyme. ;-) Dale Tronrud On 05/18/10 00:31, Vinson LIANG wrote: Dear all, Sorry for this silly biochemistory question. Thing is that I have a reversible epimerase and I want to mutate it into an inreversible one. However, I have been told that the ΔG of a reversible reaction is zero. Which direction the reaction goes depends only on the concentration of the substrate. So the conclusion is, A: I can mutate the epimerase into an inreversible one. But it has no influence on the reaction direction, and hence it has little mean. B: There is no way to change a reversible epimerase into an inversible one. Could somebody please give me some comment on the two conclution? Thank you all for your time. Best, Vinson
Re: [ccp4bb] inexpensive source of DDM
Tony, As Ed said, DDM is not, nor will ever be cheap. However, the Anatrace sol-grade is primarily for the early stages of a protein's isolation and purification, but is not recommended (or intended) for crystallization or biophysical experiments. Use their purer grade for the latter. What do you mean large quantities of DDM? If you need a lot of DDM, Anatrace is always happy to quote you lower prices for larger bulk orders. They have been always eager to help. If you find cheaper sources, ALWAYS make sure there is no batch to batch variations in quality (I have seen a batch of insoluble octyl glucoside from a very reputable German firm!!!). Run CMC measurements and TLC on all new batches and ALWAYS keep a bit of the old batch when you shift over to a new batch. It will save you many days of headaches. Good luck, Michael R. Michael Garavito, Ph.D. Professor of Biochemistry Molecular Biology 513 Biochemistry Bldg. Michigan State University East Lansing, MI 48824-1319 Office: (517) 355-9724 Lab: (517) 353-9125 FAX: (517) 353-9334Email: rmgarav...@gmail.com On Mar 12, 2010, at 4:47 PM, Edward A. Berry wrote: You get what you pay for, and this is not a cheap detergent! If your application is not sensitive to a little bit of the alpha- anomer, try Anatrace sol-grade, 25 g for $803.01: http://www.affymetrix.com/estore/browse/brand/anatrace/product.jsp?productId=131630categoryId=35662#1_1 If you find something cheaper, I'd like to know. Ed Tony Wu wrote: Hello, I am looking for an inexpensive US source for large quantities of dodecylmaltoside. Can anyone help me? Thank you!
